Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Characterization of a large DNA virus (BV-PW1) infecting the heterotrophic marine nanoflagellate cafeteria… St. John, Tanya Marie 2003

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata


831-ubc_2003-0493.pdf [ 6.68MB ]
JSON: 831-1.0090960.json
JSON-LD: 831-1.0090960-ld.json
RDF/XML (Pretty): 831-1.0090960-rdf.xml
RDF/JSON: 831-1.0090960-rdf.json
Turtle: 831-1.0090960-turtle.txt
N-Triples: 831-1.0090960-rdf-ntriples.txt
Original Record: 831-1.0090960-source.json
Full Text

Full Text

CHARACTERIZATION OF A LARGE DNA VIRUS (BV-PW1) INFECTING THE HETEROTROPHIC MARINE NANOFLAGELLATE CAFETERIA SP. by TANYA MARIE ST. JOHN BSc. (Hon.) University of Prince Edward Island, 1998  A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Department of Microbiology and Immunology) We accept this thesis as coji|ornrLifig to the required standard  The University of British Columbia May 2003 © Tanya Marie St. John, 2003  in presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department  or by his  or her representatives.  It is  understood  that  copying  or  publication of this thesis for financial gain shall not be allowed without my written permission.  Department The University of British Columbia Vancouver, Canada  DE-6 (2/88)  ABSTRACT  A virus (designated BV-PW1) infecting the marine heterotrophic nanoflagellate Cafeteria  sp. was purified and characterized. The most successful strategy for  purification of the virus particles was ultracentrifugation of culture lysates followed by sucrose-gradient purification to remove contaminating bacteria and bacteriophage. The virus particles are icosahedral, roughly 230 to 300 nm in diameter and contain at least 2 major polypeptides (55 kDa and 65 kDa) and -30 minor polypeptides. It possesses a dsDNA genome >100 kbp with a predicted G + C content of -34%. Sequence analysis of 279 amino acids at the N-terminus of a putative protein revealed the presence of the first 3 conserved motifs that delineate the helicase/NTPase superfamily II, including the NTP hydrolysis motifs GxGKT/S and DExH/D. A phylogenetic tree based on the aminoacid sequence of this region indicates the putative helicase of BV-PW1 is most similar to that of ASFV of the Asfaviridae,  however, this branch also falls within the  Iridoviridae  clade. At this time there is not enough information to classify BV-PW1 within a particular virus family. The virus infecting the nanoflagellate described in this study has several properties in common with a group of other large nucleocytoplasmic DNA viruses, such as the poxviruses, the asfaviruses, the iridoviruses and the phycodnaviruses.  iii TABLE OF CONTENTS  Abstract  ii  Table of Contents  iii  List of Tables  v  List of Figures  vi  List of Abbreviations  '•  ix  Acknowledgements  x  1  1  Introduction to viruses and protists in the marine environment 1.1  Viral abundance and diversity in the marine environment  1  1.2  Ecological and geochemical effects of marine viruses  2  1.3  Marine protists  5  1.4  Ecological significance of protistan phagotrophy  5  1.5  Viruses infecting marine eukaryotes  6  1.6  Viruses infecting heterotrophic flagellates  8  1.7  Goal and specific aims  2 Characterization of BV-PW1, a large dsDNA virus that infects a marine nanoflagellate 2.1  Introduction  10  12 12  2.2 Material and methods 2.2.1 Host cells 2.2.2 Virus isolate 2.2.3 Purification of virus particles 2.2.4 Transmission electron microscopy (TEM) 2.2.5 Analysis of structural proteins Isolation of proteins Analysis of protein sequences 2.2.6 Nucleic acids Extraction and purification of viral DNA Genome size PCR with degenerate primers Construction of a BV-PW1 clone library Sequencing of clones Analysis of DNA sequences  13 13 14 14 15 16 16 16 17 17 18 18 19 20 20  2.3 Results 2.3.1 Lysis of flagellates 2.3.2 Purification of virus particles and DNA  23 23 23  2.3.3 Analysis of structural proteins 2.3.4 Relationships of BV-PW1 protein domains to other viral sequences 2.4 Discussion 2.4.1 Viral purification 2.4.2 Amplification of DNA 2.4.3 Genome size 2.4.4 Analyses of protein and DNA sequences 2.4.5 Putative helicase of BV-PW1 2.4.6 Future directions Literature Cited  iv 27 28 34 34 35 35 36 37 41 44  APPENDIX A. Optimization of methods for the concentration and purification of viral particles and DNA extraction for BV-PW1: A guide to what didn't work 59 Concentration and filtration methods  59  Purification of virus particles  62  Purification of BV-PW1 DNA  63  v LIST O F T A B L E S  Table 1  GenBank accession numbers for protein sequences used for phylogenetic analyses  21  Table 2  Peptide sequences obtained from mass spectrometry analysis  28  Table 3  Proteins with statistically significant similarities to the putative helicase of BV-PW1. Identities show the percentage of identical amino acids in the homologous region  29  vi LIST O F F I G U R E S  Figure 1  Electron micrograph of the host heterotrophic nanoflagellate of BVPW1 (Garza and Suttle, 1995)  9  Figure 2  Electron micrograph of BV-PW1 particle  10  Figure 3  Growth of nanoflagellates with virus added (•) and without ( T ) . Values are the mean ± range, n= 2  23  Figure 4  Purification of BV-PW1 particles on 20-50% sucrose gradient  24  Figure 5  BV-PW1 particles in band 3 of sucrose gradient shown by A) Epifluoresence microscopy B) TEM (8000 x mag), Scale bar = 2 u,m C) TEM (80000 x mag), Scale bar = 100 nm  Figure 6  24  Genomic DNA isolated from sucrose bands. dsDNA genomes from bacteriophages lambda (A) (size=48.5 kbp) and T4 (size=168.9 kbp) were used as controls. Unpurified lysate (U) was run in lane 4 and standards were run in lanes marked with M  Figure 7  25  PCR analysis of DNA extracted from sucrose gradient samples using random degenerate primers. Bacteriophages T4 and lambda (A), and water (-) were used as controls. DNA standards were run in lanes M  26  VII  Figure 8  Structural proteins of BV-PW1. Proteins obtained from sucrosepurified BV-PW1 particles and run on 10% (A) and 13% (B) acrylamide gels. Molecular weight markers (lane 1). Proteins were first stained with Coomassie Blue and then Silver stain. Bands denoted by arrows were sent for sequencing by massspectrometry  Figure 9  27  Clustal X alignment of SF-2 helicases for phylogenetic analysis. Sequences were realigned to exclude the long non-conserved Nterminal regions of the cellular proteins. Abbreviations for the viruses or species are listed in Table 1. Alignments are coloured on the basis of amino acid residue if more than 85% consensus occurs at a site in the alignment. Certain groups of amino acids (W/IJV/I/M/A/F/C/Y/H/P, E/D, K/R, Q/E and T/S) are coloured together if combined they comprise a given percentage (>60%, >50%, >60%, >50% and >50% respectively) at a site. The marks shown above alignments are: asterisks (*) indicate positions which have a single, fully conserved residue; colons (:) indicate that two or more of the residues in the following 'strong' groups are fully conserved: STA, NEQK, NHQK, NDEQ, QHRK, MILV, MILF, HY and FYW; periods (.) indicate that two or more of the residues in the following 'weaker' groups are fully conserved: CSA, ATV, SAG, STNK, STPA, SGND, SNDEQK, NDEQHK, NEQHRK, FVLIM and HFY  30  vii Figure 10  Phylogenetic analysis of proteins belonging to the SF-2 group of helicases. The accession numbers and abbreviations for the viruses or species are listed in Table 1. The tree is a weighted NJ tree with bootstrap values based on 100 random alignments shown at the branch nodes. The scale bar represents the expected number of changes per residue position  Figure 11  33  Epifluorescence microscopy of BV-PW1 in filtrates. A) Prefiltered lysate. B) GC50 filtered (1.2 um pore size). C) GF75 filtered (0.8 um pore size). D) 0.45 um filtered. E) 0.22 um filtered  61  ix LIST O F A B B R E V I A T I O N S  aa ASFV Bp BV-PW1 DNA DOM dNTP D-RAPD dsDNA EsV-1 h IPTG Kb kbp kDa kV I MCP min mM M.O.I. NAPS NCLDV NJ NTPase PBCV-1 PFGE rDNA s SDS SDS-PAGE SF-2 TEM U 111 p:m uM VLPs w/v Xgal % °/ /oo  °c  amino-acid african swine fever virus base pair bodo virus-pier water 1 isolate deoxyribonucleic acid dissolved organic matter deoxy ribonucleoside triphosphate degenerate-random amplification of polymorphic DNA double stranded deoxyribonucleic acid Ectocarpus siliculosus virus hours isopropyl-1 -thio- B-D-galactopyranisode kilobase kilobase pairs kilodalton kilovolts litres major capsid protein minutes millimolar multiplicity of infection nucleic acid and protein service nucleocytoplasmic large DNA viruses neighbour joining nucleoside triphosphate phosphohydrolase Paramecium bursaria Chlorella virus pulse field gel electrophoresis ribosomal DNA seconds sodium dodecyl sulfate SDS-polyacrylamide gel electrophoresis superfamily II transmission electron microscopy units microliter micrometer micromolar virus like particles weight per volume 5-bromo-4-chloro-3-indolyl-B-D-galactopyranisode percent parts per thousand degrees celsius  X  ACKNOWLEDGEMENTS  First, I would like to thank my supervisor, Dr. Curtis Suttle, for welcoming me into his lab and giving me the opportunity to keep working on viruses. Thank you also to my committee members Dr. Francois Jean and Dr. Jim Hudson for supporting me and providing helpful feedback throughout the years. Thank you, Dr. Patrick Keeling, for the time you spent with me on phylogenetic trees. Dr. Andrew Lang deserves a big thank you for getting me back up on my feet and keeping me there, it was a pleasure working with you. Thanks to Amy Chan and Dr. Janice Lawrence for their help with the EM and their useful suggestions along the way. Must not forget Andre Comeau, who took the brunt of most of my questions while in the Suttle lab! Thanks to Alex Culley and Mandy Toperoff for their support and friendships in and outside of the lab. A big hug goes out to each of Trina Mcllhargey, Vera Tai and Sandeep Tamber for being the three most constant parts of my somewhat roller coaster lifestyle! My family fits in here too. They are always my biggest fans, even if they don't fully understand why they are cheering. I thank them for always being there for me. Then, of course, there is the 'Toonie Tuesdays' gang. Leading the bunch is the best new addition to the social circuit of the Suttle Lab ever, Emma Hambly and following in hot pursuit Jessie Clasen! Thanks to the whole gang... Pascale Loret, Cindy Frederickson, Steve Short, Alice Ortmann, Sean Brigden, Karen Reid, Nicole McLearn, Jim Rossi, Clemens Pausz, Azeem Ahmad, Janis Lattanzio, Brenton Dickieson, Nina Nemcek, Craig Kahin, Cindy Kam and Kevin Wen.  1 1  1.1  Introduction to viruses and protists in the marine environment  Viral abundance and diversity in the marine environment  There has been increasing evidence in the past two decades that viruses and VLPs (virus-like particles) are abundant (Bergh etal., 1989) and are integral members of marine environments (reviewed in: Wilhelm and Suttle, 1999; Fuhrman, 1999; Wommack and Colwell, 2000). They are ubiquitously distributed and can reach abundances of 10 to >10 particles per liter depending on the marine habitat (Wilhelm 7  11  and Suttle, 1999), with the highest abundances in more productive coastal waters (Fuhrman and Suttle, 1993). Virus abundance is typically correlated to that of bacteria, generally exceeding bacterial numbers by five to tenfold (Wommack etal., 1992; Cochlan etal., 1993; Maranger etal., 1994), although at times it can be well outside this range (Weinbauer etal., 1993). Similarly, Boehme etal. (1993) found that viral direct counts were highly correlated with chlorophyll a concentrations, particulate DNA and bacterial direct counts in the subtropical waters of the Gulf of Mexico. These results suggest that the distribution of viruses in the marine environment is tightly coupled to the distribution of microbial biomass. Viruses that infect primary producers are also abundant in the oceans. Marine phytoplankton, composed of prokaryotes (cyanobacteria and prochlorophytes) and eukaryotes, show evidence of viral infection in nature (reviewed in Fuhrman 1999). For example, viruses infecting the small flagellate Micromonas  pusilla  can exceed 10 per 5  millilitre in coastal waters (Cottrell and Suttle, 1995). Wommack etal. (1999) found natural virioplankton communities to be very diverse, with different viruses appearing and disappearing over time and space. This  viral diversity is seen at many levels; for example, there is evidence of high morphological diversity among marine bacteriophage isolates (reviewed in Borsheim, 1993) and diversity in terms of host range for given isolates has been demonstrated (Suttle and Chan, 1993; Tetart era/., 1996). Genetic diversity has also been shown for viruses infecting (Wilson  etal.,  Micromonas  pusilla  (Cottrell and Suttle, 1995), and for cyanophage  2000). Most recently, Zhong era/. (2002) used phylogenetic analysis of  gene 20 (g20) sequences in cyanophage, which code for the portal vertex protein, to reveal that natural cyanophage populations are strikingly diverse and that their genetic structures vary greatly in different marine environments.  