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Characterization and Phylogenetic Relationships of XENORHABDUS BOVIENII Strains (ENTERORBACTERIACEAE, GAMMAPROTEOBACTERI...

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Title: Characterization and Phylogenetic Relationships of XENORHABDUS BOVIENII Strains (ENTERORBACTERIACEAE, GAMMAPROTEOBACTERIA) Based on Sequence Data of Two Protein Coding Genes
Physical Description: Book
Language: English
Creator: Russell, Rachel
Publisher: New College of Florida
Place of Publication: Sarasota, Fla.
Creation Date: 2009
Publication Date: 2009

Subjects

Subjects / Keywords: Steinernema nematode
Xenorhabdus Bovienii
Entomopathogenic
Genre: bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Xenorhabdus spp. are Gram negative gamma-proteobacteria that have a mutualistic association with entomopathogenic nematodes in the genus Steinernema. These bacteria are symbiotically harbored in the intestine of the only free-living stage of nematodes, the infective juvenile. This pair is pathogenic for a range of insects and has been integrated into biological control programs worldwide. Each Steinernema species has an apparent specific association with only one Xenorhabdus species, though a single Xenorhabdus bacterial species may be associated with multiple nematode species. This is the case of X. bovienii, which is present in nine different Steinernema spp. However, why these nematode species share the same bacterial symbiont is not understood. It has been speculated that sharing of X. bovienii could have happened by horizontal transfer of symbionts during co-infection of an insect host. Phylogenies based upon 16S rRNA sequences for Xenorhabdus spp. indicate some variation exists among X. bovienii isolates from different Steinernematids. Nevertheless, this gene is considered too conservative to reflect intraspecific variation. We considered two protein coding genes, rec-A and ser-C, to examine intraspecific variation across nine X. bovienii strains and interpret evolutionary relationships with other Xenorhabdus spp.
Statement of Responsibility: by Rachel Russell
Thesis: Thesis (B.A.) -- New College of Florida, 2009
Electronic Access: RESTRICTED TO NCF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE
Bibliography: Includes bibliographical references.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The New College of Florida, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Local: Faculty Sponsor: Gilchrist, Sandra

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Source Institution: New College of Florida
Holding Location: New College of Florida
Rights Management: Applicable rights reserved.
Classification: local - S.T. 2009 R9
System ID: NCFE004164:00001

Permanent Link: http://ncf.sobek.ufl.edu/NCFE004164/00001

Material Information

Title: Characterization and Phylogenetic Relationships of XENORHABDUS BOVIENII Strains (ENTERORBACTERIACEAE, GAMMAPROTEOBACTERIA) Based on Sequence Data of Two Protein Coding Genes
Physical Description: Book
Language: English
Creator: Russell, Rachel
Publisher: New College of Florida
Place of Publication: Sarasota, Fla.
Creation Date: 2009
Publication Date: 2009

Subjects

Subjects / Keywords: Steinernema nematode
Xenorhabdus Bovienii
Entomopathogenic
Genre: bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Xenorhabdus spp. are Gram negative gamma-proteobacteria that have a mutualistic association with entomopathogenic nematodes in the genus Steinernema. These bacteria are symbiotically harbored in the intestine of the only free-living stage of nematodes, the infective juvenile. This pair is pathogenic for a range of insects and has been integrated into biological control programs worldwide. Each Steinernema species has an apparent specific association with only one Xenorhabdus species, though a single Xenorhabdus bacterial species may be associated with multiple nematode species. This is the case of X. bovienii, which is present in nine different Steinernema spp. However, why these nematode species share the same bacterial symbiont is not understood. It has been speculated that sharing of X. bovienii could have happened by horizontal transfer of symbionts during co-infection of an insect host. Phylogenies based upon 16S rRNA sequences for Xenorhabdus spp. indicate some variation exists among X. bovienii isolates from different Steinernematids. Nevertheless, this gene is considered too conservative to reflect intraspecific variation. We considered two protein coding genes, rec-A and ser-C, to examine intraspecific variation across nine X. bovienii strains and interpret evolutionary relationships with other Xenorhabdus spp.
Statement of Responsibility: by Rachel Russell
Thesis: Thesis (B.A.) -- New College of Florida, 2009
Electronic Access: RESTRICTED TO NCF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE
Bibliography: Includes bibliographical references.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The New College of Florida, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Local: Faculty Sponsor: Gilchrist, Sandra

Record Information

Source Institution: New College of Florida
Holding Location: New College of Florida
Rights Management: Applicable rights reserved.
Classification: local - S.T. 2009 R9
System ID: NCFE004164:00001


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CHARACTERIZATION AND PHYLOGENETIC RELATIONSHIPS OF XENORHABDUS BOVIENII STRAINS (ENTERORBACTERIACEAE, PROTEOBACTERIA) BASED ON SEQUENCE DATA OF TWO PROTEIN CODING GENES BY RACHEL RUSSELL A Thesis Submitted to the Division of Natural Sciences New College of Florida in partial fulllment of the requirements for the degree Bachelor of Arts Under the sponsorship of Sandra Gilchrist Sarasota, Florida May, 2009

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Acknowledgements: I would like to thank Dr. S. Patricia Stock and her lab members for all of their assistance while at the University of Arizona as well as countless hours of guidance and endless patience during my winter and summer internships during 2008. My research was sponsored through funding provided by University of Arizona Undergraduate Biology Research Program (UBRP) Fellowship and National Science Foundation funding to S.P. Stock ( NSF IOS award # 060899). Thank you as well to the members of my thesis committee: Dr. Clore and Dr. Beulig, and to my advisor, Dr. Gilchrist for all of your tireless hard work! ii

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Table of Contents Acknowledgements ii Table of Contents iii Table of Tables v Table of Figures v Abstract vi Introduction 1 II: History of the Taxonomy of Xenorhabdus and Photorhabdus II a: Multilocus Sequencing of Xenorhabdus bovienii 4 9 III: Materials and Methods III a: Axenization Procedure III b: Bacterial Harvesting III c: Bacterial DNA Extraction III d: Bacterial DNA Amplification III e: Sequence Analysis 13 14 19 20 22 24 IV: Results 25 V: Discussion 30 iii

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VI: Future Applications 35 Appendix: Appendix A : Solution Recipes Appendix B: ABI Sequencer Information 37 37 38 Bibliography 43 iv

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Table of Tables: Table of Contents iii Table of Tables v Table of Figures v Table I: Phenotypic Differences between Photorhabdus and Xenorhabdus 8 Table II: Species of Xenorhabdus and Steinernema Host 14 Table III: Primer Sequences of 16s, recA, and serC 23 Table of Figures: Figure I: Nematode/Symbiont Lifecycle 3 Figure II: Steinernema Axenization Procedure 18 Figure III: DNA extraction from Xenorhabdus species. 21 Figure IV: Procedural outline for Xenorhabdus DNA amplification and DNA sequencing. 24 Figure V: 16s rRNA Analysis 26 Figure VI: recA Analysis 27 Figure VII: serC Analysis 28 Figure VIII: Triple Gene Analysis 29 v

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CHARACTERIZATION AND PHYLOGENETIC RELATIONSHIPS OF XENORHABDUS BOVIENII STRAINS (ENTERORBACTERIACEAE, GAMMAPROTEOBACTERIA) BASED ON SEQUENCE DATA OF TWO PROTEIN CODING GENES RACHEL RUSSELL New College of Florida, 2008 Abstract Xenorhabdus spp. are Gram negative gamma-proteobacteria that have a mutualistic association with entomopathogenic nematodes in the genus Steinernema These bacteria are symbiotically harbored in the intestine of the only free-living stage of nematodes, the infective juvenile. This pair is pathogenic for a range of insects and has been integrated into biological control programs worldwide. Each Steinernema species has an apparent specific association with only one Xenorhabdus species, though a single Xenorhabdus bacterial species may be associated with multiple nematode species. This is the case of X. bovienii which is present in nine different Steinernema spp. However, why these nematode species share the same bacterial symbiont is not understood. It has been speculated that sharing of X. bovienii could have happened by horizontal transfer of symbionts during co-infection of an insect host. Phylogenies based vi

