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Molecular analysis of defective interfering RNAs associated with cucumber necrosis virus infections Finnen, Renée Louise 1996

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M O L E C U L A R ANALYSIS OF DEFECTIVE INTERFERING RNAS ASSOCIATED WITH CUCUMBER NECROSIS VIRUS INFECTIONS by RENEE LOUISE FINNEN B. Sc. (Agr.), the University of Guelph, 1988 M . Sc., the University of British Columbia, 1991 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY i n THE FACULTY OF GRADUATE STUDIES (Department of Microbiology and Immunology) We accept this thesis as conforming to the reauired standard THE UNIVERSITY OF BRITISH COLUMBIA March, 1996 © Renee Louise Finnen, 1996 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 of Nlicfco&fOtoG,^ /MMUAJQCOchy The University of British Columbia Vancouver, Canada DE-6 (2/88) Abstract In this thesis, the molecular biology of defective interfering (DI) RNAs associated with cucumber necrosis virus (CNV), a simple, well characterized plant R N A virus, was examined to gain insight into C N V RNA replication. Seven different cDNA clones of DI RNAs associated with either a laboratory strain of CNV or nucleic acid extracts from serially-passaged CNV infections (de now-generated DI RNAs) were constructed using reverse transcription followed by the polymerase chain reaction (RT-PCR). The sequence of each clone was determined and RNA transcripts generated by T7 RNA polymerase were assessed for biological activity by coinfecting with CNV. This analysis demonstrated that three or four large sequence blocks of the C N V genome are retained in C N V DI RNAs. These sequence blocks therefore likely represent essential c/s-acting elements involved in viral RNA replication. The presence of DI RNAs in CNV infected plants interfered with the development of the severe necrosis typical of CNV infections. Total nucleic acid extracts from coinfections were also found to contain RNAs which were twice the size of the DI RNA used in the coinfection. Using RT-PCR analyses and direct sequencing of RNA templates, these RNAs were established to be linear, head-to-tail repeats of DI RNA sequence, the first described in association with tombusvirus DI RNAs. Further characterization revealed that DI RNA dimers are likely generated from monomers by RNA recombination. Individual DI RNAs were found to vary in the amount of dimer which they accumulate during coinfection. Dimer accumulation was examined by coinfecting a series of chimeric and mutated DI RNAs constructed from a DI RNA which accumulated primarily as monomer and one which accumulated primarily as dimer. These analyses identified two distinct tracts of sequence in the 3' terminal region which correlate with increased dimer accumulation. The results of these investigations are discussed in terms of the insights they provide into CNV replication and recombination. A model for the generation of DI RNA dimers is also proposed. Table of Contents Abstract i i Table of Contents • i i i List of Tables v i List of Figures v i i List of Abbreviations ix Acknowledgments xi i i Dedication xiv Introduction 1 Definition of defective interfering viruses 1 Discovery and incidence of defective interfering viruses 1 Plant virus defective interfering RNAs 3 Implications of defective interfering viruses in animal and plant infections 7 Generation and evolution of defective interfering RNAs 9 Replication of positive-strand plant R N A viruses 16 Cucumber necrosis virus 20 Epidemiology .20 Classification 21 Infection cycle 22 Vector transmission 22 Viral R N A replication 25 Genomic organization and expression 26 Function of proteins 28 Movement 31 Rationale of study 33 Materials and Methods 35 Materials 35 i i i Sources of C N V DI RNAs 36 Oligonucleotides used in this study 36 Plasmids used in this study 38 Construction of cDNA clones of C N V DI RNAs 40 Construction of a head to tail repeat of DI R N A cDNA 42 Construction of chimeric DI RNAs 42 Construction of short stem and long stem mutants 43 D N A sequencing 46 Direct sequencing of RT-PCR products and restriction fragments 47 R N A sequencing 48 RT-PCR of dimer junctions 49 In vitro transcriptions 50 Coinoculation of Nicotiana clevelandii 50 Extraction of total nucleic acid from Nicotiana clevelandii leaves 51 Northern blot analysis 51 Chapter 1 - Sequence and biological properties of cloned C N V DI RNAs 53 Results 53 Sources of DI RNAs for cloning 53 Cloning and sequence analysis of DI RNAs 54 Biological activity of cloned DI RNAs 61 Involvement of coat protein in DI R N A accumulation 65 Discussion. 67 Chapter 2 - Identification and characterization of C N V DI R N A dimers 72 Results 72 Analysis of overall structure and size of dimer-sized RNAs 72 Analysis of cDNA sequence generated from dimer RNAs 75 Analysis of dimer junction sequences 78 Biological activity of synthetic dimer transcripts 80 i v Biological activity of monomers with 3' terminal extensions 81 Analysis of dimer accumulation in chimeric and mutated DI RNAs 84 Discussion 95 General Discussion 106 References 118 v List of Tables Table 1. Properties of some subviral agents 2 Table 2. Definitive members of the Tombusvirus genus 23 Table 3. Synthetic oligonucleotides used in this study 37 Table 4. Plasmid constructs unique to this study 39 v i List of Figures Figure 1. Generation of DI RNAs by nonhomologous R N A recombinationl3 Figure 2. Infection cycle of C N V 24 Figure 3. Organization and expression of the C N V genome 27 Figure 4. Construction of complementary D N A clones of C N V DI RNAs ...41 Figure 5. Strategy for construction of pDISS9 44 Figure 6. C N V DI RNAs generated de novo from wild-type inoculum 56 Figure 7. Strategy used for sequencing C N V DI R N A cDNA clones 57 Figure 8. Comparison of cloned C N V DI RNA sequences with C N V 58 Figure 9. Structure of cloned DI RNAs relative to the C N V genome 60 Figure 10. Symptom attenuation caused by C N V DI RNAs in Nicotiana clevelandii 63 Figure 11. Northern blot analysis of coinfected Nicotiana clevelandii 64 Figure 12. Analysis of DI R N A accumulation in coinfections using wild-type C N V or CP(-) synthetic transcript as helper 66 Figure 13. Northern blot analysis of dimer-sized RNAs generated during coinfections in Nicotiana clevelandii 74 Figure 14. Analysis of cDNA generated from dimeric- and monomeric-lengthDIRNA 76 Figure 15. Sequence analysis of dimer R N A junctions 79 Figure 16. Coinfection of Nicotiana clevelandii with dilutions of monomeric and dimeric-length DI R N A 42 transcripts 82 Figure 17. Analysis of transcripts from Sspl-linearized pDI42 83 Figure 18. Comparison of complementary D N A sequences of DI R N A 9 and DI R N A 21 85 Figure 19. Chimeric and mutated C N V DI RNAs 86 Figure 20. Dimer accumulation in chimeric and mutated C N V DI RNAs ....88 Figure 21. Rolling circle R N A replication 97 v i i Figure 22. Generation of turnip crinkle virus satellite R N A and DI R N A multimers by R N A recombination 98 Figure 23. Predicted stem loop structures of various DI RNAs at junctions between regions II and Ilia 105 Figure 24. Models for the generation of C N V DI R N A dimers 