1.2  Ecological and geochemical effects of marine viruses  Viruses potentially infect organisms at all trophic levels of the food web, thereby adding an extra type of control to the previously accepted concepts of control from predation and nutrient limitation (Thingstad  etal.,  1993). Results of both field and  mesocosm studies suggest that, on average, -20% or less of bacterioplankton and phytoplankton mortality is due to viral infection (reviewed in: Suttle, 1994; Wommack and Collwell, 2000). This estimate is highly variable, however, and at times the proportion of mortality resulting from viral lysis can be much greater. For example, Bratbak  etal.  mortality of  (1993) estimated that viral infection accounted for 25 to 100% of the  Emiliana huxleyi  during the collapse of a bloom.  Although viral mortality can play a modest role in controlling the total abundance of bacteria, microalgae, and protists, the important impact of viral infection may be its influence on species diversity. This highlights one difference between viral control and grazing control of microbial communities; viruses infect specific species or strains and  3 thereby affect community composition, whereas protozoan grazing is less species specific and so more likely to affect the abundance of bacteria (Fuhrman, 1999). Thingstad (2000) describes a "kill the winner" mechanism where viruses selectively destroy bacterial strains that are abundant and rapidly growing. This mechanism may explain how so many different bacteria and phytoplankton can coexist on only a few potentially limiting resources. As the competitive dominants become more densely populated, they are particularly susceptible to infection, whereas rare species are relatively protected (Fuhrman, 1999). Since viral infection is considered to be both species specific and density dependent, it has been shown by models and experimentation that viruses can control the steady-state diversity of the bacterial community (Fuhrman, 1999). This diversity is not necessarily limited to species composition, as many strains of the same species can co-exist, as is likely the case during blooms of Synechococcus  (Suttle and Chan, 1993).  The marine microbial food web is a collection of heterotrophic and autotrophic prokaryotes and protists as well as their predators (Azam, 1998). This system regulates the transfer of energy and nutrients to higher trophic levels and greatly influences global carbon and nutrient cycles (Pomeroy, 1974; Azam, 1998). Viruses have been shown to be an important component of this food web. The lysis of hosts by viruses contributes to the pool of dissolved organic matter (DOM) in the form of macromolecules, cell debris and virus particles (Thingstad etal., 1993). This, in effect, shunts organic material away from higher trophic levels and keeps recycling the carbon within the microbial loop. Calculations suggest that up to a quarter of primary production flows through the viral shunt (Wilhelm and Suttle, 1999). Middelboe and Lyck (2002) showed that viruses can have a significant impact on pelagic carbon flow,  4 and that high rates of viral infection increase the importance of bacteria for organic carbon turnover and community respiration. Sequestration of materials in viruses, bacteria and dissolved matter may lead to better retention of nutrients in the euphotic zone, because more material remains in these small non-sinking forms (Fuhrman, 1992). Viruses may also have an important impact on the global population and evolution of marine bacteria through genetic exchange between microorganisms. Transduction is a means of horizontal gene transfer, in which host and viral DNA is packaged into the capsid during the production of virus particles (reviewed in Paul, 1999). The resultant phage particles are released by viral lysis of the host and are then able to inject the DNA contained within the capsid into a new host. Transformation is another process whereby viruses might mediate genetic transfer. Viral lysis results in the release of both host and viral nucleic acids into the surrounding seawater, which may subsequently be taken up by other microorganisms. Due to the high phosphorous content in nucleic acids, this mechanism may play a significant role in the regeneration cycle of phosphorous (Thingstad etal., 1993). Considering the global significance of cyanobacteria as primary producers and nitrogen fixers, it is noteworthy to study the ecological effects of viral infection in these organisms. For example, in some nutrient poor regions of the oceans cyanobacteria are thought to contribute up to 80% of the primary productivity (Li, 1994) and globally in aquatic environments comprise 8% of prokaryotic abundance (Whitman, 1998). Two independent approaches to studying the mortality of cyanobacteria caused by viruses provided consistent results (reviewed in Suttle, 2000a). Experiments on cyanophage decay rates and observations on the proportion of infected cells demonstrated that  several percent of the cells in natural Synechococcus  communities are infected and  lysed on a daily basis. Another study by Agusti etal. (1998) examined the dissolved esterase activity in seawater samples released from phytoplankton as the result of cell lysis. They estimated phytoplankton lysis rates corresponding to as much as 50% of the estimated growth rate of the phytoplankton and proposed that the DOM released from the phytoplankton could account for high heterotrophic activity in surface waters.  1.3  Marine protists  Predation of prokaryotes in aquatic microbial food webs is dominated by phagotrophic, ubiquitously distributed protists, ranging in diameter from flagellates of 2 um or less to ciliates and dinoflagellates of >100 um (reviewed in Sherr and Sherr, 2002). Flagellates can be categorized according to size, with picozooplankton being the smallest (< 2 um), followed by nanozooplankton, (2 - 20 urn), microzooplankton (20 - 200 um), and macrozooplankton (> 200 urn) (Sieburth etal., 1978). Abundances of heterotrophic nanoflagellates in pelagic habitats can vary from 100 to10 flagellates per 4  millilitre (Boenigk and Arndt, 2002).  1.4  Ecological significance of protistan phagotrophy  The main areas of study on phagotrophic protists in aquatic ecosystems have focused on morphology, taxonomy and on the role they play as consumers of other microbial cells (reviewed in Corliss, 2002). The grazing impact of bacterivorous flagellate communities creates a complex top-down pressure on bacteria that should be at least partly responsible for the great diversity of bacteria (Boenigk and Arndt, 2002). It is well established that protistan predation can be a significant source of mortality for  pelagic bacteria, but protists can also be important grazers of phytoplankton and even other heterotrophic protists (reviewed in Sherr and Sherr, 2002). In addition, several studies have shown that heterotrophic nanoflagellates are consumers of viruses in freshwater (Manage etal., 2002) and marine environments (Suttle and Chen, 1992; Gonzalez and Suttle, 1993). These studies showed that viruses can be an important nutritional source for flagellates, but the impact of flagellates on viral turnover rates is typically relatively small (reviewed in Suttle, 2000b). Excretion or egestion by protists causes the release of substantial amounts of DOM, which can be immediately recycled via bacterial uptake (reviewed in Sherr and Sherr, 2002). Phagotrophic protists also add to the pool of regenerated nutrients in aquatic systems through the excretion of nitrogen and phosphorous compounds (Dolan, 1997) and of trace metals such as iron (Barbeau etal., 2001). Finally, an important ecosystem-level function of phagotrophic protists is to channel the production of microbes at the base of food webs to higher trophic levels (Azam et al., 1983).  1.5  Viruses infecting marine eukaryotes  Although the majority of viruses in the ocean infect bacteria (reviewed in Fuhrman 1999), viruses have also been shown to infect phytoplankton and macroalgae (reviewed in: Van Etten etal., 1991; Suttle, 2000b; Wommack and Cowell, 2000). At least 44 taxa of freshwater and marine eukaryotic algae contain viruses or VLPs (Van Etten etal., 1991). Since Van Etten's summary of algae containing VLPs, there have been a number of new host-virus systems isolated. These include large DNA viruses infecting several of the major bloom forming algae such as Heterosigma (Nagasaki and Yamaguchi, 1997; Lawrence etal., 2001; Tai etal., 2003),  akashiwo  Phaeocystis  7 pouchetti(Jacobsen al.,  etal.,  1996; Bratbak  etal.,  1998), and  Emiliana  huxleyi(Bratbak et  1993; Castberg etal., 2002). Several of the other known virus-host systems  infecting photosynthetic eukaryotes include those of the haptophyte spp. (Suttle and Chan, 1995; Sandaa Etten  etal.,  etal.,  1981) and the small flagellate  2001), a  Chlorella-Wke  Micromonas  pusilla  Chrysochromulina  green alga (Van  (Mayer and Taylor,  1979). These large double-stranded DNA viruses that infect algae are related (Chen and Suttle, 1996) and have been assigned to the family name  Phycodnaviridae  (Van  Etten and Ghabrial, 1991). The family  Phycodnaviridae  comprises 4 genera; they share large icosahedral  morphology, an internal lipid membrane and dsDNA genomes of 180 to 560 kb (Van Etten  etal.,  2002). Based on phylogenetic trees derived from DNA polymerase gene  fragments, Chen and Suttle (1996) found that the related to the  Herpesviridae  Phycodnaviridae  were more closely  than to other virus families for which DNA polymerase  sequences were available. More recently, the sequencing of two virus genomes from the  Phycodnaviridae,  etal.,  the genome of  1997) and the brown alga  Paramecium  Ectocarpus  bursaria Chlorella  siliculosus  virus (PBCV-1) (Li  virus (EsV-1) (Delaroque  etal.,  2001), has allowed for a more in depth comparison of this family to other large dsDNA eukaryotic viruses. Phycodnaviruses have 9 genes in common with 3 other families of large eukaryotic DNA viruses (poxviruses, asfaviruses, iridoviruses) and 22 more genes are present in at least three of the four families, which are collectively referred to as nucleocytoplasmic large DNA viruses (NCLDV) (Iyer  etal.,  2001). As well, this group of  viruses has unique features of genomic DNA organization and virion structure. African Swine Fever virus (ASFV) is the sole representative of the asfavirus family, while the  8 iridovirus family is composed of numerous viruses that infect insects and aquatic animals. Comparisons among representative genomes of NCLDVs suggest that their common ancestor was a nucleocytoplasmic virus with an icosahedral capsid, and which encoded complex systems for DNA replication and transcription (Iyer  etal.,  2001).  Recently, a virus (Mimivirus) was discovered in Amoebae and was added to the NCLDV group (La Scola  etal.,  2003). This giant DNA virus (diameter 400 nm) has a genome  size of 800 kbp, 21 proteins with known functional attributes and clear homologues with at least one of the virus families in Iyer  etal.  (2001).  There have been several other isolated reports of viruses found in small marine eukaryotes, other than phytoplankton, including one by Comps etal. (1991) that describes a virus infecting the rotifer  Brachionus  plicatilis.  In this study, only the  histological and cytological aspects of the disease, and the ultrastructure of the purified virions was described. As well, they reported the genome consisted of two segments of RNA similar to the family  Birnaviridae.  Others have reported viruses in apochlorotic  flagellates; Nagasaki etal. (1993) observed VLPs in flagellates in natural plankton communities, while Garza and Suttle (1995) isolated and provided preliminary characterization of a virus infecting a marine heterotrophic flagellate. The latter is the virus system that provided the impetus for my research.  