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upon 16S rRNA sequences for Xenorhabdus spp. indicate some variation exists among X. bovienii isolates from different Steinernematids Nevertheless, this gene is considered too conservative to reflect intraspecific variation. We considered two protein coding genes, rec-A and ser-C, to examine intraspecific variation across nine X. bovienii strains and interpret evolutionary relationships with other Xenorhabdus spp. Dr. Sandra Gilchrist Natural Sciences vii

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Introduction: !Nematodes are among the most successful animals on Earth, both with regard to their biomass and biodiversity. Within one handful of soil hundreds of nematodes can be found, living freely or waiting for a host to continue their lifecycle. Among the many ecological niches that nematodes occupy, one specialization that remains quite ubiquitous is that of insect pathogenicity. It is hypothesized that one of the most instrumental adaptations to the continued evolutionary success of these nematodes was the capacity to engage microbial companions (Doris and Baxter 1999). !At almost all taxonomic levels, representatives are present that harbor microorganisms. However, the nature of this afliation varies highly from parasitic, to symbiotic, to mutualistic, all of which reect differing levels of host benet. A fascinating aspect of this relationship is the way in which the symbiont species undergoes specialization to the host environment, possibly through the exchange of genetic material. Unique among the rhabditids, entomopathogenic nematodes have a symbiotic relationship with an enteric bacterium species (Burnell and Stock 2000). The mutualistic relationship between entomopathogenic Steinernema nematode species ( Nematoda: Steinernematidae ) and Xenorhabdus bacteria species (gram-negative Enterobacteriaceae) is an emerging model of such an intimate relationship. Steinernema species have a global distribution, and exhibit differences in host range, infectivity, environmental tolerances and in suitability for commercial production and formulation. This has stimulated many surveys, seeking new 1

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strains and species of entomopathogenic nematodes for biocontrol applications (Hominick et al. 1996). For the nematode life cycle to propagate, the bacterial symbiont is required to kill the insect host and liquify the host tissues and structures. This procedure then provides suitable nutrient conditions for nematode growth and development. Although this relationship is assuredly much more complex than what is currently understood thus far, it seems that one of the most integral components to the success of this organism pair is the Infective Juvenile, or IJ (Fig. I). In the life cycle, the IJ is the only free-living stage that is concurrently non-reproductive and unable to feed. For the life cycle to continue to propagate, the IJ must be colonized by symbiotic bacteria which are held within the bacterial vesicles. At this stage, the IJ can then wait in the soil for a suitable host (Poinar 1966; Wouts, 1980). The range of hosts vary widely from species to species, but one constant is that the IJ must gain entrance to the site of infection (the insect circulatory system, known as the hemocoel) via the mouth, anus or spiracles of the host. Once inside the hemocoel, or insect body cavity, the IJ recover into a feeding stage, release the bacteria from the receptacle into the hemolymph fluid of the insect, and soon the insect expires (Forst and Tabatabai 1997; Goodrich-Blair and Clarke 2007; Kaya and Gaugler 1993). Interestingly, antibiotic activity is associated with metabolites of Xenorhabdus species, thereby reducing competition of other invading bacteria (Akhurst 1982; Boemare et al. 1992; Webster et al. 2002). After insect death, the invading IJ feed and begin to reproduce, leading to juvenile and adult stages. These stages do not undergo 2

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colonization by the bacterial symbiont, but instead reproduce until a scarcity of food is sensed. At this point, IJ are then produced, and the cycle begins again. Nematode Life Cycle Infective Juvenile infects insect, and symbiotic bacteria is released. Insect succumbs to bacterial toxins, and Infective Juveniles feed and reproduce. Several generations of nematode reproduce within the cadaver. When resources diminish, Infective Juveniles are produced that contain an Intestinal Vesicle (shown here). This vesicle is then colonized by symbiotic bacteria. The Infective Juveniles then emerge. Figure I: Nematode/Symbiont Lifecycle Photos and Diagram source: S.P. Stock University of AZ 3

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II: History of the Taxonomy of Xenorhabdus and Photorhabdus Since the initial discovery and description of Xenorhabdus by Thomas and Poinar in 1979, species have been rapidly isolated from entomopathogenic nematodes worldwide (Akhurst and Boemare 1988, Boemare and Akhurst 1988). For this reason the number of species within the genus Xenorhabdus is constantly expanding. To date, the four most prevalent species are Xenorhabdus nematophilus X. bovienii X. poinarii and X. beddingii (Akhurst and Boemare 1988). Because of its close relationship to the genus Xenorhabdus, Photorhabdus was previously characterized as Xenorhabdus luminescens and the taxonomy of these genera has sometimes been a point of contention among researchers (Forst and Nealson 2006). For this reason, several methods have been utilized to attempt to solidify the taxonomic positioning of Photorhabdus and Xenorhabdus Aside from analysis of morphology, genomic comparisons have been very useful for determining the relatedness of these two genera. As the 16s rRNA gene sequences has had great success in determining phylogenetic relationship among eukaryotic and prokaryotic organisms, this technique was quickly implemented and phylogenetic analyses of Photorhabdus and Xenorhabdus were performed (Rainey et al. 1995; Woese 1984 ). These results coupled with oligonucleotide cataloging (Ehlers et al. 1998) have placed the Xenorhabdus/ Photorhabdus group within the gamma subdivision of proteobacteria. Aside from this assignment, inference based on the oligonucleotide cataloging results, led to the proposal that Xenorhabdus/Photorhabdus constituted a taxonomic unit possibly separate from the family Enterobacteriaceae (Forst and Nealson 2006). 4

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Interestingly, 16S rRNA data published by Rainey and colleagues in 1995, portrayed the relationship of Xenorhabdus/Photorhabdus as very close to each other, as well as closely related to other members of the gamma-Proteobacteria, of which many were symbionts and/or pathogens of eukaryotes. Farmer (Farmer 1984) and Farmer and colleagues (1989) have pointed out that the inability of Xenorhabdus spp. to reduce nitrate, coupled with their catalase-negative property, is a reason to question their inclusion in the Enterobacteriaceae. Similarly, Janse and Smits (1990) analyzed whole-cell fatty acid patterns. Interestingly, in contrast to the accepted view, they suggested that the Xenorhabdus spp. might be sufciently different from other enterobacteria to possibly exclude them from the Enterobacteriaceae. Having had a somewhat confusing past, the taxonomy of the Xenorhabdus/ Photorhabdus group has often rapidly undergone change in regards to phylogenetic relationships and nomenclature. One of the rst published discussions of bacteria associated with entomopathogenic nematodes appeared in 1959 (Dutky 1959), in which the nematodes studied were labeled Neoaplectana (later to be known as the Steinernema ) group. Also noted in this report, was the ability of the bacteria to produce antibiotics and inhibit the putrefaction of the insect carcasses. This nematode (now named Steinernema carpocapsae ), was later used by Poinar and Thomas (1965) as the source material for isolation of a bacterium referred to as Achromobacter nematophilus Inferring from what is now known, it seems likely that the bacteria were Xenorhabdus nematophilus 5