108 Figure 25. Some possible interactions of the 3' terminus of (+) DI R N A with other regions of DI R N A I l l Figure 26. Organization and comparison of repeated sequences found at the junction of C N V DI R N A dimers 113 v i i i Lists of Abbreviations a alpha A Angstrom unit A adenosine in the context of nucleotide sequence A M C V artichoke mottle crinkle virus A T P adenosine-5'-triphosphate P beta B B M V broad bean mottle virus BE borate/EDTA B M V brome mosaic virus B V D V bovine viral diarrhea virus B N Y V V beet necrotic yellow vein virus C cytidine in the context of nucleotide sequence °C degrees Celsius c D N A complementary D N A C i Curie C M V cucumber mosaic virus C N V cucumber necrosis virus C N V - L c laboratory culture of C N V CP(-) a C N V mutant which lacks the viral coat protein gene C P M V cowpea mosaic virus CyRSV cymbidium ringspot virus D aspartic acid dATP deoxyadenosine triphosphate dCTP deoxycytidine triphosphate ddATP dideoxyadenosine triphosphate ddCTP dideoxycytidine triphosphate i x ddGTP dideoxyguanosine triphosphate ddTTP dideoxythymidine triphosphate dGTP deoxyguanosine triphosphate DI defective interfering DIs defective interfering viruses D N A deoxyribonucleic acid dpi days post infection ds double-stranded dTTP deoxythymidine triphosphate DTT dithiothreitol EDTA ethylenediaminetetraacetic acid g gram(s) G guanosine in the context of a nucleotide sequence; glycine in the context of a protein sequence hpi hours post infection kDa kilodalton(s) 1 liter(s) |i. micro m milli M molar min minute(s) M M L V Moloney murine leukemia virus moi multiplicity of infection mol moles m R N A messenger RNA MVBs multivesicular membranous bodies n nano NTP nucleoside triphosphate x oligo(s) oligonucleotide(s) ORF open reading frame p pico PAGE polyacrylamide gel electrophoresis PCR polymerase chain reaction pK2/M5 full length infectious cDNA clone of C N V RdRp(s) RNA-dependent R N A polymerase(s) RNA(s) ribonucleic acid(s) r R N A ribosomal R N A RT reverse transcription S Svedberg unit SDS sodium dodecyl sulfate sec seconds ss single-stranded S Y N V sonchus yellow net virus T thymidine in the context of a nucleotide sequence T A E Tris acetate EDTA TBE Tris borate EDTA TBSV tomato bushy stunt virus T C V turnip crinkle virus Tris hydroxymethyl amino methane T M V tobacco mosaic virus t R N A transfer R N A T S W V tomato spotted wilt virus U uridine in the context of nucleotide sequence; units when referring to amount of an enzyme W T wild-type W T V wound tumor virus x i positive polarity (in reference to RNA) negative polarity (in reference to RNA) x i i Acknowledgments Financial assistance of this research in its early stages was provided by the British Columbia Blueberry Co-operative Association and in its latter stages by the Medical Research Council of Canada and the University of British Columbia graduate fellowship program. I wish to acknowledge the following people for their individual contributions which assisted me in completing my Ph.D. degree. My initial decision to pursue this degree was heartily encouraged by my M.Sc. supervisor, Dr. R. E. W. Hancock and also by Dr. F. Tufaro whose enthusiasm for the subject of virology was (and still is) "contagious". M y Ph.D. supervisor, Dr. D. M . Rochon, through her zealous, interactive approach to overseeing this research and the communication of its findings, allowed me to develop both personally and professionally. The members of my supervisory committee, Drs. H . Sanfagon, G. B. Spiegelman, and F. Tufaro also provided sound guidance over the direction of this research and the contents of this thesis. A l l members of the Rochon lab both past and present provided a mutually supportive (and musically diverse) work environment. Helpful technical advice was provided by M . E. Rott, J. C. Johnston, L. Lee and A. Gilchrist during the early stages which enabled me to get this research project up and running; expert technical assistance was provided by Dr. H . G. Damude during the final stages of this research. C. J. Riviere, A . Wieczorek, Dr. T. L. Sit, and J. C. Johnston acted as invaluable resource persons throughout most of this research. The greenhouse staff of the Pacific Agriculture Research Center provided a steady, dependable supply of plants used in this research. I am grateful for the often unsung efforts of the administrative staffs of the Department of Microbiology and Immunology, the Pacific Agriculture Research Center, and the Department of Plant Science and especially grateful to P. Gontier and D. G. Huchelega for assistance "above and beyond the call of duty". Finally, sincere thanks go out to many friends and my family members for providing a reliable emotional support network. This thesis is dedicated to Bruce William Banfield for his extraordinary patience and unwavering belief in my abilities and for ensuring that my sense of humor remained firmly intact throughout it all. x i v 1 INTRODUCTION Definition of defective interfering particles The term defective interfering (DI) was first proposed in 1970 in response to the growing number of defective animal viruses reported to interfere with the growth of non-defective standard virus (Huang and Baltimore, 1970). DI particles were originally defined as having the following four properties: "(1) they contain normal viral structural protein; (2) they contain a part of the viral genome; (3) they can reproduce in the presence of helper virus; (4) they interfere specifically with the intracellular replication of non-defective homologous virus" (Huang and Baltimore, 1970). The first two properties together distinguish defective interfering viruses (Dis), in particular DI RNAs, from most other known subviral agents (Table 1). The third property relates to the fact that DIs are unable to replicate in the absence of helper virus. The term helper virus refers to any virus that can replicate the DIs i n question and does not necessarily need to be the virus from which the DIs were originally generated. The last property on this list relates to the interfering properties of DIs. In a viral infection, the presence of DIs results in reduced yields of helper virus. This last property has sparked much interest in utilizing DIs as a means of controlling viral infections in both animals and plants and was responsible for the initial discovery of DIs. Discovery and incidence of defective interfering particles The concept of interference by incomplete virus particles originated over 40 Table 1. Properties of some subviral agents. 2 Subviral agent Shares sequence homology with helper virus Uses helper virus structural proteins Relies on a Interferes with helper virus helper virus for replication accumulation satellite virus no no yes yes/no^ satellite RNA no yes yes yes/nol viroid RNA N / A 2 N / A no N / A virusoid RNA no yes yes yes/no-'-defective RNA yes yes yes no DI RNA yes yes° yes yes ^Effects of satellite viruses, satellite RNAs and virusoid RNAs on helper virus can vary (see Matthews, 1991; Roossinck et al, 1992). 