1.6  Viruses infecting heterotrophic flagellates  To date, there has only been one report of a virus (BV-PW1) that has been maintained in culture and which infects a heterotrophic flagellate (Garza and Suttle, 1995). Initially, this  ca.  3  pirn  nanoflagellate (Figure 1) was identified as  subsequently demonstrated to be  Cafeteria  Bodo  sp., but  sp. by 18S ribosomal RNA sequence  9 analysis (Calvalier-Smith, personal communication). Preliminary studies elucidated that the viruses contain dsDNA, are hexagonal or pentagonal in cross section (suggesting icosahedral symmetry) and are roughly 230 to 300 nm in diameter (Figure 2). BV-PW1 is similar in appearance to VLPs observed in other heterotrophic nanoflagellates (Nagasaki  etal.,  1993), and to viruses that have been isolated which infect eukaryotic  phytoplankton (Mayer and Taylor, 1979; Nagasaki and Yamaguchi, 1997; Sandaa 2001). As well, electron microscopy of infected nanoflagellates showed viruses associated with a viroplasm outside of the intact nucleus (Garza and Suttle, 1995) Although, the group of viruses to which BV-PW1 belongs was not determined, its characteristics are consistent with NCLDVs, suggesting that it belongs to the Poxviridae,  Iridoviridae, Phycodnaviridae  or  Asfaviridae.  Figure 1. Electron micrograph of the host heterotrophic nanoflagellate of BVPW1 (Garza and Suttle, 1995).  etal.,  10  100  iun  Figure 2. Electron micrograph of BV-PW1 particle.  1.7  Goal and specific aims  Despite the ecological importance of heterotrophic flagellates as mortality agents of marine microbes, and therefore their significance to global biogeochemical cycles, almost nothing is known about viruses that infect these organisms. The existence of a largely uncharacterized virus system that infects a marine heterotrophic nanoflagellate provided an opportunity to explore the genetic make up of BV-PW1 and its relationship to other viruses. The goal of this study was to further characterize the nanoflagellate virus BVPW1, and to create a clone library of the viral genome. Knowledge of the genome of this virus is a first step towards developing molecular tools that can be used to examine  11 viruses infecting heterotrophic flagellates in natural marine communities. The approach that was used involved characterizing the major viral proteins and several genes in an attempt to identify the relationship of BV-PW1 to other viruses. This study provides the most detailed report to date of a virus infecting a marine heterotrophic flagellate. The specific aims of the study were as follows: 1. Purify BV-PW1 virus from contaminating bacteria, bacteriophage and host flagellates. 2. Isolate viral nucleic acid and create a genomic clone library. 3. Identify the size and number of major viral proteins, and if possible obtain amino acid sequence data.  12 2  Characterization of BV-PW1, a large dsDNA virus that infects a marine nanoflagellate  2.1  Introduction Viruses and VLPs have been observed in many types of marine bacteria and  algae, even in several small eukaryotes, but very few of these viruses have been isolated and characterized (Van Etten etal., 1991). Viruses have been shown to play a significant role in natural marine microbial communities (reviewed in: Suttle, 2000b; Wommack and Colwell, 2000) including carbon and nutrient cycling, changing community structure, and as termination factors of toxic algal blooms. Most research has focused on bacteriophages, and algal viruses, but very little work has been done on the viruses infecting the heterotrophic protists that are major grazers in the oceans. To learn more about the general features and ecological significance of viruses in the NCLDV group, and to develop molecular methods that may be applied in field studies, it has become necessary to bring more host-virus systems into the laboratory where they can be characterized and subjected to experimental studies. The purpose of this study is to describe the heterotrophic nanoflagellate virus, BV-PW1, by looking at its genome size, protein profile and several genes to help infer its relationship to other viruses.  13 2.2  2.2.1  Material and methods  Host cells  The heterotrophic nanoflagellate, strain E4-10, recently identified as sp. (very similar to  C. roenbergensis),  Cafeteria  was isolated from a sampling site near Yaquima  Bay, Oregon, USA (Gonzalez and Suttle, 1993). The unicellular, biflagellate protozoan is 3 to 5 um in diameter and ellipsoid in shape. Nanoflagellate cultures were grown at 20°C in the dark in f/2 medium (Guillard, 1975) using ultrafiltered (<30, 000 molecular weight) seawater at a salinity of 25 ppt (%o). The medium was enriched with 0.01% (w/v) yeast extract and autoclaved before use. Yeast extract was added to stimulate the growth of the mixed assemblage of bacteria in the cultures, which provided a primary food source for the flagellates. Flagellate culture stocks were maintained in f/2 medium where the yeast extract was replaced by a wheat grain for slower release of nutrients. Flagellate cultures used for experiments were initiated by adding 5% (v/v) stock culture to fresh medium containing yeast extract. Once exponential growth was reached, flagellates were transferred to fresh medium at an initial concentration of 10 ml" . Abundance of the 1  5  nanoflagellates was determined by removing a small sample of the culture, staining it with Lugol's Iodine fixative and counting the flagellated cells on a haemocytometer. The lower detection limit for this counting method is ~10 ml" , and cultures typically 3  reached 10 ml" . 6  1  1  14 2.2.2 Virus isolate  The flagellate virus used in this study was previously isolated from the coastal waters of Texas, USA and purified as described elsewhere (Suttle etal., 1991). In my study, virus was propagated with the addition of 0.01% (v/v) lysate to exponentially growing cultures of nanoflagellates at an m.o.i. of -0.15 of physical virus particles, incubated for 6 or 7 days until the nanoflagellate abundance dropped below the minimum detection limit of 10 ml' . To determine the presence and abundance of virus 3  1  particles, samples were stained with SYBR Green 1 (Noble and Fuhrman, 1998). Briefly, the virus lysates were fixed with glutaraldehyde (4% final concentration), diluted 100 fold in f/2 medium and filtered onto 0.02 um pore-size anodisc filters (Whatman, Kent, UK). Viruses were visualized with the AX-70 epifluorescence microscope (Olympus) under blue excitation.  2.2.3 Purification of virus particles  Virus particles from 600 ml culture lysates were purified by a series of centrifugation steps. The lysates were first diluted to 10 %o salt concentration, to reduce the buoyancy of the bacteria, with sterile distilled water and centrifuged at 13,000 x g for 1 h in a Sorvall HB-6 rotor to pellet most of the contaminating bacteria and cell debris. Supernatants were transferred to Sorvall AH-629 rotor centrifuge tubes and spun for 3.5 h at 108,000 x g to pellet the virus particles along with any remaining contaminating bacteria. Most of the supernatant was removed and the pellets were allowed to soften overnight at 4°C. The concentrated virus samples were then pooled into TLA-100.3 (Beckman, Palo Alto, USA) centrifuge tubes and centrifuged at 334,000 x g for 30 min at 20°C. The supernatant was removed and the pelleted virus particles resuspended in  15 200 ul of 0.05 M Tris (pH 7.6) by incubating at 4°C overnight. The concentrated 200 ul virus sample was diluted to 1 ml with 0.05 M Tris (pH 7.6) and spun for 5 min at 2,600 x g to pellet remaining bacteria and unwanted large material. The supernatant was transferred to a TLA-100.3 ultracentrifuge tube and the viruses were pelleted at 334,000 x g for 30 min. The pellet was resuspended in 200 ul of 0.05 M Tris (pH 7.6). The viruses were further purified through a sucrose step gradient [20, 30, 40 and 50% (w/v) in 0.05 M Tris (pH 7.6)] and linearized overnight at 4°C. The 200 ul sample was layered on top and the gradient was centrifuged for 1 h at 50,000 x g at 20°C in a SW 40 (Beckman) rotor. This resulted in three visible bands on the gradient, at 1.5, 2.0 and 6.0 cm from the top of the tube. The band at 6.0 cm (the 40-50% sucrose interface) contained the flagellate virus, the other two bands were composed of bacteriophage. Bands were extracted with an 18 gauge syringe needle from the gradient and the collected material was diluted 3-fold with sterile 0.05 M Tris (pH 7.6). The material in the bands was pelleted by ultracentrifugation in the TLA-100.3 rotor at 334,000 x g for 45 min at 20°C and the pellet was resuspended in 100 ul of sterile 0.05 M Tris (pH 7.6) and stored at 4°C.  2.2.4 Transmission electron microscopy (TEM)  One microliter of sucrose-purified virus particles was made up to 10 u.l with 0.05 M Tris (pH 7.6) and allowed to adsorb to a formvar-coated 400-mesh copper TEM grid for 15 min at room temperature. The excess sample was wicked off with hardened No1 filter paper (Whatman) and the grids stained with 1% (w/v) aqueous uranyl acetate for 10 s. Excess stain was wicked off with filter paper. The grids were examined with a Hitachi H7600 transmission electron microscope at an accelerating voltage of 80 kV.  16 2.2.5 Analysis of structural proteins Isolation of proteins  Sucrose-purified BV-PW1 particles were run on sodium dodecyl sulphatepolyacrylamide gel electrophoresis (SDS-PAGE) gels to separate the major structural proteins. A 5 ul virus sample was added to an equal volume of 2x loading buffer (4% SDS, 20% glycerol, 0.2% bromophenol blue, 100 mM Tris [pH 6.8] and 2% betamercapto-ethanol), was heated for 5 min at 95 °C and loaded onto a 10% or 12% polyacrylamide separating gel with a 4% stacking gel. Protein molecular weight standards (New England BioLabs, Pickering, Canada) ranging from 6,500 to 175,000 Da were used for size calibration. The proteins were electrophoresed for 40 min at 200 V and visualized with Coomassie brilliant blue or Silver stain (BioRad, Mississauga, Canada). For mass spectrometry protein sequencing, three of the major bands (20 kDa, 55 kDa, 65 kDa) were extracted from the SDS PAGE gels and sent to the Genome British Columbia Proteomics Centre at the University of Victoria for analysis by nanosprayquadrupole-time of flight (ESI-Q-TOF) mass spectrometry. Analysis of protein s e q u e n c e s  The six peptide sequences obtained ranged in size from 7 - 1 1 amino acids. Their sequences were screened against the Genbank database using the BLAST algorithm (Altschul etal., 1997, to find possible homologies to known proteins. Because the amino acid sequences were so short, the  17 search was performed using the criteria "for short nearly exact matches" under the BLASTp function.  