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As per the recommendation of Hendrie and colleagues (1974), the genus Achromobacter was no longer considered and the symbionts were left without a name, until the proposal of the genus name Xenorhabdus by Thomas and Poinar in 1979 (Kahn and Brooks 1974; Poinar et al. 1977; Thomas and Poinar 1979 ). To accommodate the observation of both luminous and nonluminous isolations, the nonluminous were named X. nematophilus, and the luminous were assigned X. luminescens Soon, it was recognized that heterorhabditid nematodes possessed luminous Xenorhabdus while the nonluminous X. nematophilus isolates were found associated with neoaplectanid nematodes (Thomas and Poinar 1979). The diversity of Xenorhabdus types became apparent in the early 1980s (Akhurst 1983; Akhurst and Boemare 1986; Boemare and Akhurst 1988; Grimont et al. 1984), when several subspecies of X. nematophilus were proposed on the basis of differences in phenotypes. Also of significance, about this time, the nematode genus Neoaplectana was replaced by the name Steinernema. Using DNA-DNA hybridization, it was then found that groups of Xenorhabdus once considered to constitute subspecies, should be given species status (Boemare et al 1983), as well as it was first proposed that luminous Photorhabdus luminescens should be placed in a separate genus. This proposal was then further supported by the observation that Photorhabdus luminescens strains are nitrate reduction negative, pigment positive, and bioluminescence positive. 6

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Using a different approach, Wimpee and colleagues (1991) showed that the luciferase (luxA) gene probe was positive for all strains of P. luminescens tested and showed no hybridization with any of the Xenorhabdus species. These results were confirmed by Ragudo (1999) when the genes coding for pigment formation in P. luminescens showed no cross-hybridization with any Xenorhabdus strains. As more genes are isolated and sequenced, it will be of interest to conduct crosshybridization studies between the Xenorhabdus and Photorhabdus isolates, as well as between pairs of species isolated from different environments. Because, at the phenotypic level, the traits of Photorhabdus and Xenorhabdus appear similar, but yet at the genetic level are quite distinctive, it would seem that these organisms present a very precarious position terms of their evolution. One possible view is that the niche they inhabit is one that was available and hospitable to bacteria of the gamma proteobacteria types of organisms. This type of generic environment would also help to explain the similarities observed via the 16S rRNA approach. However, it is also quite likely that convergent evolution has led to the observed similarities. Some similarities and differences between Photorhabdus and Xenorhabdus are presented in Table I. For this reason, it would be quite likely that although similar in function, the genotype would most likely present a difference in structure composition, as well as a difference in origin. Whether the convergence is due to lateral gene transfer, or another mechanism, it is most likely that future molecular studies will most likely reveal the details of gene origin (Forst and Nealson 2006). 7

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Phenotype Photorhabdus Xenorhabdus catalase positive negative bioluminescence positive negative pigments anthraquinones negative antibiotics hydroxystilbenes xenocoumacins xenorhabdins indole derivatives crystal proteins 11.6 kDa 26 kDa 11.3 kDa 22 kDa urease activity some species negative indole production some species negative aesculin hydrolysis some species negative Table I: Phenotypic Differences between Photorhabdus and Xenorhabdus (Forst and Clark, 2002) Having been more actively studied than Xenorhabdus it has been observed that the genome sequence of Photorhabdus luminescens contains a large number of mobile genetic elements, with phage remnants accounting for 4% of the genome and 195 insertion sequences or IS fragments present on the chromosome, suggesting ongoing gene transfer (Duchaud et al. 2003). It has also been observed that Photorhabdus strains contain many genomic islands, which are presumably acquired via horizontal transmission (Waterfield et al. 2002). For instance, the Photorhabdus genome contains 11 loci predicted to encode for fimbria or pili, as would be necessary for the attachment of cells involved in transmission (Duchaud et al. 2003). Whole-genome comparisons between Photorhabdus species have shown fimbrial-encoding loci are often part of the variable region of the genome. This observation would imply that these loci may have a role in the evolved specificity between the bacteria and the nematode (Gaudriault et al. 2006). 8

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As nematology continues to expand, this type of study may be applied to Xenorhabdus to determine if a similar scenario was responsible for the evolutionarily convergent genera of Xenorhabdus This type of study would be particularly interesting, as taxonomic research indicates that each Steinernema species has a natural association with only one Xenorhabdus species, though a single Xenorhabdus bacterial species may be associated with more than one nematode species (Stock and Hunt 2005; Stock and Goodrich-Blair 2008). II a: Multilocus Sequencing of Xenorhabdus bovienii Interestingly, several Xenorhabdus species have multiple Steinernema species as hosts. For example, Xenorhabdus bovienii is shared by nine Steinernema nematodes. Until now, it is not known why all of these nematode species share the same bacterial symbionts. However, it has been speculated that sharing of X. bovienii could have happened by horizontal transfer of the symbionts during co-infection of an insect host (Stock personal communication ) Molecular studies based on 16S rRNA sequence data indicate there is variation among X. bovienii isolates belonging to different Steinernema hosts. Previous studies suggest association between the nematodes and their natural bacterial symbionts may be highly specialized. Sicard and co-workers (2006) conducted interspecies competition studies and concluded that nematode fitness is highly dependent on their natural symbionts. In this study, it was observed that nematodes reared with symbionts of another nematode species do not perform as well as with their own natural symbionts, this having been 9

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reflected in their generational offspring production. However, no studies have yet been published to assess competition of Xenorhabdus symbionts at the intraspecific level. We attempted such a study, in which species of Steinernema that are naturally paired with Xenorhabdus bovienii were reared in absence of symbiotic bacteria, in other words were axenized. These species would have then been exposed to a non-native strain of X.bovienii, and observed to assess reproductive fitness. However in the process of rearing the axenic species, a significant contamination issue was observed and could not be rectified in a timely fashion. Therefore, unfortunately this study was abandoned. The source of contamination, which at the time was viewed as disastrous, later yielded insight into the project that composes the focus of this work. The specific goal of this study was to evaluate genetic variation of X. bovienii strains that are symbionts of different Steinernema species considering the 16s ribosomal gene and two protein-coding genes: recA and serC. Although the type strains have been well studied, and both an Xenorhabdus nematophila strain and an Xenorhabdus bovienii strain have been sequenced ( http:xenorhabdus.danforthcenter.org ), the extent of diversity within the species is largely unknown. Therefore, whether populations exhibit enough diversity to warrant screening for novel toxins and/or antibiotics, not present in the type strains, is not yet known. The 16s ribosomal RNA gene is widely used for phylogenetic studies as it is highly conserved, with several species specific variable regions that allows for 10

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comparative analysis (Sergeant et al. 2006). As a result, 16S rRNA gene sequencing has become prevalent in medical microbiology as a rapid and fairly accurate alternative to phenotypic methods of bacterial identification. However because this gene is highly conserved within species of bacteria, a more variable gene is needed to determine the presence of subspecies more accurately. One canidate, the recA gene catalyzes the hydrolysis of ATP in the presence of single-stranded DNA, the ATP-dependent uptake of single-stranded DNA by duplex DNA, and the ATP-dependent hybridization of homologous single-stranded DNAs. It interacts with lexA causing its activation and leading to its autocatalytic cleavage. It also plays a role in DNA recombination, repair, the SOS response, DNA dependent ATPase activity, as well as the binding of singlestranded DNA ( http://xenorhabdus.danforthcenter.org ). This gene is known to be less conservative than the 16s rRNA gene. Another candidate, the serC gene product is thought to synthesize serine, which is required for growth in the vesicular environment. It is known that a serC mutant strain cannot colonize the IJ in absence of X. nematophila (Kim and Forst 2005). Because this gene is believed to govern this highly specialized nematode symbiont interaction, it is thought to be extremely variable. Statistical analysis done on this gene of ten specimens of X. bovienii by Sergeant and colleagues (2006) implied that recombination had occurred at the serC locus and that moderate amounts of interallele recombination most likely takes place. Liu and co-workers (1997) noticed that there may also be an apparent geographical correlation within possible clades of Steinernema and 11