2 N / A = not applicable. ^Some exceptions do exist (see Rochon et al, 1994). years ago from experiments with influenza virus preparations. The usual method of propagating influenza virus at this time was to dilute stock virus 1000-fold and then use the diluted virus to inoculate embryonated eggs. Virus propagated in this manner was found to be highly infectious. However, when undiluted virus was used as the inoculum, the resultant progeny virus was found to be less infectious and had a heterogeneous sedimentation rate indicative of a heterogeneous population of viruses (Gard and von Magnus, 1947; von Magnus, 1947). This phenomenon was hypothesized to be a result of incomplete virus particles in the progeny virus which interfered with the growth of influenza virus (von Magnus, 1954). These early experiments highlight the importance of using dilute virus stocks when propagating virus in order to avoid Dis in virus stocks and, moreover, the ability to use serial passaging of undiluted virus as a means of amplifying DIs for study. In these early studies, heterogeneous sedimentation rates were interpreted as evidence of incomplete virus particles. The first physical demonstration of incomplete virus particles came much later in 1964. Undiluted passages of vesicular stomatitis virus were demonstrated by electron microscopy to contain particles which were shorter than those of the standard virus (Hackett, 1964). 3 By 1970 defective viruses with interfering properties had been reported i n association with undiluted passages of 10 different animal viruses, including both D N A and R N A viruses, prompting the establishment of the term DI (Huang and Baltimore, 1970). DIs are now known to be virtually ubiquitous amongst the animal viruses, having been reported in association with every R N A virus in which their existence has been investigated as well as with most D N A viruses (Perrault, 1981; Lai, 1992). In addition, DIs have been described for R N A and D N A bacteriophage (Griffith and Romberg, 1974; Enea and Zinder, 1975; Mills et al, 1975; Schaffner et al, 1977; Chen and Ray, 1978; Ravetch et al, 1979), the double-stranded R N A killer virus of yeast (Ridley and Wickner, 1983; Theile et al, 1984), plant D N A viruses (Stanley and Townsend, 1985) and plant R N A viruses (see next section). Plant virus defective interfering RNAs As early as 1983, Dl-like particles were reported in association with passaged potato yellow dwarf virus (a member of the Rhabdoviridae) infections (Adams et al, 1983) and, later, deletion mutants of wound tumor phytoreovirus (WTV) were reported to interfere with the accumulation of standard virus (Nuss and Summers, 1984) . The first definitive description of a DI R N A in association with a plant R N A virus, however, did not occur until 1987. The molecular organization of a small R N A previously described in association with tomato bushy stunt tombusvirus (TBSV) infections (Hillman et al, 1985) was determined to be a linear deletion mutant of the helper virus (Hillman et al, 1987). Coinfections of this R N A and helper virus were also demonstrated to attenuate normal symptom development and resulted in a decreased recovery of progeny virus. These characteristics established this RNA as an authentic DI RNA. Soonafter, DI RNAs were described in association with other plant R N A viruses. At the commencement of the studies in this thesis, DI RNAs had been definitively described in association with TBSV (Hillman et al, 1987), WTV (Anzola et al, 1987), sonchus yellow net virus (SYNV, also a member of the Rhabdoviridae) (Ismail and Milner, 1988), cymbidium ringspot tombusvirus (CyRSV; Burgyan et al, 1989), turnip crinkle carmovirus (TCV; L i et al, 1989), and mutants of cucumber necrosis tombusvirus (CNV) which lacked expression of the 20 kDa protein (Rochon, 1991). By the completion of these studies, DI RNAs had also been described in association with tomato spotted wilt tospovirus (TSWV; Resende et al, 1992) and broad bean mottle bromovirus (BBMV; Romero et al, 1993). There are many examples in the literature of defective (D) RNAs associated with plant R N A viruses, which, unlike the RNAs described in the aforementioned studies, have not been demonstrated to interfere with the accumulation of helper virus. D R N A s have been described in association with clover yellow mosaic potexvirus (White et al, 1991), TSWV (Resende et al, 1991), three furoviruses: beet necrotic yellow vein virus (BNYVV; Bouzoubaa et al, 1991); peanut clump virus (Manohar et al, 1993); and soil-borne wheat mosaic virus (Chen et al., 1994), citrus tristeza closterovirus (Mawassi et al, 1995), and cucumber mosaic cucumovirus (CMV; Graves and Roossinck, 1995). Having now been reported with at least 10 divergent plant R N A virus genera, D and DI RNAs are emerging as a common theme in plant virology. It is very likely that new reports of D and DI RNAs in association with plant viruses wil l continue to surface. In systemic plant hosts, where the viral infection can spread from the original inoculation site to infect the whole plant, the effect of DI RNAs on symptom development can be quite dramatic. In the case of all tombusvirus DI RNAs, S Y N V DI RNAs and TSWV DI RNAs symptoms are attenuated in comparison to those produced by standard virus. Transgenic plants expressing CyRSV DI RNAs also show attenuated symptoms upon challenge with helper virus (Kollar et al, 1993). In addition, a positive correlation exists between the level of symptom attenuation 5 and the abundance of C N V DI RNAs in the infection (Chang et al, 1995). Attenuation of viral symptoms seems predictable in light of the decrease in virus yield caused by DI RNAs. There are, however, two notable exceptions. The presence of TCV DI RNAs and BBMV DI RNAs during infection leads to more severe symptoms. These exceptions illustrate that the observed biological effects of any DI is the result of a complex three-way interaction between host, virus and DI and, therefore, these effects are often unpredictable. Lower virus levels resulting from competition for replicase is generally accepted as the predominant factor contributing to the symptom attenuation observed in the presence of Dis (Huang and Manders, 1972; Perrault and Holland, 1972; Giachetti and Holland, 1989; Roux et al, 1991). Although both the yield of progeny virus and the accumulation of genomic R N A have been repeatedly demonstrated to be decreased in the presence of DI RNAs (for specific examples, see Hillman et al, 1987; L i et al, 1989; Burgyan et al, 1991; Jones et al, 1990; Romero et al, 1993; Scholthof et al, 1995c), the molecular basis for the modification of disease progression by plant virus DI RNAs remains largely unexplored. Only one study to date has attempted to correlate viral protein levels with symptom development i n the presence of DI RNAs (Scholthof et al, 1995c). In this study, the accumulation of RNAs (both genomic and subgenomic RNAs) and viral proteins in plants coinfected with TBSV and TBSV DI RNAs was monitored. Reduced levels of both genomic R N A and subgenomic RNAs were found in the presence of DI RNAs. Reduced levels of all viral proteins were also observed. The proteins which were most affected were those implicated in viral cell-to-cell movement and in induction of necrotic symptoms (p22 and pl9, respectively). These authors suggest that a decreased rate of viral cell-to-cell movement and a reduced general tissue necrosis occur in the presence of DI RNAs which could contribute to the observed symptom attenuation. 