2.2.6 Nucleic acids Extraction and purification of viral D N A  All reagents used in the purification and amplification of the viral DNA were purchased from Invitrogen Life Technologies (Burlington, Canada). Viral DNA was extracted from the same concentrated sucrose-purified virus sample that was used for the protein analysis and TEM work. The protocol for the extraction of DNA from viruses was followed (Cottrell and Suttle, 1995) and wide-bore pipet tips were used to reduce shearing of the DNA. Basically, the method used was a modified version of a standard phenol-chloroform DNA extraction, in which the 100 ul purified virus sample was topped up to 250 ul with 0.05 M Tris (pH 7.6) and incubated for 20 min at 68°C with 0.5 % (w/v) SDS, 50 mM EDTA (pH 8.0) and 50 ug of Proteinase K. DNA from the lysed particles was extracted once with an equal volume of phenol and once with an equal volume of phenol:chloroform (1:1). The nucleic acids were precipitated by adding NaOAc (pH 5.2) to a final concentration of 0.3 M and 2 volumes of ice-cold 100% ethanol and incubated on ice for 20 min. This preparation was centrifuged for 10 min at 20,000 x g, the supernatant was removed and the nucleic acid pellet was washed with 70% ethanol. After air drying for 20 min, the pellet was resuspended in 30 u.l of TE buffer (10 mM Tris, 1 mM EDTA, pH 7.8) and stored at 20°C.  18 G e n o m e size  To determine the size of the BV-PW1 genome, viral nucleic acid from sucrosepurified particles extracted in phenol-chloroform as described above, was analyzed by pulsed-field gel electrophoresis (PFGE) using a BioRad DR2 CHEF unit. BV-PW1 DNA and phage lambda concatamer molecular weight markers (New England BioLabs), were loaded into the wells of 1 % agarose (Invitrogen Life Technologies) gels in 0.5x TBE buffer (45 mM Tris-borate and 1 mM EDTA, pH 8.0) and electrophoresed at 200 V with pulse ramps of 20-45 s at an angle of 120° at 14°C for 23 h in 0.5x TBE buffer. After electrophoresis, gels were stained overnight with SYBR Green 1 (Molecular Probes, Eugene, USA) and visualized with a UV transilluminator (Fisher, Nepean, Canada). P C R with degenerate primers  Viral DNA extracted as described above was used as a template for the amplification of random segments of the virus genome using the D-RAPD (degenerate random amplification of polymorphic DNA) primers 5'-GTCASSWSSW-3' (S=G or C, W= A or T) (Comeau et al., 2003 manuscript submitted). One ul of viral DNA was added to a 49 ul PCR mixture containing Taq DNA polymerase assay buffer (50 mM KCI, 20 mM Tris-HCI [pH 8.4]), 4.5 mM MgCI , 0.6 mM of each deoxy ribonucleoside 2  triphosphate (dNTP), 2 uM of primer and 1.5 U of Platinum Taq polymerase. Negative controls contained all reagents except the template. PCR was carried out with the following cycle parameters: denaturation at 95°C for 2 min, 40 cycles of denaturation at 95°C for 45 s, annealing at 40°C for 3 min, extension at 72°C for 2 min, and a final extension at 72°C for 9 min. PCR products were electrophoresed in 1.5% agarose in  19 0.5 x TBE buffer (45 mM Tris-borate, 1 mM EDTA [pH 8.0]) and examined by ethidium bromide staining. Construction of a BV-PW1 clone library  DNA products from the D-RAPD PCR were cleaned with the QIAquick PCR Purification Kit (Qiagen, Mississauga, Canada) and ligated with T4 DNA ligase (Invitrogen Life Technologies) into the pGEM®-T vector (Promega, Madison, USA). The recombinant plasmids were transformed into chemically competent Escherichia  coli  DH5a cells (Invitrogen Life Technologies) and the cells were plated on LB agar plates containing 200 ug ml" ampicillin and 40 ug ml" 2% 5-bromo-4-chloro-3-indolyl-B-D1  1  galactopyranisode (Xgal). This resulted in the growth of white colonies for cells containing a recombinant plasmid or blue colonies if the transformants did not contain an insert in the vector. Single white colonies were chosen from the plates and used as templates in colony PCR with the following reagents (all from Invitrogen Life Technologies, except the primers): Taq DNA polymerase assay buffer (50 mM KCI, 20 mM Tris-HCI [pH 8.4]), 1.5 mM MgCI , a 0.16 mM of each dNTP, 100 pmol of each primer (m13R and -21m13 2  from the Nucleic Acid and Protein Service [NAPS] centre, Vancouver, Canada) and 0.5 U of Platinum Taq polymerase. The PCR conditions were as follows: 95°C for 1.5 min, followed by 30 cycles of 95°C for 45 s, 50°C for 45 s, and 72°C for 3 min, and a final cycle of 72°C for 5 min. To determine the presence and size of the amplified segments, the PCR products were run on a 1.25% agarose gel with a 100 bp ladder for size calibration and stained with ethidium bromide.  20 Sequencing of clones  Eight clones containing the highest molecular weight inserts  0.6 kb) were  chosen for sequencing. The corresponding colony PCR products were cleaned with the mini-elute purification kit (Qiagen) and sequenced in the forward and reverse directions using the plasmid primers m13R and -21m13. Sequencing reactions were performed with the Big Dye Terminators versions 3.0 or 3.1 (Applied Biosystems, Foster City, USA) and analyzed by the University of British Columbia NAPS Facility. Analysis of D N A s e q u e n c e s  Nucleotide sequences from the eight clones, translated into all 6 reading frames, were used for BLAST searches of the Genbank database (Altschul etal., 1997). Phylogenetic trees were created with the putative BV-PW1 helicase and other known proteins containing the superfamily II (SF-2) helicase motifs from the Genbank database (Table 1). Conserved blocks of amino acids spanning the first three Nterminal motifs of the SF-2 helicases (Gorbalenya and Koonin, 1989) were aligned using CLUSTAL X v1.81 (Thompson etal., 1997) with default settings, except the BLOSUM series was used for the protein weight matrix (Henikoff and Henikoff, 1992).  21 Table 1. GenBank accession numbers for protein sequences used for phylogenetic analyses. Virus or species  Abbreviation  Protein  Accession number  Regina ranavirus  RRV  similar to SWI/SNF2 family helicase  AF367980_1  Infectious spleen and kidney necrosis virus  ISKNV  putative NTPase  NP_612285.1  Rana tigrina ranavirus  TFV  putative NTPase I  NP_571991.1  Lymphocystis disease virus  LCDV-1  SWI/SNF2 family helicase  NP_078720.1  African swine fever virus  ASFV  helicase  NP_042799.1  Cowpox virus  CPV  V117  NP_619907.1  Vaccinia virus  ETF1VACC  early transcription factor 70 kDa subunit  Q9JFA3  SNF2  G A M 1 / S N F 2 protein  CAA40969.1  Drosophila melanogaster  BRM  brahma protein  AAA19661.1  Homo sapiens  S N F2 h  global transcription activator homologous sequence  N P 003060.2  Saccharomyces  cerevisiae  The protein sequence alignments from Clustal X were used to generate a weighted NJ tree using BIONJ (Gascuel, 1997). Bootstrap values shown in Figure 10 were generated in the following manner: TREE-PUZZLE v5.0 (Strimmer and von Haeseler, 1996) was used to create distances from the Clustal X alignment file (substitution matrix = WAG [Goldman and Whelan, 2000]; gamma distributed rates with 8 rate categories). After conversion using PUZZLEBOOT v1.03 (by Micheal Holder and  Andrew Roger []), the programs TREEBOOT, Seqboot v 3.6 and CONSENSE were used to generate the bootstrap values (iterations = 100) and a concensus tree. Trees were plotted using TreeView v1.6.6 (Page, 1996). Sequences from cellular helicases (yeast,  Drosophila  and human) were used as the outgroup based on Sonntag etal. (1994). These cellular proteins are distantly related to the viral helicases, but still members of the SF-2 helicases.  23 2.3  2.3.1  Results  Lysis of flagellates  When 0.01% inoculum was added to exponentially growing flagellate cells, the number of flagellates decreased to near detection levels (ca 10 ml" ) within 6-7 days 3  1  (Figure 3).  Ie  +7  I c"**^ I 0  I  »  20  40  — T "  1  60  —I  80  Time  r—  1  J00  120  r  140  —r  tSO  *-f  180  C^s~)  Figure 3. Growth of nanoflagellates with virus added (•) and without (T). Values are the mean ± range, n= 2.  2.3.2  Purification of virus particles and DNA Sucrose gradients were used to separate the contaminating bacteriophages and  bacteria from BV-PW1 particles. Three bands and a pellet containing bacteria were visible in the gradient (Figure 4). The top two bands (1 and 2) were blue under white  24 light and were shown by epifluorescence microscopy to contain much smaller particles than those seen for BV-PW1. Band 3 was the broadest, yellow in colour and comprised of BV-PW1 particles (Figure 5).  I haul 2  | hand .1  Figure 4. Purification of BV-PW1 particles on 20-50% sucrose gradient.  Figure 5. BV-PW1 particles in band 3 of sucrose gradient shown by A) TEM (8000 x mag), Scale bar = 2 um B) TEM (80000 x mag), Scale bar = 100 nm.  25 The content of the sucrose bands was confirmed by running DNA from the excised sucrose bands on a PFGE gel (Figure 6). The band seen near the 30 kbp mark in the unpurified sample (lane 4) corresponds to the bacteriophage bands seen in the lanes for bands 1 and 2 (lanes 5 and 6, respectively). Despite repeated attempts, I was not successful in producing unsheared DNA from band 3 (lane 7). Instead, a smear ranging from -30 to 100 kbp was consistently produced from the BV-PW1 particles.  Figure 6. Genomic DNA isolated from sucrose bands. dsDNA genomes from bacteriophages lambda (A) (size=48.5 kbp) and T4 (size=168.9 kbp) were used as controls. Unpurified lysate (U) was run in lane 4 and standards were run in lanes marked with M.  26 Random amplification of DNA extracted from the three sucrose bands from Figure 4 resulted in a unique banding pattern for each virus (Figure 7): BV-PW1 (lane 6), the bacteriophage bands (lane 4 and 5) and the controls (lanes 2 and 3). PCR products from sucrose bands 1, 2 and 3 correspond with the bands seen in the unpurified sample (U). The two strongest bands from bands 1 and 2 (lanes 4 and 5) are greatly reduced in the band 3 sample (lane 6).  Figure 7. PCR analysis of DNA extracted from sucrose gradient samples using random degenerate primers. Bacteriophages T4 and lambda (A), and water (-) were used as controls. DNA standards were run in lanes M.  27 2.3.3 Analysis of structural proteins  The size and number of structural proteins in the virus particles were determined by polyacrylamide gel electrophoresis (PAGE). BV-PW1 has at least 2 major structural proteins and -30 minor proteins (Figure 8). The major proteins were 55 kDa and 65 kDa.  Figure 8. Structural proteins of BV-PW1. Proteins obtained from sucrose-purified BV-PW1 particles and run on 10% (A) and 13% (B) acrylamide gels. Molecular weight markers (lane 1). Proteins were first stained with Coomassie Blue and then Silver stain. Bands denoted by arrows were sent for sequencing by massspectrometry.  28 Only two of the three SDS-PAGE protein bands that were submitted were successfully internally sequenced (Table 2). Analysis of the 65 kDa peptide sequences by BLAST did not reveal significant homologies to any viral proteins, however, a portion of the single peptide sequence obtained for the 55 kDa protein matched up exactly with the residues NYNVLR of the major capsid protein of the PBCV-1 virus that infects Chlorella.  Table 2. Peptide sequences obtained from mass spectrometry analysis.  2.3.4  Protein Size  Amino acid sequences  65 kDa  NLEQELVGAQR, NVDQPTYR, VLGDLTLDTK, FLALPXALER, TNGDXTK  55 kDa  LYDLNYNVLR  20 kDa  —  R e l a t i o n s h i p s of B V - P W 1 protein d o m a i n s t o o t h e r viral s e q u e n c e s  Analysis of the translated DNA sequences of the eight clones by BLAST revealed that only one sequence (279 aa) had significant similarities to other proteins in the GenBank database (Table 3), with its best matches to viruses from the family.  Iridoviridae  29  Table 3. Proteins with statistically significant similarities to the putative helicase of BVPW1. Identities show the percentage of identical amino acids in the homologous region. Organism  Protein  BLAST E-value  Regina ranavirus  Similar to SWI/SNF2 family helicase  2 x 10"  Infectious spleen and kidney necrosis virus  putative NTPase  4 x 10"  Rana tigrina ranavirus  putative NTPase I  7 x 10"  Lymphocystis disease virus 1  SWI/SNF2 family helicase  1 x 10"  African swine fever virus  Helicase  0.001  22  Melanoplus sanguinipes entomopoxvirus  putative early transcription factor small subunit V E T F - S  0.004  28  Variola minor virus  F6R protein  0.007  38  Vaccinia virus  Early transcription factor 70 kDa subunit (VETF small subunit)  0.007  38  Identity (%)  11  11  11  9  28 26 28 29  Three conserved domains were found in the 279 aa sequence of BV-PW1 that correspond to the first 3 motifs of the DEAH/D family of the SF-2 helicases (Gorbalenya and Koonin, 1993). These N-terminal regions are highly conserved in the aa sequences from BV-PW1, poxvirus, iridovirus and cellular members of the SF-2 group of proteins (Figure 9). The three motifs are as follows: GxGKT (residues 107-110), DExH (286-289) and the region around the P at residue 226 (223-227).  30 Figure 9. Clustal X alignment of SF-2 helicases for phylogenetic analysis. Sequences were realigned to exclude the long non-conserved N-terminal regions of the cellular proteins. Abbreviations for the viruses or species are listed in Table 1. Alignments are coloured on the basis of amino acid residue if more than 85% consensus occurs at a site in the alignment. Certain groups of amino acids (i.e. W/L7V/I/M/A/F/CA7H/P, E/D, K/R, Q/E and T/S) are coloured together if combined they comprise a given percentage (>60%, >50%, >60%, >50% and >50% respectively) at a site. The marks shown above alignments are: asterisks (*) indicate positions which have a single, fully conserved residue; colons (:) indicate that two or more of the residues in the following 'strong' groups are fully conserved: STA, NEQK, NHQK, NDEQ, QHRK, MILV, MILF, HY and FYW; periods (.) indicate that two or more of the residues in the following 'weaker' groups are fully conserved: CSA, ATV, SAG, STNK, STPA, SGND, SNDEQK, NDEQHK, NEQHRK, FVLIM and HFY.  31  1  STflVACC  aw  -PA  LCOV-1  t u L t r X. * * *# * • Xfc • • * *»* - .  