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Heterorhabditis symbiotic bacteria. These clades correlated with host nematode species as follows: clade 1, for symbiotic bacteria of Steinemema feltiae and Steinemema intemedia equivalent to X. bovienii ; clade 2, for symbiotic bacteria of Steinemema glasen equivalent to X poinarii ; clade 3, for symbiotic bacteria of Steinemema carpacapsae and Steinemema riobravis equivalent to X. nematophilus ; clade 4, for symbiotic bacteria Heterorhabditis marelatus and four other isolates from the Oregon coast region, equivalent to an undescribed Photorhabdus sp.; clade 5, for symbiotic bacteria of Heterorhabditis megidis equivalent to Photorhabdus luminescens ; and clade 6, for symbiotic bacteria of two isolates of Heterorhabditis indicus two isolates of Heterorhabditis bacteriophora and four isolates of Heterorhabditis spp. from noncoastal regions, equivalent to Photorhabdus luminescens Because these clades are associated with different nematode species, this may be initial evidence of coevolution between the nematodes and their symbiotic bacteria (Liu et al. 1997). The close correspondence between the taxonomic grouping of the bacteria and the taxonomic grouping of their nematode associates has been confirmed by phenotypic studies (Akhurst et al. 1983; Akhurst and Boemare 1983; Boemare and Akhurst 1988). Multilocus sequence typing (MLST) types bacteria based on the sequence of a number of housekeeping genes and is thought to reect the phylogeny of strains more accurately than other methods (Seargent et al. 2006). For this reason, it was determined that this procedure would be most appropriate to 12

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accomplish the goal of this research, which was to assess the diversity within Xenorhabdus bovienii. III: Materials and Methods For the current study, the 16s rRNA gene as well as the protein coding recA and serC genes for the species presented in Table II were sequenced to determine a possible phylogenetic relationship. However, before beginning the MLST of the Xenorhabdus species, we attempted to rear axenic samples of the Steinernema listed in Table II. These species were raised on cultured Xenorhabdus nematophila while we were under the impression that this species of Xenorhabdus would not allow bacterial colonization of the IJ. This attempt at rearing generations of axenic nematodes was conducted as preparation for a competition experiment in which the ultimate goal would have been to assess the reproductive fitness of species of Steinernema nematodes that contain a different strain of their native symbiotic bacteria. This study would have been similar to that conducted by Sicard and colleagues in 2006, but expanded to compare Steinernema species that contain a different strain X. bovienii. The procedure for axenization follows Table II, and is described in subsection III a, which is followed by III b outlining the procedure for Xenorhabdus extraction from Steinernema IJ. 13

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Xenorhabdus Species Steinernema Species Steinernema Strain Steinernema Host Clade reference bovienii weiserii Turkey III Mr‡cek et al. 2003 bovienii feltiae Canada 14 lig 7A III Nguyen et al. 2006 bovienii feltiae Florida, USA III Stock pers com bovienii feltiae Anatolia, Turkey III Hazir et al. 2003 bovienii affine United Kingdom I Poinar 1988 bovienii feltiae SN France III Poinar 1989 bovienii feltiae Bodega Bay, Jordan III Hazir et al. 2003 bovienii oregonense Bubbling Ponds, Arizona, USA III Liu and Berry 1996 bovienii jollieti Monsanto, Missouri III Spiridonov et al. 2004 szentirmaii rarum Sargento Cabral, Argentina III Lengyel et al. 2005 doucetiae diaprepesi Florida, USA V Nguyen and Duncan DATE nematophila anatoliense Anatolia, Turkey IV Hazir et al. 2003 nematophila carpocapsae A10, Wisconsin, USA II Poinar 1967 cabanillasi riobrave Texas, USA Cabanillas et al 1994 Table II: Species of Xenorhabdus and Steinernema Host III a:Nematode Axenization This procedure was modified by S. Kim (S. Patricia Stock Lab, University of Arizona) from a procedure created by D. Renneckar (H. Goodrich-Blair Lab, University of Wisconsin, Madison). Samples of infective juveniles (IJ) from nine species of Steinernema were obtained from stock cultures which contain 14

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individuals from either in vitro or in vivo propagation (Figure II). First examined for viability (mortality level, population density etc. ) using low magnification, the samples that were taken from the stock collection were stored in distilled water from 15 ¡ C storage. Once samples in good condition were verified, 1 mL of nematode stock solution was added to a 10 cm x 2 cm petri dish or 0.5 mL was added 5 cm x 1 cm petri dish that was inverted, and the lid lined with two appropriately sized filter papers Greater wax moth larvae, Galleria melonella (L.) ( Lepidoptera: Pyrallidae ) were surface-sterilized after a brief dip in 95% ethanol to attempt to ensure that the nematodes that infest the larvae will not succumb to illness from harmful bacteria and viruses. Ten larvae were then added to the larger petri dishes (10 cm. diam.), while five larvae were added to the smaller petri dish (5cm diam.). Petri dishes were then isolated in ziplock bags to prevent dessication and incubated in dark 25 ¡ C storage. They were checked daily after initial three days had passed in which initial infection and one to two generations had usually taken place. Daily observation consisted of dissection of one cadaver until an adult female nematode was obtained and characterized as either gravid or not gravid. Also upon first observation, any larvae that survived the initial infection were removed, while those that had obviously succumbed to other causes of death (such as bacterial infection) were removed to prevent further contamination. Each dissection was conducted using deep concave glass with M9 buffer (see Appendix for description of M9), and the cadaver agitated to induce the 15

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nematodes to enter the buffer solution. Gravid females were then removed and placed in fresh M9 buffer to await introduction into axenizing solution. The axenizing solution (see Appendix for description of Axenizing Solution) was freshly prepared before each axenization to prevent microbial growth. During agitation this solution effectively disintegrates the collagen rich nematode cuticle. Approximately 1 mL of M9 buffer nematode solution (containing approximately 150 females) was then added to approximately 12 mL of axenizing solution, and subsequently centrifuged for ten minutes at 13,000 rpm. The supernatant was then discarded, and the egg pellet resuspended with 13 mL of axenizing solution and constantly agitated for ten minutes. This solution was centrifuged at 13,000 rpm for another ten minutes, and the supernatant discarded. The egg pellet was then resuspended with 13 mL of sterile LB broth, agitated continuously for ten minutes, and centrifuged for another ten minutes at 13,000 rpm. After discarding the supernatant, the pellet was transferred to 1.5 mL microfuge tube, resuspended in 1 mL of sterile Luria B ertani (LB) broth (see Appendix), and centrifuged at 2,000 revs/min for one minute. The supernatant was then discarded, and the pellet was resuspended with 1 mL of sterile LB broth. This wash was repeated five times. Upon the discard of the final wash, 1 mL of sterile LB broth was added, and this solution added to a 6 cm sterile petri dish containing 4 mL of sterile LB broth and 5 L of ampicillin (concentration 150 mg/mL). The eggs in solution were then allowed to hatch for approximately 48 hours before extraction of the juveniles from the broth. The extracted juveniles were subsequently washed three times with 1 mL of sterile LB as above, and 5 16