6 It has long been appreciated with bacterial and animal virus systems that DI RNAs can be powerful tools for elucidating cz's-acting elements involved in viral R N A accumulation (Mills et al, 1967; Levis et al., 1986). The more established plant virus DI RNAs have also been utilized in this capacity. Two general approaches have been employed to define cz's-acting elements contained in plant virus DI RNAs: introduction of deletions and/or insertions into cloned DI RNAs followed by assessment of the ability of the altered DI RNAs to accumulate (Li and Simon, 1991; Burgyan et al, 1992; Zhang and Simon, 1994; Chang et al, 1995; Havelda et al, 1995); and sequence analysis of DI RNAs selected for increased ability to accumulate using competition assays (White and Morris, 1994a; White and Morris, 1994c). Details of these studies and the cz's-acting elements identified by them are discussed where appropriate in subsequent chapters of this thesis. Perhaps the most practical application of plant virus DI R N A research is the development of transgenic plants expressing DI RNAs as a means of controlling viral infection. DI RNAs constitutively produced in transgenic plants should be amplified only upon challenge with appropriate helper virus. The presence of DI RNAs at the onset of viral infection should then prevent the development of disease. This idea has been around since the initial discovery of DI RNAs i n association with plant viruses and has shown initial promise in at least one experimental host. Nicotiana benthamiana plants transformed with CyRSV DI R N A cDNA sequence were found to constitutively express DI RNA-related transcripts and were protected from subsequent challenge with helper virus (Kollar et al, 1993). This disease resistance strategy provides an alternative to strategies involving the expression of viral proteins (see Lomonossoff, 1995). It may be possible to extend this strategy to control plant viruses not known to be associated with DI RNAs by utilizing artificially constructed DI RNAs. 7 Implications of defective interfering particles in animal and plant infections The discovery and characterization of Dis have largely taken place in a laboratory setting with model systems. It is well established in both animal and plant model systems that Dis can influence disease progression. As early as 1950, it was appreciated that inoculation of animals with virus stocks containing Dis resulted in fewer fatalities than inoculations with virus stocks devoid of Dis (Bernkopf, 1950). The protective effect of Dis in animals has been reported in a number of laboratory studies (for examples see Doyle and Holland, 1973; Welsh et al, 1977; Dimmock and Kennedy, 1978; Rabinowitz and Huprikar, 1979; Fultz et al., 1982; Morgan and Dimmock, 1992; Morgan et al, 1993; Noble and Dimmock, 1994). The effect of Dis on symptom development in experimental plant hosts is a very obvious visual example of the biological effects of Dis. Transgenic plants expressing Dis have also been reported to be protected from subsequent viral challenge (Stanley et al, 1990; Kollar et al, 1993). It has been observed that Dis can facilitate the establishment and maintenance of persistent infections of a wide variety of animal viruses in cultured cell lines (for examples see Roux and Holland, 1979; De and Nayak, 1980; Dauenhauer et al, 1982; Moscona, 1991). Persistent viral infections in either animals or plants can also be established with coinoculations of Dis and helper virus (Barrett and Dimmock, 1986; Morris and Knorr, 1990). The ability of a virus to generate Dis spontaneously is generally accepted and has been definitively demonstrated for several plant viruses using an intact plant host (a model more analogous to the natural situation; see L i et al, 1989; Burgyan et al, 1991; Knorr et al, 1991; Rochon, 1991). On the basis of these numerous experimental observations, it is widely believed that Dis do exist i n natural disease states of both animals and plants and can influence natural disease progression, particularly in the case of chronic or persistent diseases and recurrent infections. 8 A large body of information documenting attempts to demonstrate the presence and involvement of defective viruses or Dis in natural disease states of animals and humans has accumulated. Despite this, few clear examples of a l ink between defective viruses or Dis and a particular natural disease state exist. This is due to the technical difficulties in obtaining suitable amounts of tissue specimens for the isolation of Dis and the complicating effects of the immune response (Barrett and Dimmock, 1986; Roux et al, 1991). New technologies such as the polymerase chain reaction have made it easier to detect Dis in tissue samples taken from natural infections. However, even if Dis can be repeatedly demonstrated to be present in clinical material, there is still the challenge of proving or disproving that their existence has a specific biological role. In two well-studied natural diseases, the progressively fatal subacute sclerosing panencephalitis in humans and a fatal immunodeficiency disease in both cats and mice, the involvement of defective measles viruses and defective retroviruses, respectively, have been implicated (Cattaneo et al, 1988; Overbraugh et al, 1988; Aziz et al, 1989). These defective viruses, however, do not appear to have interfering properties. The sole report of Dis causing a natural disease state i n animals came only recently from studies on a fatal mucosal disease of cattle known to be caused by bovine viral diarrhea virus (BVDV) (Tautz et al, 1994). Persistent infections by a noncytopathic strain of BVDV in cattle were found to be associated with a DI R N A which was able to exert a cytopathic effect, resulting in an outbreak of fatal disease. This particular DI R N A had an internal deletion of 4.3 kb and encoded one long open reading frame (ORF). The p80 protein product expressed from this ORF, a known marker protein of cytopathic strains of BVDV that is not expressed in noncytopathic strains (Corapi et al, 1988), was responsible for the observed cytopathic effect. The incidence of Dis in natural plant virus infections or disease states has not yet been investigated, probably owing to their more recent discovery. These types of 9 investigations are certainly warranted in light of the fact that transgenic plants expressing DIs have shown initial promise as a means of controlling plant virus infections (Stanley et al, 1990; Kollar et al, 1993). The first report of a D R N A i n association with a natural disease of plants came only very recently (citrus tristeza closterovirus infections in Citrus macrophylla; Mawassi et al, 1995). DIs have not yet been implicated in natural diseases of plants. Plant viruses which are known to be associated with DIs in laboratory settings would be prime candidates to search for the occurrence of naturally