rrrwAcc CPV  Ml  I  tti  ita  •tt • • •» »•••• *« * * *  %•*»*•  tit til  itt  ttt 230 22*  Ml art  23 23 tt tt *t tt 12 it «3 ti tt  32 The base composition of the sequenced region (1276 bp) of BV-PWTs putative helicase gene was found to be 71.7% A + T and 28.3% G + C. The average base composition for all the clones that were sequenced was 66.4% A + T and 33.6% G + C. A phylogenetic tree was constructed to evaluate the evolutionary relationship of the BV-PW1 helicase motifs to helicase sequences from other members of the SF-2 helicases. The entire 279 aa sequence obtained from the sequencing of the single BVPW1 clone and the corresponding regions from the other proteins in the SF-2 group (Table 1) were aligned. This region contains the first 3 conserved helicase motifs nearest to the N-terminus of the protein. The resulting NJ tree (Figure 10) places the BV-PW1 sequence among the viral helicases and closest to ASFV.  33 SNF2h  100 BRM  —  SNF2  CPV  1100 ETF1VACC  ASFV  59 BV-PW1  LCDV-1  ;55 62  TFV  100  40  RRV  — — — —  IS K N V  0.1  Figure 10. Phylogenetic analysis of proteins belonging to the SF-2 group of helicases. The accession numbers and abbreviations for the viruses or species are listed in Table 1. The tree is a weighted NJ tree with bootstrap values based on 100 random alignments shown at the branch nodes. The scale bar represents the expected number of changes per residue position.  34 2.4  Discussion  This thesis presents the first genetic information for a virus that infects a heterotrophic marine nanoflagellate. I was able to show that BV-PW1 produces a putative helicase protein belonging to the helicase/NTPase superfamily, most closely related to proteins from two other large dsDNA virus families, the Asfaviridae Iridoviridae.  and the  This is the first marine virus that has been shown to have homology to  these groups, granted, it is only a single gene phylogeny. I also isolated and analysed the major structural proteins of BV-PW1 and showed that the capsid diameter, morphology and base composition are similar to members of the NCLDV group (poxviruses, asfaviruses, iridoviruses and phycodnaviruses). These findings are discussed in detail below. Garza and Suttle (1995) isolated and described the physical characteristics and host range of BV-PW1. My work confirms their findings on morphology, particle size and growth characteristics of infected cultures, but contributes new information regarding some of the molecular aspects of BV-PW1. I optimized culture techniques and viral purification methods, obtained protein and nucleic acid sequence data and described the first virus of this type to infect a protist. Knowledge gained from this study can be used to acquire information on the temporal and spatial distribution of this pathogen in natural marine environments.  2.4.1 Viral p u r i f i c a t i o n  Purification of the virus particles was complicated by the nanoflagellate cultures not being axenic. Therefore, the cultures included the nanoflagellates and their  35 accompanying virus, bacteria, which served as the flagellate food source and bacteriophages that infected the bacteria. In the end, ultracentrifugation and sucrosegradient purification gave the purest and most concentrated BV-PW1. The different viral purification and DNA extraction techniques explored during the course of this study will be discussed at greater lengths in the appendix.  2.4.2 Amplification of D N A  The amplification of purified viral DNA using degenerate primers proved to be a useful method for determining the purity of BV-PW1 extracts. In Figure 7, evidence of bacteriophage contamination can be seen by comparing the BV-PW1 fraction of the sucrose gradient in lane 6 with the banding pattern from the bacteriophages in lanes 4 and 5. It is clear that BV-PW1 DNA is more abundant than the bacteriophage DNA in purified sample. Studies using D-RAPD PCR by Comeau etal. (2003, manuscript submitted) found that representative viral banding patterns are still seen as long as the abundance of contaminating bacterial is at least two orders of magnitude less than the abundance of viruses. As bacteria were below the detection limit (ca. 10 ml" ) by 5  1  epifluorescence microscopy and the abundance of BV-PW1 particles from band 3 (Figure 4) was 10 ml" , bacterial contamination was not an issue. 11  1  2.4.3 G e n o m e size  The genome size of BV-PW1 was not determined as a smear of DNA was consistently observed ranging between -30 and 100 kbp (Figure 6, lane 7). It is not clear why several purification techniques appeared to produce sheared or degraded DNA (see Appendix). The large size of the virus particle (-230 to 300 nm) suggests  36 that BV-PW1 should also have a large genome > 100 kbp. This is true for iridoviruses, which are 120 to 300 nm in diameter and have dsDNA genomes of 100 to 210 kbp (Williams, 1996). As well, the recently discovered 400 nm in diameter Mimivirus has a genome size of 800 kbp (La Scola etal., 2003). Other members of the NCLDV group also have large genome sizes: Phycodnaviridae kbp) and Asfaviridae  2.4.4  (160-380 kbp), Poxviridae  (130-375  (170-190 kbp) (van Regenmortel etal., [eds] 2000).  Analyses of protein and D N A s e q u e n c e s  Analysis of the proteins revealed that BV-PW1 has 2 major structural proteins (55 kDa and 65 kDa) and at least 30 minor proteins (Figure 8). Comparitively, ASFV has 6 major structural proteins (Cobbold and Wileman, 1998), while more than 50 proteins have been found in extracellular virus particles (Jacobs etal., 1998). Iridoviruses have only one major structural protein (-50 kDa) and more than 20 minor structural proteins (Fields, 1996). Only part (6 amino acid residues) of the 55 kDa protein fragment of BV-PW1 that was sequenced was an exact match to a viral sequence in the GenBank database. This sequence was the major capsid protein (MCP) of the phycodnavirus PBCV-1. Although there is too little information to say with certainty that the MCP of BV-PW1 is related to MCPs from the other members of the NCLDV group, this prediction does not seem unfounded. Homology has been found among the amino acid sequences of the MCPs from iridoviruses, ASFV and PBCV-1 (Tidona and Darai, 1997). MCP is the major structural component of virus particles, comprising 45-50% of the total protein (Black et al., 1981). It would be beneficial to have the complete MCP sequence for BV-PW1, as it tends to be highly conserved and is a valuable indicator in the study of viral evolution (Tidona etal., 1998).  37 The DNA sequence data revealed that for the part of the genome represented by the 8 sequenced clones, BV-PW1 has a G + C content of only 34%. This low G + C content is also found in other members of the NCLDV group. For example, ASFV of the Asfaviridae  has a base composition of 39% G + C (Yanez etal., 1995), and the two  iridoviruses CIV and LCDV both have a G + C content of 29% (Jakob etal., 2001; Tidona and Darai, 1997, respectively). Moreover, the other two members of the NCLDV, the poxviruses and the phycodnaviruses, have G + C contents of 33% for the Cowpox virus and 40% for PBCV-1 (NCBI database, Base composition analysis from LCDV and TFV, the only other iridoviruses that have been sequenced to date, reveals a higher G + C content than observed for BV-PW1 (55%).  2.4.5 Putative helicase of BV-PW1  Of the 8 cloned fragments sequenced, only one was significantly similar to anything when subjected to BLAST searches. Analysis of this translated 279 aa sequence gave the best matches to a putative helicase and nucleoside triphosphate phosphohydrolase (NTPase) belonging to the superfamily II (SF-2) helicases of several iridoviruses (E-value 2 x 10" ). Gorbalenya and Koonin (1993) recognized that subsets 11  of distantly related helicase proteins share short conserved amino acid sequence fingerprints, or motifs. They proposed a general classification of helicases into 5 major groups based on these motifs. A total of 7 conserved motifs were identified for SF-2, the first 2 of which encompass the A and B motifs of the NTP-binding site, GxGKT/S and DExH/D, respectively (Walker et al., 1982). The Walker A box is common to many  38 proteins that bind and hydrolyze NTPs, but it is not necessary for all helicases (Martins etal., 1999).  The portion of the putative helicase region of BV-PW1 to which sequence information was obtained, as seen in Figure 9, carries the Walker A phosphate-binding loop and the Mg -binding aspartic acid motif of Walker B. The third motif in SF-2 2+  helicases is less conserved than the first two and its functions are not completely understood (Caruthers and McKay, 2002). It is interesting to note, however, that the residues SATP, which comprise motif III in BV-PW1, match those of the UvrB protein, a component of the bacterial UvrABC nucleotide excision repair system. Motif III of UvrB participates in the interaction via hydrogen bonds between domains 1 and 2 (Caruthers and McKay, 2002). The helicase SF-2 is composed of a large number of DNA and RNA helicases from archaea, eubacteria, eukaryotes and viruses (Caruthers and McKay, 2002). NTPbinding pattern proteins are typical in dsDNA viruses, being found in all groups for which complete genome sequences have been reported (Gorbalenya and Koonin, 1989). All have the ability to bind ATP, but not all have been shown to have helicase activity (i.e. the ability to promote DNA, RNA or DNA-RNA duplex unwinding) (Gorbalenya etal., 1989). At the time of the report by Yanez etal. (1993), no helicase activity had been reported for poxvirus, yeast-plasmid and ASFV proteins; hence their classification within SF-2 is based solely on amino-acid sequence similarity. Recently, Martins etal. (1999) reported helicase activity for one (NPH-II) of the two DExH box NTPases encoded by the vaccinia virus. The other protein, NPH-I, is a component of the virus RNA transcription elongation complex and serves as a  39 transcription termination factor during the synthesis of viral early mRNAs. NPH-I is the prototype member of a distinct subgroup of DExH proteins that includes Snf2, a component of SWI/SNF global transcription activator complex in yeast and its homologues, including the brahma protein (BRM) in Drosophila  (Elfring etal., 1998). In  no case has helicase activity been demonstrated for any member of this group of proteins, all function in ATPase hydrolysis (Hall and Matson, 1999). Martins etal. (1999) concluded that the functions of the conserved amino acids of the NTPase motifs are context dependent and must therefore be established empirically for any given member of the DExH family. To help answer the question of how the putative helicase of BV-PW1 is related to other helicases from SF-2, a phylogenetic tree, with representation from cellular and other viral members of the family, was created (Figure 10). Three points can be made about the BV-PW1 putative helicase from the resulting tree: 1) it does not group with the cellular proteins, 2) it does not group with the poxvirus helicases, 3) the BV-PW1 helicase groups amongst the viral clade, is distantly related to the iridovirus helicases and most similar to the helicase from ASFV. One must keep in mind that although the BV-PW1 helicase groups closest to that of ASFV, the branch lengths separating them are quite long, indicating that the relationship between them is not very close. It must also be emphasized that this tree is based solely on a short fragment of translated sequence of one gene and that phylogentic relationships for the whole genome cannot be deduced from this data. Certainly, knowing the complete genome sequence or even the entire sequence for the helicase gene and several other genes, such as the DNA polymerase and the MCP, would provide a better understanding as to where BV-PW1 fits with respect to other virus families.  40 Assessment of the information obtained for the MCP, G + C content and partial putative helicase gene of BV-PW1 does not clearly discern the family to which it belongs. BV-PW1 shares characteristics with the in the case of the MCP, the  Phycodnaviridae,  Asfaviridae  and the  Iridoviridae  and,  as well. Incidentally, ASFV was initially  classified as an iridovirus because of the similar morphology. It was the genome analyses that revealed intermediate characteristics between poxviruses and iridoviruses that led to the exclusion of ASFV from the family  Iridoviridae  (Brown, 1986).  