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" L removed for counting under the dissecting scope. After counting, the concentration of axenic juveniles was assessed and the number of bacterial cultures needed to achieve the desired concentration of 1,000 juveniles per culture determined. The axenic nematodes were then added to Xenorhabdus nematophila cultures grown on Lipid Agar (see Appendix), and placed in 28C dark storage. The nematode/bacteria cultures were then observed 48 hours later for signs of nematode inhabitance. 17

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! !!!! !! !! !!!!! !!!! !!!!!! Dissection of a Steinernema infected cadaver to recover gravid females for axenization procedure. S. kraussei gravid female. Gravid females are introduced into axenizing solution, then subsequently agitated, centrifuged, and the waste decanted. Luria Bertani broth is then added before the nal centrifugation. The LB broth is then decanted, and the eggs are condensed into an microfuge tube with a small amount of LB broth for nal centrifugation repeats. Photo by S. Kim, University of AZ After several LB washes, the eggs are injected onto LB agar plates for rearing of several generations. Shown are adult male and female S.oregonense Photo by R. Russell Nematode Axenization Procedure. Photo by S. Kim, University of AZ Figure II: Steinernema Axenization Procedure. 18

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III b: Bacterial Harvesting This procedure was modified by Samkyu Kim (of S. Patricia Stock Lab of the University of Arizona) from a procedure created by Darby Renneckar of the Heidi Goodrich-Blair Lab (University of Wisconsin, Madison). After the initial infection of insects and the first generations of Steinernema had reproduced, cadavers from each group were placed in white traps for collection of IJ colonized by X. bovienii White traps were constructed from 10 cm x 20 mm petri dish bases with a superficial layer of distilled water added to surround an uncovered 50mm x 10mm petri dish base which was lined with 2 sheets of filter paper, on which 6 to 10 properly infected larvae were arranged (Figure III). Once infective juveniles were observed in the surrounding distilled water, the solution was collected for use in symbiotic bacteria extraction. Once collected, the infective juvenile solution was washed three times with distilled water, and then aliquoted to produce the concentration of 1,000 nematodes to 1 mL, in a 15 mL glass centrifuge tube. The IJ were then surface sterilized by adding approximately 12 mL of a 0.5% NaOCl solution was then added to the tubes for 3 minutes, before Millipore vacuum filtration with three distilled water rinses. After the third wash was decanted, the IJ were removed from the filter in approximately 1 mL of distilled water, and placed in 5 mL of sterile LB broth (see Appendix). The surface sterilized IJ were then sonicated for two minutes, and if cuticle disruption was complete, a milky white solution was produced. 19

PAGE 27

Aliquots of 20 l of this solution as well as 5 microliters of ampicillin solution (concentration of 100 g/ml) were then plated on NBTA (Appendix) plates. The NBTA plates were then placed in 30 ¡ C dark storage to ensure proper growth. After two days had passed (2 days is peak Xenorhabdus bovienii colonization) (Sergeant and colleagues 2006), plates that showed evidence of characteristic Xenorhabdus colonies (blue, anaerobic and small) were subsequently subcultured by placing one small colony in 2 mL of fresh sterile LB broth in a dark incubator at 30 ¡ C and rotated at 120 rpm. After 48 hours, the tube was then removed and plated on NBTA. This process was repeated until only blue colonies were present on the NBTA plate, and then was repeated once or twice more. From these plates, colonies were once again subcultured in 2 mL of LB broth, and then prepared for the DNA phenol chloroform extraction. III c: Bacterial DNA extraction The procedure of the phenol chloroform extraction began with the transfer of the 1 mL of bacteria culture into a microcentrifuge tube (Figure III). The tube was then placed in a centrifuge for five minutes at 10,000 rev/min. The supernatant was then discarded. Then, to the pellet while stirring, 500 L of TE buffer (pH 8), as well as 15 L of 20% SDS, and 20 L of proteinase K added. This solution was then incubated at 55 0 C for 30 minutes. The pellet then underwent three freezing cycles in liquid nitrogen for ten minutes before thawing at 85 0 C. The bacterial solution was then vortexed before performing the extraction with a ratio of 25:24:1 phenol: chloroform: isoamyl alcohol. 20

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Figure III: DNA extraction from Xenorhabdus species. In this procedure, Xenorhabdus colonies were isolated from IJ, subsequently cultured, and the DNA extracted and quantified. This solution was then centrifuged for ten minutes at 13,000 revs/min, and the supernatant transfered to a new tube (the pellet was discarded). An equal 21

PAGE 29

volume of 24:1 chloroform: isoamyl alcohol was then stirred into the supernatant, and the combined solution centrifuged for ten minutes at 13,000 revs/min. The supernatant was then pipetted 100 L at a time to a new tube, while the pellet was discarded. Sodium acetate was added (to encourage precipitation) in the concentration of 10 L per every 100 L of supernatant recovered. One mL of ethanol was then added, before incubating at 20 0 C overnight. The pellet was then dried in the dessicator for another 12 hours. Following dessication, the was resuspended in 50 L of sterile molecular grade water, and RNAse added to a final concentration of 20 L/mL. This solution was then incubated for one hour at room temperature. The concentration of DNA was then assessed (in ng/" L) using the Nano Drop spectrophotometer. If needed, the concentration of DNA was diluted with molecular grade water to approximately 200 ng/ L (Figure III). At this point the DNA extract was either placed in storage (at -20¡C) for later processing, or prepared for amplification via Polymerase Chain Reaction (PCR). III d: Bacterial DNA amplification An internal fragment of the 16S rRNA, recA, and serC housekeeping genes was amplified by PCR. Each reaction was carried out in a 50 L volume containing 1 M of each primer (Table III), 200 ng chromosomal DNA, and 12 L of Red Taq Ready Mix. PCR cycles consisted of 94¡C for 2 minutes and 25 cycles of 94¡C for 15 seconds, 50 to 55¡C for 30 seconds, and 72¡C for 45 seconds. The PCR products were then purified for sequencing with Exo-Sap, in the concentration of 2 L per 5 L of product. Sequencing reactions were 22

PAGE 30

performed using dye-terminator sequencing chemistry, and reaction products were separated and detected using an ABI 3730 capillary DNA sequencer at the University of Arizona Sequencing Facility. Sequence contigs were assembled using SeqMan software (DNA Star, Inc) (Figure IV). Gene Forward Primer Reverse Primer Fragment Size (BP) Annealing Temperature 16S rRNA GATGGAGGG GGATAACCAC T TTGTCCAGGG GGCCGCCT 720 54¡C recA CCAATGGGC CGTATTGTTG A TCATACGGAT CTGGTTGATG AA 420 54¡C serC CCACCAGCAA CTTTGTGTCC TTTC AAAGAAGCAG AAAAATATTG CA 670 53¡C Table III: Primer Sequences of 16s, recA, and serC (purchased from Invitrogen) 23

PAGE 31

Figure IV: Procedural outline for Xenorhabdus DNA amplification and DNA sequencing. Photos taken by R. Russell III e: Sequence Analysis Support for nodes of the trees was evaluated by bootstrap analysis with 500 replications using a heuristic search and by examination of all trees 1 % longer than the shortest trees. When several most-parsimonious trees 24