occurring DIs. Apart from their role in influencing disease progression, DIs represent destabilizing forces in virus infections that can exert a selective pressure on the virus, leading to the evolution of viruses which are resistant to the interfering effect. This phenomenon has been documented in the case of several viruses (for examples see Kawai and Matsumoto, 1977; Horodyski and Holland, 1980; Jacobsen and Pfau, 1980; Enea and Zinder, 1982). Dl-resistant viruses themselves then generate new DIs and the cycle of selection begins again (DePolo et al, 1987; Bangham and Kirkwood, 1993). Consequently there exists, at least in experimental systems, a constantly coevolving population of virus and DIs. This has important implications for vaccine production and maintenance of genetically-defined virus stocks and, moreover, could potentially be a driving force for the generation of new viruses in nature. The emergence of Dl-resistant viruses is a significant possibility to bear in mind when developing transgenic plants or animals expressing DIs as a means of viral disease control. Generation and evolution of DI RNAs In many different viral systems, the generation of DI RNAs from viral genomic R N A and the subsequent evolution of DI RNAs have been suggested to occur via RNA recombination (Lazzarini et al, 1981; Perrault, 1981; Kuge et al, 1986; 10 Makino et al, 1988; Cascone et al, 1990; Furuya et al, 1993; L i and Ball, 1993; Romero et al, 1993; White and Morris, 1994a). Evidence of the ability of single-stranded (ss) R N A viruses to undergo genetic recombination was first described for poliovirus and related picornaviruses in the early 1960's (Hirst, 1962; Ledinko, 1963; Pringle, 1965). Since then, R N A recombination has been demonstrated for many different ss R N A viruses (reviewed in Lai, 1992; Bujarski et al, 1994; Simon and Bujarski, 1994) and is now appreciated as part of their general biology. Two general mechanisms can be proposed to account for the joining of non-contiguous segments of either R N A or D N A ; breakage-and-rejoining or copy-choice. Mechanisms of the first type are involved in D N A recombination (Meselson and Weigle, 1961), in cis and trans-splicing of mRNAs (Sharpe, 1987; Blumenthal and Thomas, 1988) and in RNAs which can undergo self-processing (Symons, 1992). A l l of the aforementioned examples are post-replication mechanisms. In contrast, copy-choice mechanisms are mediated by the process of replication. Copy-choice mechanisms are involved in rare types of D N A recombination (Brunier et al, 1988) and in R N A recombination (Kirkegaard and Baltimore, 1986; Lai, 1992; Bujarski et al, 1994; Simon and Bujarski, 1994). Although considerable evidence exists i n support of a copy-choice mechanism of R N A recombination (see below), breakage-and-rejoining has not been unequivocally ruled out. The copy-choice mechanism was proposed originally as a possible mechanism involved in R N A recombination by Cooper, (1974). Copy-choice requires several sequential events to occur: (1) the replication complex (viral R N A dependent R N A polymerase (RdRp) plus other accessory proteins) pauses during R N A synthesis; (2) this complex, along with the nascent R N A strand, switches to another R N A template or a new site on the same R N A template; (3) R N A synthesis continues. The first and still most direct evidence for the copy-choice mechanism came from studies on R N A recombination in poliovirus (Kirkegaard and Baltimore, 1986). As part of this study, the frequency of recombination between a wild-type (WT) 11 guanidine-sensitive virus and a temperature-sensitive guanidine-resistant virus (3NC-202guaR) was assessed under conditions which would inhibit the replication of each of these viruses independently (presence of guanidine in the case of W T virus and 39°C incubation temperatures in the case of 3NC-202guaR virus). The results of this study established that R N A synthesis is required for R N A recombination, and further, that in the poliovirus system, R N A recombination occurs during the synthesis of negative polarity [(-)] RNA. Other evidence i n support of the involvement of R N A synthesis in R N A recombination comes from the observation that nucleotides which are not encoded by the template R N A (non-templated nucleotides) are observed at the junctions between recombined sequences (Cascone et al, 1990; Carpenter et al, 1991a; Carpenter et al, 1991b; Nagy and Bujarski, 1993; White and Morris, 1994a). In addition, recombination in the TCV system appears to be directed by replicase recognition signals (Cascone et al, 1990; Cascone et al, 1993; Simon and Bujarski, 1994). Replication conditions and mutations in the gene encoding the replicase also alter recombination in brome mosaic virus (BMV; Nagy and Bujarski, 1993; Simon and Bujarski, 1994; Nagy et al, 1995). Copy-choice also requires that the replication complex pause during synthesis. Pausing during RNA synthesis has been shown to be associated with both RNA-dependent and DNA-dependent R N A polymerases (Mills et al, 1978; Kassavetis and Chamberlin, 1981; Baric et al, 1987; Landick et al, 1987). R N A recombination events are usually categorized as either homologous or nonhomologous (Lai, 1992). In homologous recombination, the RNAs involved share extensive sequence homology and crossovers occur at homologous sites on the two RNAs. Crossovers within this homologous site can be precise, leading to a recombinant R N A with the exact sequence and organization of the parental R N A s (e.g. maintenance of an open reading frame) or imprecise, leading to duplications or deletions in the recombinant R N A relative the parental RNAs. Parental RNAs are often termed donor and acceptor RNAs with the donor R N A being the template 1 2 from which R N A synthesis commences and the acceptor R N A being the template to which the replicase complex switches. Homologous recombination is thought to be one mechanism responsible for strain variation amongst R N A viruses and also a means by which errors of R N A replication can be repaired (Simon and Bujarski, 1994). Nonhomologous recombination involves parental RNAs which do not share appreciable sequence homology, allowing viruses to acquire sequences from other unrelated R N A viruses (Lai, 1992; Simon and Bujarski, 1994), from transgenically-expressed viral RNAs (Greene and Allison, 1994) and even cellular R N A sequences (Khatchkian et ah, 1989; Meyers et ah, 1991; Weiss and Schlesinger, 1991). It is therefore appreciated as a major driving force in the evolution of R N A viruses. Nonhomologous recombination is also believed to be responsible for the generation of different types of DI RNAs (Figure 1). The mechanism of nonhomologous R N A recombination has been most extensively studied in two plant virus systems, BMV and TCV. Two different renditions of the general copy-choice mechanism have emerged from these studies. BMV was the first R N A plant virus for which genetic recombination was demonstrated experimentally (Bujarski and Kaesberg, 1986). Analysis of the crossover sites of nonhomologous recombinants of BMV revealed that the recombining RNAs had the potential