One final note regarding the iridovirus connection is worth mentioning. In his review on iridoviruses, Williams (1996) contemplates the role of "alternative" hosts and discusses the possibility of one-virus-multihost systems for iridoviruses. He speculates that aquatic organisms of overtly different taxa than arthropods may be involved in the transmission and persistence of iridoviruses. Referring to PBCV-1, which infects a Chlorella-like alga, he discusses the observation of iridovirus-like particles in other aquatic organisms. With virus particles of 150-190 nm in diameter, containing 5-10% lipid, possessing a major structural protein of -54 kDa, and having a dsDNA genome, it is easy to follow his conjecture. However, it is now clear that the algal viruses belong to their own family  {Phycodnaviridae)  based on genome structure (Van Etten  Similarly, iridovirus-like particles observed in a daphnid,  Cerodaphnia  etal.,  dubia,  1991).  may be an  alternate host for mosquito iridoviruses (Ward and Kalmakoff, 1991). It is interesting to speculate that BV-PW1 may also infect another eukaryotic marine host? In conclusion, this report makes a number of significant advances to our knowledge of BV-PW1. First, a protocol was established to purify the virus particles, isolate viral DNA and major structural proteins and create a clone library. Although the family to which BV-PW1 belongs was not elucidated, the data strongly suggest that BV-  41 PW1 belongs within the NCLDV group of viruses. Whether the virus is part of an established family or the first member of a new family is not yet known.  2.4.6  Future directions  This study provided the initial steps required in the process leading to full characterization of the BV-PW1 virus and a study on the significance of this virus in different natural marine environments. Several possible future directions come to mind and they are outlined here briefly. A portion of the sequence encoding the putative helicase gene of BV-PW1 is now known and the next obvious step would be to obtain the full sequence. An attempt was made to do this by creating a specific primer at the 3' end of the putative helicase DNA sequence and using it in a sequencing reaction with viral genomic DNA as the template. The NAPS unit at the University of British Columbia was unsuccessful at obtaining sequence data from this reaction. This was only tried once and is worth repeating, as this approach has been used to sequence other viruses in the lab. In light of all the work that has been done on the structure and function of proteins belonging to the helicase superfamilies (reviewed in Caruthers and McKay, 2002), it would be of interest to produce the putative helicase protein of BV-PW1 in an expression vector system and look at its properties. Many of the interactions seen in three-dimensional crystal structures are specific to particular helicases, correlating with the family-to-family variations in the sequences of these motifs (Caruthers and McKay, 2002). An activity assay could also be conducted to determine if this protein has NTPase activity or solely functions in ATP hydrolysis.  42 A second method to obtain the rest of this putative helicase gene would involve primer walking with oligonucleotides created from the known nucleic-acid sequences in the BV-PW1 genome. Fluorescently-labelled probes could be made corresponding to specific sequences taken from fragments in the clone library and used in hybridization assays with restriction enzyme digested genomic DNA. The corresponding labelled bands could then be cut out of the agarose gel and used as a template for the sequencing reaction. Similarly, having the sequences from other highly conserved genes such as the DNA polymerase and MCP would provide insight on the evolutionary relationship of BVPW1 to other large dsDNA viruses. Possible primer sites are the universally conserved YGDTDS motif, previously used to amplify DNA polymerase fragments from viruses infecting microalgae (Chen and Suttle, 1995) and another region upstream or downstream that is conserved among iridoviruses and asfaviruses. A number of different strategies could be explored in order to sequence the gene encoding the MCP of BV-PW1. Specific primers were created by Webby and Kalmakoff (1998) to compare the sequences of the MCP gene from 18 diverse iridoviruses. Amplification with these primers was attempted with BV-PW1 genomic DNA, but no product was observed following the described protocol of Webby and Kalmakoff (1998). Further effort should be made to optimize the PCR conditions before ruling out the possibility of amplification using this primer set. Alternatively, degenerate primers could be designed based on these primer sequences. All of the iridoviruses compared by Webby and Kalmakoff (1998) were from the genus Iridovirus, one of the four genera within the family Iridoviridae.  BLAST analysis indicated that the Iridoviridae helicases  that were most similar to the putative helicase from BV-PW1 were from the genera  Lymphocystis  43 and Ranavirus, which infect vertebrates. It might be worthwhile to create  a new primer set based on the sequences from the MCP genes of these viruses. A final suggestion for acquiring the full sequence of the MCP would be to create a degenerate probe based on the peptide sequence information obtained from the major structural proteins that were sequenced internally by mass spectrometry. The viral DNA clone library and/or purified genomic DNA could then be screened for cross hybridization with this probe. Fragments that hybridize with the probe could then be sequenced. Sequence data unique to the DNA polymerase or MCP genes of BV-PW1 would serve as the basis for designing primers that could be used to examine the diversity of similar viruses in nature.  44 L I T E R A T U R E CITED  Agusti, S., M P . Satta, M P . Mura, and E. Benavent. 1998. Dissolved esterase activity as a tracer of phytoplankton lysis: evidence of high phytoplankton lysis rates in the northwestern Mediterranean.  Limnology and Oceanography  43: 1836-1849.  Altschul, S.F., T.L. Madden, A.A. Schaffer, J.H. Zhang, Z. Zhang, W. Miller, and D.J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs.  Nucleic Acids Research  25: 3389-3402.  Arisaka, F., S. Kanamaru, P. Leiman, and M.G. Rossmann. 2003. The tail lysozyme complex of bacteriophage T4.  International Journal of Biochemistry  & Cell  Azam, F. 1998. Microbial control of oceanic carbon flux: the plot thickens.  Science  Biology  35: 16-21.  280:  694-696.  Azam, F., T. Fenchel, J.G. Field, J.S. Gray, L.A. Meyer Reil, and F. Thingstad 1983. The ecological role of water-column microbes in the sea. Marine Progress  Series  Ecology  10: 257-263.  Barbeau, K., E.B. Kujawinski, and J.W. Moffett. 2001. Remineralization and recycling of iron, thorium and organic carbon by heterotrophic marine protists in culture. Aquatic Microbial Ecology  24: 69-81.  45 Bergh, O., K.Y. Borsheim, G. Bratbak, and M. Heldal. 1989. High abundance of viruses found in aquatic environments.  Nature  340: 467-468.  Black, P.N., C D . Blair, A. Butcher, J . L Capinera, and G.M. Happ 1981. Biochemistry and ultrastructure of iridescent virus type 29. Journal  of Invertebrate  Pathology  38: 12-21.  Boehme, J., M.E. Frischer, S . C Jiang, C A . Kellogg, S. Pichard, J.B. Rose, C. Steinway, and J.H. Paul. 1993. Viruses, bacterioplankton, and phytoplankton in the southeastern Gulf-of-Mexico - Distribution and contribution to oceanic DNA pools.  Marine Ecology-Progress  Series  97: 1-10.  Boenigk, J . and H. Arndt. 2002. Bacterivory by heterotrophic flagellates: community structure and feeding strategies. Antonie of General and Molecular  Microbiology  Van Leeuwenhoek  International  Journal  81: 465-480.  Borsheim, K.Y. 1993. Native marine bacteriophages.  FEMS  Microbiology  Ecology102:  141-159.  Bratbak, G., J.K. Egge, and M. Heldal. 1993. Viral mortality of the marine alga huxleyi (Haptophyceae) Progress  Series  and termination of algal blooms.  Marine  Emiliania  Ecology-  93: 39-48.  Bratbak, G., A. Jacobsen, and M. Heldal. 1998. Viral lysis of bacterial secondary production.  Phaeocystis  Aquatic Microbial Ecology  pouchetiiand  16: 11-16.  46 Brown, F. 1986. The classification and nomenclature of viruses - summary of results of meetings of the International Committee on Taxonomy of Viruses in Senday, Sept. 1984.  Intervirology  25: 141-143.  Caruthers, J.M. and D.B. Mckay. 2002. Helicase structure and mechanism. Opinion in Structural Biology  Current  12: 123-133.  Castberg, T., R. Thyrhaug, A. Larsen, R.A. Sandaa, M. Heldal, J.L. Van Etten, and G. Bratbak. 2002. Isolation and characterization of a virus that infects /7ux/ey/(Haptophyta).  Journal of Phycology  Emiliania  38: 767-774.  Chen, F. and C A . Suttle. 1996. Evolutionary relationships among large doublestranded DNA viruses that infect microalgae and other organisms as inferred from DNA polymerase genes.  Virology  219: 170-178.  Cobbold, C. and T. Wileman. 1998. The major structural protein of African Swine Fever Virus, P73, is packaged into large structures, indicative of viral capsid or matrix precursors, on the endoplasmic reticulum.  Journal of Virology 72:  5215-5223.  Cochlan, W.P., J. Wikner, G.F. Steward, D.C. Smith, and F. Azam. 1993. Spatialdistribution of viruses, bacteria and chlorophyll-a in neritic, oceanic and estuarine environments.  Marine Ecology-Progress  Series  92: 77-87.  Comeau, A.M., S.M. Short, and C A . Suttle 2003. Strain typing of viruses using Degenerate-Random Amplification of Polymorphic DNA (D-RAPD). Environmental  Microbiology,  [submitted].  Applied  47 Comps, M., B. Menu, G. Breuil, and J.R. Bonami. 1991. Viral-infection associated with rotifer mortalities in mass- culture.  Aquaculture  93: 1-7.  Corliss, J.O. 2002. Biodiversity and biocomplexity of the protists and an overview of their significant roles in maintenance of our biosphere.  Acta Protozoologica  41:  199-219.  Cottrell, M.T. and C A . Suttle. 1995. Genetic diversity of algal viruses which lyse the photosynthetic picoflagellate Environmental  Microbiology  Micromonas pusilla (Prasinophyceae).  Applied  and  61: 3088-3091.  Delaroque, N., D.G. Muller, G. Bothe, T. Pohl, R. Knippers, and W. Boland. 2001. The complete DNA sequence of the Virology  Ectocarpus  Siliculosus  virus EsV-1 genome.  287: 112-132.  Dolan, J.R. 1997. Phosphorus and ammonia excretion by planktonic protists. Geology  Marine  139: 109-122.  Elfring, L.K., C. Daniel, O. Papoulas, R. Deuring, M. Sarte, S. Moseley, S.J. Beek, W.R. Waldrip, G. Daubresse, A. Depace, J.A. Kennison, and J.W. Tamkun. 1998. Genetic analysis of brahma: the remodeling factor SWI2/SNF2.  Drosophila  Genetics  homolog of the yeast chromatin  148: 251-265.  Fields, B. M., Knipe, D. M., and Howley, P. M. Fields Virology. 1996. Philadelphia, Lippincott-Raven Publishers.  48 Fuhrman, J.A. 1992. Bacterioplankton roles in cycling of organic matter: the microbial food web, p. 361-383.  In  [eds.], P.G. Falkowski and A.D. Woodhead  Productivity and Biogeochemical  Cycles in the Sea.  Primary  Plenum Press.  Fuhrman, J.A. 1999. Marine viruses and their biogeochemical and ecological effects. Nature  399: 541-548.  Fuhrman, J.A. 1992. p. 361-383.  In  [eds.], P.G. Falkowski and A.D. Woodhead  Productivity and Biogeochemical  Cycles in the Sea.  Primary  Plenum Press.  Fuhrman, J.A., D.E. Comeau, A. Hagstrom, and A.M. Chan 1988. Extraction from natural planktonic microoganisms of DNA suitable for molecular biological studies. Applied  & Environmental  Microbiology  54: 1426-1429.  Fuhrman, J.A. and C A . Suttle 1993. Viruses in marine planktonic systems. Oceanography  6: 50-62.  Garza, D.R. and C A . Suttle. 1995. Large double-stranded DNA viruses which cause the lysis of a marine heterotrophic nanoflagellate marine viral communities.  (Bodo  Aquatic Microbial Ecology  sp.) occur in natural  9: 203-210.  Gascuel, O. 1997. BIONJ: an improved version of the NJ algorithm based on a simple model of sequence data.  Molecular Biology and Evolution  17: 685-695.  49 Goldman, N. and S. Whelan. 2000. Statistical Tests of Gamma-Distributed Rate Heterogeneity in Models of Sequence Evolution in Phylogenetics. Biology and Evolution 17:  Molecular  975-978.  Gonzalez, J.M. and C A . Suttle. 1993. Grazing by marine nanoflagellates on viruses and virus-sized particles - ingestion and digestion. Series 94:  Marine  Ecology-Progress  1-10.  Gorbalenya, A.E. and E.V. Koonin. 1993. Helicases - amino-acid-sequence comparisons and structure- function-relationships. Biology  Current Opinion in Structural  3: 419-429.  Gorbalenya, A.E. and E.V. Koonin. 1989. Viral-proteins containing the purine NTPbinding sequence pattern.  Nucleic Acids Research  17: 8413-8440.  Gorbalenya, A.E., E.V. Koonin, A.P. Donchenko, and V.M. Blinov. 