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had been obtained, a 50% consensus tree was formed and treated as the most parsimonious tree for constructing figures (Liu et al 1997). IV: Results The preliminary experiment in which species of Steinernema nematodes were axenized yielded surprising results. The axenization procedure was constructed such that all nematode adults, juveniles and X.bovienii were disintegrated, leaving only unhatched eggs. Although IJ obtained from the axenization procedure were likely axenic prior to growth on the bacterial culture of Xenorhabdus nematophila it was observed via microscope that bacteria were present in the vesicle of several juveniles from different species. It was determined that the colonies that were present were most likely from the growth culture of X. nematophila It then became clear that the competition experiment between what needed to be axenic IJ and symbiotic Xenorhabdus bovienii from other species of nematodes could not be completed under the assumption that the nematodes were initially axenic. Unfortunately, we were unable to extract these bacteria from the individuals in which they were observed via microscope. Upon the decision to pursue the molecular genetics approach to attempt to elucidate the evolutionary relationship between X. bovienii bacteria and Steinernema nematode, the Xenorhabdus isolates were initially compared to each other and characterized by sequence analysis of the 16S rRNA gene. These results can be seen in Figure V. As expected, the strains of X. bovienii clustered with each other, while the same behavior was observed for the other species of Xenorhabdus Also as expected, strains of the same Xenorhabdus 25

PAGE 33

species (isolated from different locations) clustered together. The strains of X.bovienii are phenotypically indistinguishable from other strains of the same species (Sergeant et al. 2006). /-------------------------------------------------------------------bovienii.feltiae.anatolia | +-------------------------------------------------------------------X.bovienii.weiseri | +-------------------------------------------------------------------X.bovienii.feltiae.14lig7A | +-------------------------------------------------------------------X.bovienii.feltiae.4reg6A | +-------------------------------------------------------------------X.bovienii.affine.uk | +-------------------------------------------------------------------X.bovienii.feltiae.4reg6A | +-------------------------------------------------------------------X.bovienii.feltiae.sn.france | | /-----------X.bovienii.oregonense.bub +---------------------------83---------------------------+ | \----------X.bovienii.feltiae.bodega | | /-------------------------------------------------------X.bovienii.jollieti | | | | /-----------------------------------------------X.szentirmaii.rarum \----86-+ | | | /-------------------------X.doucetiae.diaprepesi \-100---+ | | /----70----+ /--------------X. nematophila.anatoliense | | \----99--+ \-100--+ \--------------X.nematophila.carpocapsae | \---------------------------------X.cabanillasi.riobrave Figure V: 16s rRNA Analysis The housekeeping gene recA was then sequenced, and the results presented in Figure VI. The tree generated from these data also demonstrated clear homoplasies amongst strains of the same species. 26

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/-------------------------------------------------------------------X.bovienii.feltiae.fl | | /--------------------------------------------------------X.bovienii.feltiae.anatolia | | | | /----------X.szentirmaii.rarum | | /----61-+ | | | \----------X.cabanillasi.riobrave | | | | | +----------------------X.doucetiae.diaprepesi | | /---100+ | | | | /----------X.nematophila.carpocapsae | | | | | | | | \-100--+----------Xenorhabdus.nematophila(ref) | | /----67+ | | | | | \----------X.nematophila.anatoliense | | | | | | | \---------------------------------X.bovienii.jollieti | | | \--------+ +--------------------------------------------X.bovienii.kraussei | | | +--------------------------------------------X.feltiae.canada.4reg | | | | /----------X.bovienii.oregonense.bubbling +----91+----------------66-------+ | | \----------X.bovienii.feltiae.bodega | | | +--------------------------------------------X.bovienii.feltiae.can.14lig | | | +--------------------------------------------X.bovienii.affine.uk | | | +--------------------------------------------Xenorhabdus.bovi(ref) | | | \---------------------------------------------X.bovienii.feltiae.sn | \--------------------------------------------------------X.bovienii.weiserii Figure VI: recA Analysis Sequencing of serC on the other hand, depicted a much more complex evolutionary relationship between strains of X.bovienii as well as the relationship 27

PAGE 35

of X.bovienii with other species of Xenorhabdus These results are presented in Figure VII. /-------------------------------------------------------------------X.cabanillasi.riobrave | | /---------X.doucetiae.diaprepesi | /------------------69------------------+ | | \---------X.szentirmaii.rarum | | | | /---------X.bovienii.feltiae.fl | | | | | /---95---+---------X.bovienii.feltiae.canada.4reg | | | | | | | \---------X.bovienii.weiseri | | /-94-+ | /-97--+ | \---------X.bovienii.affine.UK | | | | | | | /---63----+ | | | | | | | | | \----------------------------X.bovienii.jollieti | | | | | | | | /------------------X.bovienii.feltiae.can.14lig | | 100---+ | \-------+ | /---75-+ /---------X.bovienii.feltiae.sn.france | | | | | | | | \--72+ | \--94--+ | | | \---------X.bovienii.feltiae.anatolia | | | | /---------X.bovienii.oregonense.bubbl | \-------100--------+ | \---------X.bovienii.feltiae.bodegabay | | /------------------X.nematophila.anatoliense | | \-----------------100---------+ /---------X.nematophila (ref) \--69+ \---------X.nematophila.carpocapsae.A10 Figure VII: serC Analysis 28

PAGE 36

The sequences of the three genes were then connected contiguously to assemble a triple gene phylogenetic tree (Figure VIII). /-------------------------------------------------------------------X.cabanillasi.riobrave | | /-----------X.bovienii.feltiae.can.14li | | | /--94--+ | | | /-----X.bovienii.feltiae.anatolia | | \-85--+ | | \-----X.bovienii.feltiae.sn | /----94--+ | | | /-----X.bovienii.oregonense.bub | | \----100--+ | | \-----X.bovienii.feltiae.bodegabay | /-54-+ | | | /-----------X.bovienii.feltiae.canada.4re | | | | | | +-----92-----+ /-----X.bovienii.weiseri | /--------100-+ | \--97-----+ | | | | \-----X.bovienii.feltiae.fl | | | | | /-80--+ | \------------------------------X.bovienii.affine | | | | | | | \------------------------------------X.bovienii.jollieti | | | | /-89+ \--------------------------------------------X.szentirmaii.rarum | | | | | \-------------------------------------------------------X.doucetiae.diaprepesi \-+ | /-----X.nematophila.carpocapsae \--------------------------100--------------+ \-----X.nematophila.anatoliense Figure VIII: Triple Gene Analysis 29

PAGE 37

V: Discussion To fully contextualize the genetic relationships between the X.bovienii isolates, a sense of the distribution, or biogeography, of the nematode/bacteris pair is required. In this respect, Steinernema have a very wide distribution, having been recovered from all continents except Antarctica (Burnell and Stock 2000). Within this genus, two species appear to have the most global distribution: Steinernema feltiae and Steinernema carpocapsae (Hominick et al. 1996). At this point, according to the academic community, the remaining Steinernema species appear to represent a more restricted geographical distribution, with observations recorded at the national or sometimes continental level only (Hominick et al. 2002). However as surveying and research expands, nematodes that were once considered to be limited in distribution, may soon be isolated on other continents and thus represent a global distribution. One example of such a case that has already occurred is that of Steinernema kraussei Steinernema kraussei was originally thought to be limited in habitat to the Geggen Mountains, Westphalia, in Germany only. However, as research expanded, this species was soon recorded in many other locations in Germany as well as other European countries, such as the Czech Republic, the Netherlands Switzerland, the United Kingdom, and Spain (Hominick 1996). This species was then thought to represent a Paleoarctic distribution. Then, following more research, the geographic range was further expanded to include North America (Stock and Burnell 2000) and thought to represent a Holarctic distribution. It is thought by 30