to form double-stranded heteroduplexes suggesting that heteroduplex formation might mediate nonhomologous recombination (Bujarski and Dzianott, 1991). This was experimentally verified using a donor R N A which carried sequence complementary to an acceptor R N A (Nagy and Bujarski, 1993). Mutations in the helicase domain of the BMV replicase have also been demonstrated to influence recombination (Nagy et al, 1995). Apart from further confirming that replication and recombination are interrelated, these results suggest that the inability of the replicase complex to unwind heteroduplexes may cause R N A synthesis to pause, thereby providing an opportunity for a switch to a less structured region. On the basis of these studies, a processive, heteroduplex-13 5'- 3' B internal deletion 6 OF 7 7 C i i i i i copy-back Figure 1. Generation of DI RNAs by nonhomologous R N A recombination. Thin lines represent viral R N A template; bold lines represent DI R N A ; dashed line indicates deleted sequence; hatched ball represents replication complex. A . Generation of internal deletions. This mechanism can occur intermolecularly (detachment-reinitiation) or intramolecularly (processive looping out). Mosaic type of DI RNAs containing three or more tracts of non-contiguous sequence can be formed by sequential events of the type shown in A . B. Generation of copy back R N A . 1 4 mediated model for strand switching in the BMV system was proposed whereby the replication complex functions in two different modes (Simon and Bujarski, 1994). In replication mode, the helicase portion of the replication complex efficiently unwinds any heteroduplexes which may form between the donor R N A and a potential acceptor R N A and synthesis continues along the donor RNA. In recombination mode, the donor R N A is released from the helicase. Consequently, upon encountering a heteroduplex, the replicase switches to a ss region on the acceptor template, thereby bypassing the double-stranded region. This mechanism can theoretically operate both inter- and intra-molecularly (e.g. for the generation of BBMV DI RNAs; see Pogany et al, (1995) and the elimination of introduced hairpin structures in BMV RNA; see Bujarski et al, (1994)). Recombination in BMV is believed to take place primarily during (-) R N A synthesis (Nagy and Bujarski, 1993). Unlike BMV recombinants, TCV recombinants (between TCV satellite R N A and TCV genomic R N A or between TCV satellite RNAs) show no obvious potential for heteroduplex formation at the crossover site. Instead, one of three conserved sequence motifs is invariably located at the right side of the crossover site (Cascone et al, 1990). The primary sequence of these motifs resembles putative promoters of positive polarity [(+)] R N A synthesis: motif I resembles the TCV 5' end; motif II resembles the 5' ends of some TCV satellite RNAs and DI RNAs; and motif III resembles the 5' end of a TCV subgenomic RNA. Secondary structure analysis of sequences around, and including, motifs I and III revealed similar stem loop structures (Cascone et al, 1993; Simon and Bujarski, 1994). The majority of crossover sites occur immediately upstream of this stem loop structure and mutations which disrupt the stem loop have been demonstrated to alter the crossover site (Cascone et al, 1993; Simon et al, 1994). Although this is suggestive of a specific involvement of this secondary structure in R N A recombination, the interrelationship is unclear at present (Simon and Bujarski, 1994). On the basis of these observations, a nonprocessive model was proposed where the TCV replicase 1 5 complex along with the nascent R N A detaches from the donor template and recognizes a specific motif on the acceptor template (or possibly the secondary structure surrounding the motif) as a reinitiation site (Simon and Bujarski, 1994). This same basic mechanism is though to be involved in the generation of TCV DI RNAs (Cascone et al, 1990) and in the generation of multimeric forms of T C V satellites and DI RNAs ' (Carpenter et al, 1991a). By contrast to both BMV and poliovirus, recombination in TCV appears to occur primarily during the synthesis of (+) R N A , although recent evidence suggests that recombination can also take place during synthesis of (-) R N A (Carpenter and Simon, 1994). As suggested by Simon and Bujarski, (1994), the differences in the models proposed for BMV and TCV could be a reflection of the differences in the RdRps of these viruses. B M V RdRp is a more "complete" enzyme than TCV RdRp as it contains a helicase domain whereas the TCV RdRp does not (Koonin and Dolja, 1993). A n important point not to be overlooked is that once generated, a DI R N A will be subjected to selective pressures which wil l ultimately determine whether or not it accumulates and how it wil l evolve. The maintenance of ds-acting elements involved in replication and packaging are two obvious factors which could influence DI R N A accumulation and evolution. In addition, the maintenance of an ORF appears to influence the accumulation of certain D and DI RNAs (de Groot et al, 1992; White et al, 1992; Kim et al, 1993; Romero et al, 1993; van der Most et a l , 1995). The selective pressures involved can be expected to vary depending on the parental virus, the host, and environmental conditions. Competition between DI RNAs can be used to study these selective pressures. Competition between TBSV DI RNAs in cucumber protoplasts results in a predominant accumulating DI R N A (White and Morris, 1994a; White and Morris, 1994c). Replication competency was identified as a major factor dictating DI R N A competitiveness in these studies. These studies also demonstrated directly that replication competency is a fundamental determinant of DI R N A evolution because the primary structure of DI 1 6 RNAs which accumulate from larger precursors after serial passaging were found to be the same as DI RNAs which are highly competitive. A recent study has also demonstrated DI R N A selection in animals. When mice were coinfected with a mixture of influenza DI RNAs, only a subset of the inoculum DI R N A s accumulated in lung tissue (Noble and Dimmock, 1995). Replication of (+) ss R N A viruses A l l (+) ss R N A viruses encode an RdRp which, in complex with viral-encoded and host-encoded subunits (replication complex), synthesizes copies of the viral genome (replication) and, in the case of some viruses, synthesizes sub genomic RNAs (transcription). The above definition of a replication complex has recently been expanded to incorporate viral R N A and certain host membranous structures (see Ahlquist et al, 1994; de Graaff and Jaspars, 1994; Quadt et al, 1995). Despite vast differences in host range, virion morphology, genome organization, and genome expression strategies, the molecular anatomy of the RdRps of (+) ss R N A viruses is remarkably similar. The known or putative RdRps of all (+) ss R N A viruses contain a glycine-aspartic acid-aspartic acid (G-D-D) motif (Kamer and Argos, 1984), which has been definitively correlated with polymerizing activity in the case of QP bacteriophage and poliovirus (Blumenthal and Carmichael, 1979; Jablonski et al, 