1989. Two related superfamilies of putative helicases involved in replication, recombination, repair and expression of DNA and RNA genomes.  Nucleic Acids Research  17: 4713-  4730.  Guillard, R.R.L. 1975. Culture of phytoplankton for feeding marine invertebrates. In: Smith, W.L and Chanley, M. H. (eds.) Culture of marine invertebrate animals  60.  Hall, M . C and S.W. Matson. 1999. Helicase motifs: the engine that powers DNA unwinding.  Molecular Microbiology  34: 867-877.  29-  50 Henikoff, S. and J.G. Henikoff. 1992. Amino-acid substitution matrices from protein blocks.  Proceedings  America89  of the National Academy  of Sciences  of the United States of  : 10915-10919.  Iyer, L.M., L. Aravind, and E.V. Koonin. 2001. Common origin of four diverse families of large eukaryotic DNA viruses. Journal  of Virology 75:  11720-11734.  Jacobs, SC, Dixon, LK, Brookes, SM, and Smith, GL. Expression of African swine fever virus envelope protein J13L inhibits vaccinia virus morphogenesis.  J Gen Virol  79(5), 1169. 98.  Jacobsen, A., G. Bratbak, and M. Heldal. 1996. Isolation and characterization of a virus infecting  Phaeocystis  pouchetii (Prymnesiophyceae).  Journal of Phycology  32:  923-927.  Jakob, N.J., K. Muller, U. Bahr, and G. Dara. 2001. Analysis of the first complete DNA sequence of an invertebrate iridovirus: coding strategy of the genome of chilo iridescent virus.  Virology  286: 182-196.  La Scola, B., S. Audic, C. Robert, L. Jungang, X. De Lamballerie, M. Drancourt, R. Birtles, J.M. Claverie, and D. Raoult. 2003. A giant virus in amoebae.  Science  299: 2033.  Lawrence, J.E., A.M. Chan, and C A . Suttle. 2001. A novel virus (HaNIV) causes lysis of the toxic bloom-forming alga Heterosigma Journal of Phycology  37: 216-222.  akashiwo  (Raphidophyceae).  51 Lee, J . J., Leedale, G. F., and Bradbury, P. (eds). The Illustrated Guide to the Protozoa. 2000. Lawrence, Allen Press Inc.  Li, W.K.W. 1994. Primary production of prochlorophytes, cyanobacteria, and eukaryotic ultraphytoplankton - measurements from flow cytometric sorting.  Limnology  and  39: 169-175.  Oceanography  Li, Y., Z. Lu, L. Sun, S. Ropp, G.F. Kutish, D.L. Rock, and J.L. Van Etten 1997. Analysis of 74 kb of DNA located at the right end of the 330-kb genome.  Virology  Chlorella  virus PBCV-1  237: 360-377.  Manage, P.M., Z. Kawabata, S. Nakano, and Y. Nishibe. 2002. Effect of heterotrophic nanoflagellates on the loss of virus- like particles in pond water. Research  Ecological  17: 473-479.  Maranger, R., D.F. Bird, and S.K. Juniper. 1994. Viral and bacterial dynamics in arctic sea-ice during the spring algal bloom near resolute, NWT, Canada. Ecology-Progress  Series 111:  121  Marine  -127.  Martins, A., C H . Gross, and S. Shuman. 1999. Mutational analysis of Vaccinia virus nucleoside triphosphate phosphohydrolase I, a DNA-dependent ATPase of the DExH box family.  Journal of Virology  73: 1302-1308.  Mayer, J.A. and F.J.R. Taylor 1979. A virus which lyses the marine nanoflagellate Micromonas  pusilla. Nature  281: 299-301.  52 Middelboe, M. and P.G. Lyck. 2002. Regeneration of dissolved organic matter by viral lysis in marine microbial communities.  Aquatic Microbial Ecology  27: 187-194.  Nagasaki, K., M. Ando, I. Imai, S. Itakura, and Y. Ishida. 1993. Virus-like particles in an apochlorotic flagellate in Hiroshima Bay, Japan. Marine  Ecology-Progress  Series  96: 307-310.  Nagasaki, K. and M. Yamaguchi. 1997. Isolation of a virus infectious to the harmful bloom causing microalga Microbial Ecology  Heterosigma Akashiwo {Raphidophyceae).  Aquatic  13:135-140.  Noble, R.T. and J.A. Fuhrman. 1998. Use of SYBR Green I for rapid epifluorescence counts of marine viruses and bacteria. Aquatic  Microbial Ecology^:  113-118.  Page, R.D.M. 1996. Treeview: an application to display phylogenetic trees on personal computers.  Computer Applications  in the Biosciences  12: 357-358.  Paul, J.H. 1999. Microbial gene transfer:an ecological perspective. Microbial Biotechnology  Journal of Molecular  1: 45-50.  Pomeroy, L.R. 1974. The ocean's food web, a changing paradigm.  Bioscience  24: 499-  504.  Proctor, L.M. and J.A. Fuhrman 1991. Roles of viral infection in organic particle flux. Marine Ecology Progress  Series  69: 133-142.  53 Sandaa, R.A., M. Heldal, T. Castberg, R. Thyrhaug, and G. Bratbak. 2001 . Isolation and characterization of two viruses with large genome size infecting Chrysochromulina  ericina (Prymnesiophyceae)  (Prasinophyceae).  Virology  and  Pyramimonas  orientalis  290: 272-280.  Sherr, E.B. and B.F. Sherr. 2002. Significance of predation by protists in aquatic microbial food webs.  Antonie Van Leeuwenhoek  and Molecular Microbiology  International Journal oi  General  81: 293-308.  Sieburth, J.M., V. Smetacek, and J. Lenz 1978. Pelagic ecosystem structure: heterotrophic compartments and their relationship to plankton size fractions. Limnology  and Oceanography  23: 1256-1263.  Sonntag, K.C., P. Schnitzler, E.V. Koonin, and G. Darai. 1994. Chilo iridescent virus encodes a putative helicase belonging to a distinct family within the DEAD/H superfamily - Implications for the evolution of large DNA viruses.  Virus Genes  8:  151-158.  Strimmer, K. and A. Vonhaeseler. 1996. Quartet Puzzling: a Quartet MaximumLikelihood Method for Reconstructing Tree Topologies. Evolution  Molecular Biology  and  13: 964-969.  Suttle, C A . 2000a. Cyanophages and their role in the ecology of cyanobacteria, p. 563589.  In  [eds.], B.A. Whitton and M. Potts  Diversity in Time and Space.  The Ecology of Cyanobacteria:  Kluwer Academic Publishers.  Their  54 Suttle, C A . 2000b. The ecological, evolutionary and geochemical consequences of viral infection of cyanobacteria and eukaryotic algae, p. 247-296. In [ed.], C.J. Hurst Viral Ecology.  Academic Press.  Suttle, C A . 1994. The significance of viruses to mortality in aquatic microbial communities.  Microbial Ecology  28: 237-243.  Suttle, C A . and A.M. Chan 1993. Marine cyanophages infecting oceanic and coastal strains of  Abundance, morphology, cross-infectivity and growth  Synechococcus:  characteristics.  Marine Ecology Progress  Series  92: 99-109.  Suttle, C A . and A.M. Chan 1995. Viruses infecting the marine Prymnesiophyte Chrysochromulina  abundance.  spp.: Isolation, preliminary characterization and natural  Marine Ecology Progress  Series  118: 275-282.  Suttle, C.A., A.M. Chan, and M.T. Cottrell 1991. Use of ultrafiltration to isolate viruses from seawater which are pathogens of marine phytoplankton. Environmental  Microbiology  Applied and  57: 721-726.  Suttle, C A . and F. Chen. 1992. Mechanisms and rates of decay of marine viruses in seawater.  Applied and Environmental  Microbiology  58: 3721-3729.  Tai, V., J.E. Lawrence, A.S. Lang, A.M. Chan, A.I. Culley, and C A . Suttle. 2003. Characterization of HaRNAV, a single-stranded RNA virus causing lysis of Heterosigma  akashiwo  [Raphidophyceae).  Journal of Phycology  39: 343-352.  55 Taylor, F.J.R. 1999. Ultrastructure as a control for protistan molecular phylogeny. American  Naturalist  154: S125-S136.  Tetart, F., F. Repoila, C. Monod, and H.M. Krisch. 1996. Bacteriophage T4 host range is expanded by duplications of a small domain of the tail fiber adhesin. Molecular  Biology  Journal of  258: 726-731.  Thingstad, T.F. 2000. Elements of a theory for the mechanisms controlling abundance, diversity, and biogeochemical role of lytic bacterial viruses in aquatic systems. Limnology  and Oceanography  45: 1320-1328.  Thingstad, T.F., M. Heldal, G. Bratbak, and I. Dundas 1993. Are viruses important partners in pelagic food webs?  Trends in Ecology and Evolution  8: 209-213.  Thompson, J.D., T.J. Gibson, F. Plewniak, F. Jeanmougin, and D.G. Higgins. 1997. The Clustal_X Windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools.  Nucleic Acids Research  25: 4876-4882.  Tidona, C A . and G. Darai. 1997. The complete DNA sequence of lymphocystis disease virus.  Virology 230:  207-216.  Tidona, C.A., P. Schnitzler, R. Kehm, and G. Darai. 1998. Is the major capsid protein of iridoviruses a suitable target for the study of viral evolution? 66.  Virus Genes  16: 59-  56 Van Etten, J.L. and S.A. Ghabrial 1991.  Phycodnaviridae,  p. 137-139.  Francki, C M . Fauguet, D.L. Knudson, and F. Brown Nomenclature  of Viruses.  In [eds.],  Classification  R.I.B.  and  Springer-Verlag.  Van Etten, J.L., M.V. Graves, D.G. Muller, W. Boland, and N. Delaroque. 2002. - Large DNA algal viruses. Archives  Phycodnaviridae  of Virology\47:  1479-  1516.  Van Etten, J.L., L.C Lane, and R.H. Meints. 1991. Viruses and virus-like particles of eukaryotic algae.  Microbiological  Reviews  55: 586-620.  Van Etten, J.L., R. Meints, D. Burbank, D. Kuczmarski, D. Cuppels, and L. Lane 1981. Isolation and characterization of a virus from the intracellular green alga symbiotic with  Hydra viridis. Virology  113: 704-711.  van Regenmortel, M.H., C M . Fauquet, and D.H.E. Bishop 2000. Virus taxonomy. Seventh report of the International Committee on Taxonomy of Viruses.  Walker, J.E., M. Saraste, M.J. Runswick, and N.J. Gay 1982. Distantly related sequences in the a- and a-subunits of ATP synthase, myosin, kinases and other ATP-requiring enzymes and a common nucleotide binding fold . EMBOJ.  8:  945-951.  Ward, V.K. and J. Kalmakoff 1991. Invertebrate Kurstak  Viruses of Invertebrates.  Dekker.  Iridoviridae,  p. 197-226. /n[ed.], E.  57 Webby, R. and J. Kalmakoff. 1998. Sequence Comparison of the Major Capsid Protein Gene From 18 Diverse Iridoviruses.  Archives  of Virology\42:  1949-1966.  Weinbauer, M.G., D. Fuks, and P. Peduzzi 1993. Distribution of viruses and dissolved DNA along a coastal trophic gradient in the northern Adriatic Sea. Environmental  Microbiology59:  Applied and  4074-4082.  Whitman, W.B., D.C. Coleman, and W.J. Wiebe 1998. Prokaryotes: The unseen majority.  Proceedings  of America  of the National Academy  of Sciences  of the United States  95: 6578-6583.  Wilhelm, S.W. and C A . Suttle. 1999. Viruses and nutrient cycles in the sea - Viruses play critical roles in the structure and function of aquatic food webs.  Bioscience  49: 781-788.  Williams, T. 1996. The iridoviruses.  Advances  in Virus Research  46: 345-412.  Wilson, W.H., N.J. Fuller, I.R. Joint, and N.H. Mann 2000. Analysis of cyanophage diversity in the marine environment using denaturing gradient gel electrophoresis, p. 565-570. In [eds.], C R . Bell, M. Brylinsky, and P. JohnsonGreen  Microbial Biosystems:  New Frontiers.  Atlantic Canada Society for  Microbial Ecology.  Wommack, K.E. and R.R. Colwell. 2000. Virioplankton: Viruses in aquatic ecosystems. Microbiology  and Molecular Biology Reviews  64: 69-114.  58 Wommack, K.E., R T . Hill, M. Kessel, E. Russekcohen, and R.R. Colwell. 1992. Distribution of viruses in the Chesapeake Bay. Applied  and  Environmental  58: 2965-2970.  Microbiology  Wommack, K.E., J . Ravel, R.T. Hill, J.S. Chun, and R.R. Colwell. 1999 . Population dynamics of Chesapeake Bay virioplankton: Total- community analysis by pulsed-field gel electrophoresis.  Applied and Environmental Microbiology  65:  231-240.  Yanez, R.J., J.M. Rodriguez, M. Boursnell, J.F. Rodriguez, and E. Vinuela. 1993. Two putative African Swine Fever Virus helicases similar to yeast DEAH premessenger-RNA processing proteins and Vaccinia virus ATPases D111 and D6R.  Gene  134: 161-174.  Yanez, R.J., J.M. Rodriguez, M.L. Nogal, L. Yuste, C. Enriquez, J.F. Rodriguez, and E. Vinuela. 1995. Analysis of the complete nucleotide-sequence of African Swine Fever Virus.  Virology  208: 249-278.  Zhong, Y., F. Chen, S.W. Wilhelm, L. Poorvin, and R.E. Hodson. 2002 . Phylogenetic diversity of marine cyanophage isolates and natural virus communities as revealed by sequences of viral capsid assembly protein gene g20. Applied Environmental  Microbiology  68: 1576-1584.  and  59 APPENDIX A. Optimization of methods for the concentration and purification of viral particles and D N A extraction for BV-PW1: A guide to what didn't work  Concentration and filtration methods  The goal of this study was to do a molecular characterization of the BV-PW1 virus that infects a marine nanoflagellate, described previously (Garza and Suttle, 1995). This required the separation of the BV-PW1 particles from other components in the system, namely the contaminating bacteria and bacteriophage. Initially, the same protocol used in the Garza and Suttle (1995) study, developed by Suttle  etal.  (1991),  was followed for my study. This method included the prefiltration of 18 to 120 I of natural seawater samples through glass-fiber (MFS GC50, 1.2 um nominal pore size) then polyvinylidine difluoride (0.2 or 0.45 um pore size, Millipore) membrane filters in order to remove plankton and bacteria. The filtrates were then concentrated to 60 to 120 ml using a spiral cartridge ultrafiltration membrane with a 30,000 molecular weight cutoff. Although this method worked for their natural samples, it became obvious it was not going to work for the cultured lysates in my study. This was demonstrated by the poor recovery of virus particles ranging between 0.1 and 3% of the initial particles in 10 to 20 I lysates. There are several possible reasons as to why the prefiltration/ultrafiltration method did not produce good virus recoveries. Firstly, one must consider that Garza and Suttle (1995) were dealing with natural virus communities in seawater as opposed to cultured lysates. The extra nutrients and yeast extract, added to the seawater base in the f/2 media, provided excellent growth conditions for both the flagellates and the required bacterial food source. Typically, bacterial abundance in the lysates of infected  60 nanoflagellate cultures, was 10 times greater than virus abundance following the 6 to 7 2  day incubation period. This is contrary to what is normally seen in natural water samples, whereby, virus abundance usually outnumbers bacterial abundance up to ten fold (Wommack etal., 1992; Cochlan etal., 1993; Maranger etal., 1994). Another possible factor could have been the size of bacterial cells, natural bacteria are much smaller than cultured ones (Proctor and Fuhrman, 1991). These issues not only created problems for separating the viruses from the bacteria, it also made it difficult to count the virus particles with epifluorescence microscopy. High viral particle losses were observed during the prefiltration steps in the ultrafiltration method and in any additional filtration of concentrated lysates using HVLP 0.45 um or GV 0.22 urn low protein-binding filters (Millipore). This was most likely due to the extra organic material and cellular debris from the lysates that possibly clogged up the pores of the filters. As seen in figure 11, all filtering attempts resulted in substantial BV-PW1 particle loss, visible with epifluorescence microscopy. Glass fibre membranes (0.8 urn pore size, Whatman) and 0.22 u.m produced the worst results, allowing only 2.2% and 1.2% of the initial virus particles though, respectively. Recovery of BV-PW1 from GC50 (1.2 urn pore size) and 0.45 um filtrates were 7.1% and 37.2%, respectively.  61  Figure 11. Epifluorescence microscopy of BV-PW1 in filtrates. A) Prefiltered lysate. B) GC50 filtered (1.2 urn pore size). C) GF75 filtered (0.8 um pore size). D) 0.45 um filtered. E) 0.22 um filtered.  Due to the large particle size of BV-PW1, -230 to 300 nm, it is reasonable that the virus did not pass through this filter or cause lysis of cells when reinoculated into fresh cultures. This, however, contradicts the results reported by Garza and Suttle (1995) that 0.22 um filtrates of lysed cultures caused lysis of fresh nanoflagellate cultures. The reason for the discrepancy is unclear, as both situations involved cultured viruses, but perhaps greater force was used by Garza and Suttle (1995) in the filtration process that pushed the BV-PW1 particles through the membrane. The situation of the glass fibre membrane (0.8 um pore size) is also unclear. Perhaps there was a charge associated with the membrane that caused the virus particles and bacteria to remain bound to the filter. Clearly the best membrane for BV-PW1 particle recovery was the 0.45 um GV, most likely due to its low-protein binding surface, however, it was not  62 selective against the bacteria (Figure 11, B, C, D). Because the purpose of filtration was to remove the contaminating bacteria prior to the concentration step, this method was abandoned. Bacteria were clearly still getting through the membrane at abundances of roughly 1 x 10 ml" . 8  1  This additional organic matter from the bacteria also may have created problems in the concentration step. It is possible that the viruses and bacteria alike adsorbed to the spiral membrane accounting for losses of up to 74% between the prefiltration and concentration steps. Due to the poor recoveries and large culture volumes involved, all ultrafiltration efforts were abandoned and concentration of BV-PW1 was carried out with the centrifugation method described in chapter two.  Purification of virus particles  After concentration of the lysates, the problem of separating the BV-PW1 particles from the contaminating bacteria still existed. Initially, sucrose gradient purification was attempted, but most likely due to low virus yields from infected nanoflagellate cultures at that time, no bands on the sucrose gradients were seen. Another method that was investigated for the purification of BV-PW1 made use of the reagent called Bactozyme (Molecular Research Center, Cincinnati, USA). In attempt to cause bacterial cell lysis without harming the integrity of the viral capsids, the Bactozyme solution, containing activated lysozyme, was used following the manufacturer's protocol. Since lysozyme functions in targeting the covalent bonds in the peptidoglycan of bacterial cell walls, it was hypothesized that the viral particles would remain intact. Further support of this hypothesis came from reports that bacteriophage T4 produces lysozyme to disrupt the intermembrane peptidoglycan layer,  63 promoting subsequent entry of phage DNA into the host (Arisaka etal., 2003). Therefore, bacteriophage T4 isn't susceptible to the effects of lysozyme, so perhaps BV-PW1 had a chance of resistance as well. The concentrated lysate was incubated at 50°C for 1 h in the presence of Bactozyme resulting in the lysis of both the bacteria and BV-PW1 particles. Although viruses lack the outer peptidoglycan layer that constitutes a bacterial membrane, it is possible that the lysozyme targeted bonds essential for the structure of the viral capsid or destroyed the virus particles through a different mechanism. Midway through my research, several changes that were made to the method for culturing the nanoflagellates resulted in higher virus yields. It became apparent that higher virus titres were produced from smaller culture volumes, therefore cultures were reduced from 10 I to 600 ml. In addition, a new incubator room became available that provided a constant temperature of 20°C. Under these new conditions, the growth of the flagellates and the production of viruses increased. Previously, the nanoflagellates were subjected to the fluctuating temperatures of the lab. With the resulting high virus yields and the ultraconcentration method outlined in chapter two, bands were finally seen on sucrose gradients. Sucrose gradient purification was used to isolate BV-PW1 particles from that point onward.  Purification of BV-PW1 D N A  A number of different DNA extraction methods were carried out in attempt to obtain a pure, non-degraded sample of genomic BV-PW1 DNA. In all cases, DNA quality was monitored by pulsed-field gel electrophoresis.  64 Initially, the concentrated viral lysates were treated with nucleases (deoxyribonuclease I and ribonuclease A [DNase I and RNase A], each at 1 U ml" final 1  concentration) for 30 min at room temperature before the standard phenol chloroform DNA extraction method as described previously in the material and methods section of chapter two. This step proved to be unnecessary as the same shearing of the DNA result was produced on the PFGE gels as seen in figure 6, lane 7 with or without the nucleases. After this discovery the DNase and RNase treatment was no longer used. Another modified version of the standard phenol chloroform extraction was pursued in attempt to reduce handling of the viral DNA. Effort was taken to remove the bottom non-aqueous layer at each step of the procedure, leaving the interface and the DNA-containing aqueous layer untouched until the very last extraction step, as described previously (Fuhrman  etal.,  1988). Even with these precautions, the final  result on the PFGE gel was the same smearing pattern seen for DNA extracted from the standard protocol (Figure 6, lane 7). Another idea involved the addition of tRNA (1% v/v of 10 mg ml" stock) to the 1  final ethanol precipitation step of the standard phenol chloroform DNA extraction method. Yeast tRNA is often used as a co-precipitant in nucleic acid precipitation, acting as a carrier for small amounts of DNA (Ambion, Austin, USA). Once again the same degraded smear was produced no different than that of DNA extracted without tRNA present. The TEGED (Tris, EGTA [egtazic acic], EDTA [edetic acid]) method for DNA extraction was also considered. Purified viral pellets from the 334,000 x g final ultracentrifugation spin were resuspended in 60 ul of 10x TEGED buffer (100 mM Tris,  65 10 mM EGTA, 10 mM EDTA) and incubated at 60°C for 40 min. Extraction with the DNase and RNase pre-step was also carried out, but again there was no difference seen between the two conditions. Proteinase K (1 % v/v of 20 mg ml" stock) was added 1  to the reaction to help disrupt the capsid proteins and release the viral DNA. One interesting result to note is that without the addition of proteinase K, there was absolutely no DNA seen on the PFGE gel. Only with the addition of proteinase K was there DNA smearing in the lane. After this discovery, proteinase k was always added to all subsequent DNA extraction protocols. The agarose plug method, commonly used for extraction of DNA prior to PFGE (Wommack etal., 1999) was the next method to be tried. Pellets were resuspended in 450 ul of SM buffer (0.1 M NaCI, 8 mM MgS0 - 7 H 0 , 50 mM Tris-HCI, 0.005% [w/v] 4  2  glycerin) and incubated at 4°C overnight. Another millilitre of SM Buffer was added before centrifuging the mixture in a suspended 1.5 ml tube with an SW-40 rotor for 1.5 h at 50,000 x g at 20°C. The supernatant was removed and the pellet was resuspended in 40 |il of SM Buffer and vortexed to mix. Equal volumes of virus concentrate and molten 2.5% NuSieve GTG low melt agarose (Cambrex, East Rutherford, USA), were mixed, dispensed into plug molds, and left to solidify. The plugs were then punched out from the molds into a small volume of buffer (250 mM EDTA, 1% SDS) containing 1 mg ml" proteinase K and incubated in the dark at room temperature overnight. The 1  digestion buffer was decanted and the plugs were washed three times for 30 minutes each in TE buffer. Virus-agarose plugs were stored at 4°C in TE 20:50 buffer (20 mM Tris, 50 mM EDTA, pH 8.0) until loaded into the wells of 0.5x TBE agarose PFGE gels like the DNA extracts from the other methods. Extracted DNA gave the same smeared banding pattern as seen in the other previously described methods.  66 Positive results were not anticipated from the final method used to extract the genomic DNA of BV-PW1, but, in theory, the Trizol LS extraction method for RNA should have worked. The first steps in the method outline how to precipitate DNA from a sample of viral RNA; these steps were followed as per manufacturer's instructions (Life Technologies) and the steps for the purification of the RNA were omitted. The trizol extraction method did not produce any visible DNA on the PFGE gel. In summary, all of the DNA extraction methods produced a smear ranging from -30 kbp to >100 kbp or an empty lane on PFGE gels. This apparent shearing of the genome was seen with all extraction methods even when precautions were taken, such as the use of wide bore tips and not vortexing to mix samples (except in the case of the plug extraction method). There are no obvious reasons as to why this curious phenomenon is happening. The genome size of BV-PW1 is most likely quite large for a virus, but other larger genomes have been resolved as a clean band on PFGE gels, such as the 800 kbp Mimivirus genome (LaScola etal., 2003) and the Phycodnavirus infecting Pyramimonas  algae (560 kbp) (Sandaa etal., 2001). Perhaps the very high A  + T content (-66%) of the BV-PW1 genome makes it more susceptible to shearing. In spite of the putatively sheared genomic DNA, efforts to create a clone library from this sample were still successful. Although it would be beneficial to have the genome size for BV-PW1, it is not necessary for future experimentation and characterization of this virus.  


Citation Scheme:


Citations by CSL (citeproc-js)

Usage Statistics



Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            async >
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:


Related Items