PAGE 38

Stock and Burnell, that habitat preferences may reflect not only the distribution of suitable insect hosts, but also physiological and behavioral needs that require species certain niches. However, as a rule, biogeographical correlations are difficult to draw given that many factors such as spatial distribution, seasonality, and sampling size differences are often variable and uncontrollable. Furthermore, complications easily arise due to misidentification of species due to morphological or genetic differences. In this respect, although several studies have documented what appears to be preferences of several species for different habitats, these data are at times contradictory and require much more study before a definitive correlation can be drawn (Hominick et al. 2002). It is also of note that some Steinernema species prefer a wider range in habitat than other less distributed species. One example of such an observation in that of S. kraussei which has been found in both coniferous and deciduous forests as well as in grassland areas (Stock and Burnell 2000) as compared to S.rarum which is found only in the warm temperate climate of Argentina (Stock, personal communication). Given the diversity of the habitat preference as well as the variability at the species level of the adaptive ability of Steinernema species, our attempt to evaluate the genetic diversity of their shared symbiont, Xenorhabdus bovienii, has the potential to provide much insight into the evolutionary events involved in the formation of this symbiotic relationship. Phylogenetic analysis of the subspecies of X. bovienii using outgroups such as X.nematophila and X.szentirmaii illustrated without question that the 31

PAGE 39

Xenorhabdus bovienii group is positioned as a sister species relative to the other species sampled. Then, after analyzing each gene independently, it seems that the 16s rRNA gene displayed the expected results in which very little diversity was visible amongst the strains of X.bovienii In these samples the bootstrapped values were all quite high, and were most likely due to the highly conservative nature of the 16s rRNA gene. As stated earlier, the relationships presented most likely artificially represent the strains of X.bovienii as more closely related than they may actually be. Of note, in this regard, is the identication by Farmer et al. (1983) of five separate isolates that were classed as X. luminescens. DNA-DNA hybridization showed these ve isolates to be closely related, but sufciently different from other Photorhabdus species such that they may in fact constitute a new species. To date, these are the only members of the Xenorhabdus/ Photorhabdus group to be isolated as saprophytes (i.e., with no nematode host), and the major property that allowed for their identification was that of bioluminescence (Farmer et al. 1983). Analysis of the recA gene also displayed a great deal of interrelatedness amongst the strains of X. bovienii with a few notable close clusters evident in X.bovienii of S. feltiae of Bodega Bay, S. oregonense of Bubbling Ponds, as well as X.bovienii of S. feltiae of Anatolia, and that of S. weiseri Overall, as compared to the other genes sequenced, this single gene probably gives the best estimate as to possible existence of subspecies or strains within X. bovienii because there is not a great deal of interallele recombination, but more variation inherent than what is presented on the 16s rRNA gene (Sergeant et al 2006). 32

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However, it is clearly most likely that the phylogenetic relationships of the three genes assembled contiguously provides the most balanced view, in regard to genes that are more or less conservative. Analysis of the serC sequences showed even more specialized clustering of the strains of X. bovienii which was as predicted due to the higher variability inherent in this gene. Clearly, from all phylogenetic trees, the X. bovienii are all more closely related to each other than to the outgroups: X. nematophila X. cabanillasi X. szemtirmaii and X. doucetiae. However, the affinity shown by some strains of X.bovienii to each other does appear to reveal certain trends in the adaptation to selection pressures, and allows for the postulation of several theories as to the formation of the nematode/symbiont relationship. One hypothetical possibility is that Xenorhabdus bovienii was transported around the world with the descendants of the ancestral Steinernema nematode. As the species of Steinernema spread and adapted to their environment, Xenorhabdus bovienii remained relatively unchanged. It would then follow that whatever variation that is present in the sequences was most likely the result of random mutation. However, in refute of this hypothesis is the observation of the close alignment of the X. bovienii strains of S. feltiae nematode srtains. If the theory was to have been supported, one might have expected to see an equal distribution of X.bovienii with any close alignment of strains absent any correlation to the identity of the Steinernema host. Another theory that also helps to explain the observed close relationships present between X. bovienii strains of the same nematode symbiont species is 33

PAGE 41

that a similar feature in the ancestor to X. bovienii strains was required to fully colonize the nematode vesicle. This type of novel adaptation is similar in nature to the independent origin of the toxic symbiont release adaptation in Photorhabdus (Doris et al. 1999). This scenario would imply access of aposymbiotic IJ to an undifferentiated Xenorhabdus ancestor, possibly already in hemolymph of an insect, prior to nematode infection. As it has been observed that aposymbiotic nematodes can kill some insects, this event may have occurred (Kim and Forst 2005). Then, after exposure to the Xenorhabdus ancestor, the colony that was able to best exploit the vesicle was able to propagate, and then differentiated accordingly. From a different approach, it is quite possible that a combination of the above named scenarios occurred. For instance, it may be that upon infection of a host insect by two Steinernema nematodes, such that one Steinernema is aposymbiotic and the other contains Xenorhabdus bovienii the second infective juvenile may take up the bacteria into its open vesicle. For example, this theory explains the close relationship (bootstrapped value of 97) of X. bovienii of S. weiseri to X.bovienii of S. feltiae from Florida observed in the triple gene analysis, and may have occurred such that once one strain of X.bovienii was inside the vesicle, the strain of X.bovienii adapted such that only the colonies that were best able to colonize and exploit the vesicle of the nematode were retained. In other words, the adaptation of the X.bovienii colony was stimulated by the opportunity to exploit the niche of the nematode vesicle. Furthermore, this idea is also supported by our preliminary experiment in which the Steinernema species that 34

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normally contain X. bovienii were able to be colonized by what composed the bacterial lawn on which the axenic Steinernema were raised, Xenorhabdus nematophila In this particular instance, it seems most likely that devoid of the preferred symbiont, some IJ were able to internalize X.nematophila into their vesicle (instead of allowing it to simply pass through the digestive system). Due to the unfortunate fact that we were unable to extract these bacteria from the individuals from which they were observed via microscope, we do not know what genotypic modifications may have been evident in these colonies. Nevertheless, this observation coupled with the MLST results, seem to project a complicated symbiotic relationship, that appears to favor a dynamic and changing relationship between nematode host and bacteria symbiont over one in which members of each pair is rigidly bound to each other. VI: Future Applications Because of what was determined to be the very closely related genotypes and nearly identical phenotypes of the Xenorhabdus bovienii strains studied, future studies could be carried out to determine the interchangeability of the X.bovienii symbiont of the Steinernema host with respect to nematode reproductive fitness. These studies would then have broad applications in regards to the use of Steinernema nematodes as entomopathogenic biocontrol agents. For instance, if Xenorhabdus bovienii was found to be virtually interchangeable between Steinernema symbionts, upon coinfection of an insect 35

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host, it would not seem likely that the commercially placed nematodes would become ineffective if exposed to a strain of X.bovienii from a native Steinernema Also of interest is the value of this mutualistic pair as a model to teach students about symbiosis. Because of the relative ease of rearing Steinernema this model would lend itself quite well to hands on classroom experiments for students of all ages. 36

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Appendix: Appendix A: Solution Recipes Axenizing Solution: 32.2 mL of 6% NaOCl 5.0 mL of 5M NaOH 80.0 mL distilled water Luria-Bertani broth: 10.0 g Tryptone 5.0 g yeast extract 10.0 g NaCl 800 ml distilled water 0.5 mL 4 M NaOH Lipid agar: 8 g nutrient broth 15 g bacto-agar 5.0 g yeast extract 890 mL of distilled water 10 mL .2 g/mL MgClx6H20 100 mL of Karo corn syrup mixture 4.0 mL corn oil M9 Buffer: 3.0 g KH2PO4 6.0 g Na2HPO4 5.0 g NaCl 1.0 mL MgSO4 (1N) 1.0 L distilled H2O NBTA agar: 1.00 L nutrient Agar 0.04 g Triphenyltetrazolium Chloride 0.025 g Bromothymol Blue PCR Prep 12.5 L ReadyMix Taq PCR Reaction Mix 1.0 L DNA 9.0 L Nanopure Water 1.5 L Forward Primer 1.5 L Reverse Primer 37