1991). Other conserved sequence motifs and domains can also be observed among disparate (+) ss R N A viruses (nucleoside triphosphate binding motifs, helicase domains, and methyltransferase domains). Comparative analysis of the sequence of RdRps has been used to group (+) ss R N A viruses into three distinct supergroups (see Koonin, 1991) and to predict how (+) ss R N A viruses may have evolved from a common ancestor (see Koonin and Dolja, 1993). Another property common to RdRps is their high frequency of replication errors due primarily to the lack of inherent correction mechanisms (Holland et al, 1982; Reanney, 1982; Steinhauer 1 7 and Holland, 1987). The misincorporation rate for R N A replication has been estimated to be on the order of 10"3 to 10~4 per nucleotide per replication round for Qp\ vesicular stomatitis virus, and poliovirus (Batschelet et al, 1976; Steinhauer and Holland, 1986; Ward et al, 1988) which is 10 4 to 10 7 times higher than for D N A replication (Koonin and Gorbalenya, 1989). Replication of (+) ss R N A viruses in vivo can be thought of as occurring i n three successive phases. First, viral replication proteins (including the RdRp) must be translated directly from the genomic R N A and assembled into a functional complex with other host proteins in a specific cellular location. Complementary (-) R N A is then synthesized from genomic R N A templates, commencing at the 3' terminus of the template. Subsequently, (+) R N A is synthesized from (-) R N A templates, commencing at the 3' terminus of the template for the synthesis of genomic R N A or from an internal promoter for the synthesis of subgenomic R N A s . Synthesis of (+) R N A can be associated with replicative intermediates consisting of a (-) R N A template with bound nascent (+) RNAs of different lengths (David et al., 1992). The synthesis of (-) and (+) R N A is thought to be mechanistically different because more (+) than (-) R N A is produced during infections (asymmetric replication). The ratio of (+) to (-) RNAs has been estimated to be greater than 100 : 1 in BMV infections (Marsh et al., 1991), and 20 to 30 : 1 poliovirus infections (Andino et al., 1990). The identification of different cz's-acting structures used for initiating replication of RNAs of different polarity (see examples below) is also consistent with mechanistic differences between (+) and (-) R N A replication. Cz's-acting elements involved in (+) ss R N A virus replication have been characterized for many viruses. In general, sequences located at the 3' terminus of (+) R N A are recognized by the viral replication complex for the initiating (-) R N A synthesis. Specific structures have been implicated in .many viruses. tRNA-l ike structures which mimic cellular tRNAs both structurally and functionally (Hall et al, 1972; Bastin and Hall, 1976; Bujarski et al, 1985; Bujarski et al, 1986; Rao et al, 1 8 1989; Marsh et al, 1991) have been implicated in the case of barley stripe mosaic hordeivirus, BMV, C M V , cowpea chlorotic mottle bromovirus, tobacco mosaic tobamovirus (TMV), and turnip yellow mosaic tymovirus (David et al, 1992; Duggal et al, 1994) and possibly encephalomyocarditis virus, mengovirus, enterovirus, and rhinovirus (Lindley and Stebbing, 1977; Salomon and Littauer, 1974; Pilipenko et al, 1992). Pseudoknot structures have been implicated in the case of some viruses with tRNA-like 3' termini and poliovirus (Duggal et al, 1994; Jacobson et al, 1993). Stem loop structures have been implicated in the case of alfalfa mosaic ilarvirus, B N Y V V ; cowpea mosaic comovirus (CPMV), CyRSV; rubella virus and Sindbis virus (Koper-Zwarthoff and Bol, 1980; Houwing and Jaspars, 1982; Jupin et al, 1990; Eggen et al, 1989; Zoltan and Burgyan, 1995; Nakhasi et al, 1991; Strauss et al, 1990). There is a general requirement for an unstructured 3' terminus to facilitate the initiation of (-) R N A synthesis; an unstructured poly (A) tract in the case of some viruses and a C C A O H ending which is not involved in secondary structure in the case of viruses with a tRNA-like ending (Pogue et al, 1994). Cz's-acting sequences involved in (+) R N A synthesis from (-) R N A template reside in the 3' terminus of the (-) R N A and also in the 5' terminus of the (+) RNA. It has been suggested that the requirement for specific sequences at the 5' terminus of (+) R N A may facilitate the release of nascent (-) R N A from (+) R N A template thereby allowing it to be used as a template in the final phase of replication (Pogue et al, 1994). Again, specific structures have been implicated in many viruses: tRNA-like endings in the case of the 3' terminus of B M V (-) R N A (which accepts methionine in contrast to the tRNA-like structure at the 3' terminus of the (+) R N A which accepts tyrosine) stem loop structures at the 3' terminus of (-) R N A in the case of rubella virus and Sindbis virus (Nakhasi et al, 1991; Pardigon and Strauss, 1992); stem loop structures at the 5' terminus of (+) R N A in the case of B N Y V V , BMV, alfalfa mosaic virus, and Sindbis virus (Gilmer et al, 1993; Pogue and Hall, 1992; van der Vossen et al, 1993; Strauss et al, 1990); and a cloverleaf-like structure involving three stem loops in the case of the 5' terminus 1 9 of poliovirus (+) R N A (Andino et al, 1990). Interestingly, the cz's-acting sequences at the 5' end of B M V (+) R N A resemble the internal control region A and B boxes which promote the transcription of tRNA genes (Marsh and Hall, 1987), raising the possibility that they could be recognized by host transcription factors such as TFIIIC (Pogue et al, 1994). The requirement for sequence elements contained within viral coding regions (internal elements) in R N A replication has also been demonstrated (see Meyer et al, 1981; Ball and L i , 1993; Duggal et al, 1994; Pogue et al, 1994; Song and Simon, 1994). The mechanism by which these internal elements exert their influence on R N A replication is not well understood. In an effort to work out the mechanism of (+) ss R N A virus replication and identify host proteins involved, in vitro replication systems have been developed (reviewed in Blumenthal and Carmichael, 1979; Ishihama and Nagata, 1988; David et al, 1992; de Graaff and Jaspars, 1994; Ishihama and Barbier, 1994). Isolation of an active replication complex has proven to be an difficult task as replication proteins are produced at low levels in infections, multiple protein subunits are generally involved, and replication complexes are often membrane-associated and relatively insoluble. A n added complication is the presence of a virus-induced host RdRp i n plant virus infections (David et al, 1992). In fact, only a few replication complexes are capable of directing the complete replication (i.e. production of R N A of the same polarity as the input template) of an input R N A template in vitro (QP bacteriophage, see Blumenthal and Carmichael, 1979; cucumber mosaic C M V , see Hayes and Buck, 1990; flockhouse nodavirus, see Wu et al, 1992). Most replication complexes are only capable of synthesizing a complementary copy of input R N A template ((-) R N A from input (+) R N A or, in fewer