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Appendix B: ABI Sequencer Information Information retrieved from: http://www.gmi-inc.com/BioTechLab/ABI%20377.htm USE OF AN ABI 377 FOR SEQUENCING Introduction This SOP describes how to use an ABI 377 for sequencing. Materials 377 sequencing gel (SOP/QH58Z02/1999/0062) 1xTBE (SOP/QH58Z02/1999/0027) Sequencing Samples Method 1) Preparing the machine After the previous run is completed remove the heating plate and place handle side down on the bench [DO NOT PLACE FLAT SIDE DOWN AS THIS CAN RUIN THE HEATING PLATE]. Remove the used cassette and take it complete with the upper buffer chamber to the sink.Take off upper buffer chamber and wash with RO water before use. Take used gel out of the cassette and place in tray provided ensuring that there is tissue between the top of the gel already in the tray and the gel to be put in. This prevents scratching of the glass plates. Wash the cassette with RO water and dry before use. Discard buffer from the lower buffer chamber and wash with RO water. Collect a sequencing gel from the gel pouring room wash off excess acrylamide with warm water followed by deionized water and dry plates using white kimwipe, lint free tissues (red box). Take out the pouring comb and make sure there is no acrylamide left in-between the gel plates at the top. Clean between the plates with an old loading comb to remove any of the old acrylamide. 38

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Wash the well left by the sequencing comb with 1 x TBE and carefully insert the sharks tooth comb into the gel so that the tips of the teeth just penetrate the top of the gel. Do not push the comb into the gel too far otherwise it will be difficult to load the gel. If the comb has been pushed into the gel too far but is still loadable DO NOT remove the comb and try again as this will ruin the wells. Clip the gel into the ABI cassette ensuring the gel plates are flat in the cassette. 2) Preparing the computer Before each run, restart the Mac computer by going to the Special menu and selecting Restart This ensures that the RAM memory is not fragmented and will restart the collection software. Select ABI Collection Software, this nornmally comes up automatically when the computer is restarted. In the File menu select New then select Sequencing Run. In the Run window that appears make sure that the following settings are present. For dRhodamine and BigDye Terminators. PLATE CHECK: Plate Check A. PRE RUN: Seq PR 36E-2400 RUN: SEQ RUN 36E-2400. MATRIX: ABIX_BDT_ACCUGEL_DEC99 For the matrix setting the X refers to the machine so ABI A will have a matrix called ABIA_BDT_ACCUGEL_DEC99 [Note: On the Monaco ABI the pre run and run modules have the suffix chiller added because of the additional water bath installed] To change the number of lanes run select either 36 and full scan or 48 and XL scan. Plate Check Press plate check and then the scan window will appear and after approximately 1 min 4 coloured lines will appear. These should be flat and below 2048. If there are peaks in this trace this indicates that the plates are not clean and the cassette needs to be taken out of the machine and cleaned with kimwipes again. If the peak is not removed the corresponding lane may have to be missed out during loading. 39

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If the plate check is OK Clip on the buffer chamber and fill with 1 x TBE. Wait a minute to ensure there is no leakage of buffer down the front of the plate. This could cause arching of electrical current if left and severely damage the ABI. Next clip on the heating plate and fill the lower buffer chamber with 1 x TBE. Rinse out the wells thoroughly to ensure that there are no air bubbles or excess acrylamide present. Pre Run Select pre-run. Then go to Window: Status and check the run is for 1 hour and that the gel gets up to temperature 51 o C. You should allow at least 10 minutes for the pre-run. While the gel is pre running prepare the sample sheet by selecting File New -Sequencing Sample sheet and typing in the appropriate lanes. Save the sample sheet. Prior to loading your samples and performing a run, add 3 l loading dye for ABI377. Make sure the pellet, which is not visible, is thoroughly resuspended either by pipetting the loading dye up and down the tube a few times or by vortexing gently for 10-15 seconds to ensure that your sample is resuspended in the loading dye. Centrifuge samples briefly. Denature the samples for 2 mins @ 95 o C and immediately store on ice until ready to load (this is important as DNA will re-anneal if not immediately placed on ice). Pause the Pre Run and wash out the wells once more. Load sample sheet into the run window [ the machine will not be able to start a run without a sample sheet] Load 1-2 l per lane, loading every other lane and then cancel the pausepress continue and run the samples in for 3 minutes. Pause the gel again and wash out the wells and load the well not loaded in the first round. Run the samples in for a further 3 minutes and then cancel the Pre Run. Start the Run and then a window will appear asking to name the run. After the run has finished the run folder can be transferred to another computer for further analysis 40

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[The reason for loading every other lane is that the machine can recognize the individual lanes and track the gel in the analysis program. If every lane is loaded at the same time with a sharks tooth comb the machine will only see one broad lane] PRODUCTION OF SEQUENCING MATRIX USING DATA UTILITY Introduction This S.O.P. describes the production of a sequencing matrix on both 373 and 377 ABI machines. Materials 373 or 377 gel (see SOP/HH5TA02/1998/0023 and SOP/HH5TA02/1998/0010) BigDye dRhodamine matrix dye set Deionised Formamide Method Set up the gel in ABI machine as SOP/QH58Z02/1999/0066, SOP/ QH58Z02/1999/0067 or SOP/HH5TA02/1998/0012, for ABI373, ABI 373-XL and ABI 377 respectively. Combine 2.5 l of each matrix standard with 2.5 l deionised formamide, in duplicate. This will give you 8 samples, 2 of each dye type. Heat each sample to 96 o C for 2 minutes to denature before loading. Create a sample sheet to assign sample (see SOP/QH58Z02/1999/0064) but do not assign a matrix to the samples and do not allow autoanalysis. Load the Matrix Standards in duplicate, 2.5 l of the matrix standards, i.e. load 8 lanes 2 dR110, 2 dR6G, 2 dROX and 2 dTAMRA, leaving an empty lane between each standard. Select the appropriate filter-set, and start the run. 377 ? Run modules containing filter set E, 373 ? filter-set A. After the gel has run Open up Sequence Analysis software 41

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Go to File then Open then Collection gel and open the gel you wish to produce a matrix from. Track the lanes containing the Matrix standards this is done by selecting a lane and going to Gel on the menu and selecting Mark lane used then move the tracking lanes so that they follow exactly the lane of that well. When this has been done for all the lanes used, go to Gel and Extract the lanes. From within the Utilities folder of the Sequence analysis software, open the DataUtility program. In the Utilities menu, choose Make Matrix. In the dialogue box that appears make sure that the Dye Primer matrix button is selected. In the next box that appears press the "C" nucleotide button and select the appropriate sample file for that nucleotide according to the table below. For each sample fill in a start point, where analysis of each sample will start, start with the default value of 2000 and for the points choose 1500, for the number of data points to analyse. Click New File and name the new matrix e.g. BigDye ABI10 Oct99. Then press OK. The computer will then try and make the matrix, if it is unable to do this an error box will appear telling at which point it has failed. One solution if it has failed to make a matrix is to increase the amount of data points to analyse. If the matrix has been successful go back to the Utilities menu, choose Make Matrix This time make sure that the Taq Terminator button has been selected before choosing the sample files. After selecting the sample files, select Update the File and choose the correct matrix name e.g. BigDye ABI10 Oct99 to update. Repeat again as above but this time making sure that the T7 Terminator Matrix button has been selected. The matrix is automatically saved in the ABI folder. To check the matrix, go to the utilities menu and select copy matrix. Under Source, select Instrument file and choose the matrix you wish to check. 42

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