instances, (+) R N A from input (-) RNA). The best characterized replication complex in terms of host proteins components is that of QP bacteriophage. Three different bacterial proteins have been identified: ribosomal protein SI, and translation elongation factors Tu and Ts 20 (Blumenthal and Carmichael, 1979). Ribosomal protein SI is known to have an R N A helicase activity which is thought to resolve R N A duplexes formed during R N A replication (Cole et al, 1982). A n additional protein component, HF-I, is also required specifically for (-) R N A synthesis (Ishihama and Barbier, 1994). The bacterial gene encoding HF-I {hfq) has been cloned from E. coli (Kajitani and Ishihama, 1991), however the function of this protein in E. coli is not yet known. A n interesting parallel to the QP replication complex has emerged from studies on the BMV replication complex, where translation elongation factor eIF3 has been implicated as a host protein required for BMV replication (Quadt and Jaspars, 1990; Quadt et al, 1993). A ribosome-associated protein has also been identified as a host factor required for (+) R N A synthesis in poliovirus (Andino et al, 1993). Host proteins with functions other than translation have also been implicated in (+) ss R N A virus replication. For example, calreticulin (a cellular calcium binding protein) and Ro/SS-A-associated antigens (targets of the autoantibodies implicated in autoimmune diseases) have been demonstrated to bind to cz's-acting elements involved in the translation and replication of rubella virus; however, their necessity for viral replication has not yet been demonstrated (Nakhasi et al, 1994). Cucumber necrosis virus Epidemiology C N V was first isolated in southwestern Ontario in the early 1950's as the causative agent of a severe disease in greenhouse cucumbers (McKeen, 1959). In greenhouse cucumbers, C N V causes necrotic spotting and malformation of the foliage, stunting of growth and reduced fruit size. The infection can become systemic, usually resulting in the death of the plant six to eight weeks after infection. The disease is most severe during the autumn and winter seasons and mildest i n the summer. The outbreaks in southwestern Ontario throughout the 1950's remain 2 1 the only reports of C N V infection and therefore cucumber remains the only known natural host of CNV. C N V infections can be effectively prevented by steam sterilization of soil, burning of diseased plants and sterilization of contaminated cultivation tools (Menzies and Jarvis, 1994). Consequently, C N V is not currently considered an economically important disease. Nevertheless, C N V is still recognized as a potential pest of greenhouse cucumbers in Canada. Classification C N V was originally classified as a member of the Necrovirus genus because the biology of transmission, and symptoms induced, were similar to tobacco necrosis virus, the type member of this genus (Matthews, 1982). Later, nucleic acid hybridization studies demonstrated that C N V is a member of the Tombusvirus genus even though, serologically, C N V appeared unrelated to TBSV, the type member of this genus (Rochon and Tremaine, 1988). The Tombusvirus group was first established as a distinct group of plant viruses in 1971 (Harrison et al, 1971). Currently, this former taxonomic group is now recognized as one of the two genera encompassed by the Tombusviridae family of plant viruses (Mayo and Martelli, 1993). The Tombusvirus genus presently consists of at least 13 definitive members (see Table 2). Tombusviruses as a whole have a wide geographic distribution, with the majority of the species found i n temperate regions. Although the natural host range of individual tombusviruses is limited, the experimental host range is very broad comprising over 100 plant species (Martelli et al, 1988). Nicotiana clevelandii and N. benthamiana are commonly used as experimental systemic hosts. Both TBSV and petunia asteroid mosaic virus are known to have weed hosts in addition to cultivated hosts, which probably serve as natural reservoirs of virus (Martelli et al, 1988). A l l tombusviruses are mechanically transmissable. In terms of natural vectors, C N V remains the only definitive member of the Tombusvirus genus whose transmission by a soil 22 organism has been unequivocally demonstrated (McLean et al, 1994; see below ). Tombusviruses share a similar particle structure of small icosahedrons with an approximate diameter of 30 nm. The three dimensional structure of the capsid and coat protein subunits of TBSV has been determined at a resolution of 2.9 A by X-ray crystallography (Harrison et al, 1978). The tombusviruses all have a similar genomic organization as deduced from the complete nucleotide sequences of C N V (Rochon and Tremaine, 1989), CyRSV (Grieco et al, 1989), TBSV (Hearne et al, 1990) and artichoke mottle crinkle virus (AMCV; Tavazza et al, 1994). Infection cycle The general infection cycle of C N V is shown schematically in Figure 2. The best understood steps of this cycle are vector transmission and expression of viral proteins. Certain aspects of viral R N A replication as well as the functions of some of the viral proteins are also known. A n overview of some of these steps are provided in the following subsections. Although the structure of the C N V virion is well established based on analogy with TBSV, no specifics are known regarding virion disassembly or assembly mechanisms. Vector transmission Most plant viruses depend on specific invertebrate or fungal vectors for their efficient transmission in nature (Matthews, 1991). C N V is transmitted to cucumber roots by the soil-inhabiting fungus Olpidium bornovanus (Dias, 1970; Stobbs et al, 1982; Campbell and Sim, 1994). The biology of this transmission is very specific; C N V is not transmitted by related Olpidium species (Dias, 1970) and viruses closely related to C N V are not transmissible by O. bornovanus (Martelli et al, 1988; McLean et al, 1994). O. bornovanus (also known in older literature as O. radicale Schwartz and Cook and O. cucurbitacearum Barr and Dias) is a Chytridiomycete and is an obligate parasite of cucurbits, including cucumber, squash, and melon (Barr, 1968). Table 2. Definitive members of the Tombusvirus genus. 23 Virus and acronym Artichoke mottle crinkle (AMCV) Carnation Italian ringspot (CIRV) Cucumber necrosis (CNV) Cymbidium ringspot (CyRSV) Eggplant mottled crinkle (EMCV) Grapevine Algerian latent (GALV) Lato River (LRV) Morrocan pepper (MPV) Neckar River (NRV) Perlargonium leaf curl (PLCV) Petunia asteroid mosaic (PAMV) Sikte waterborne (SWBV) Cultivated host artichoke carnation aiarmber cymbidium, white clover eggplant grapevine unknown pepper, tomato, eggplant, perlagonium unknown pelargonium petunia, cherry, grapevine, hop, pepper, plum, privet unknown Geographical distribution Mediterranean Great Britain, Italy, USA, Germany Canada Great Britain Lebanon Algeria Italy Morocco, Germany Germany Europe, Mediterranean, USA Central Europe, Canada Germany Original description Martelli (1965) Hollings et al, (1970) McKeen (1959) Hollings et al, (1977) Makkouk et al, (1981) Galitelli et al, (1989) Vovlas et al, (1989) Fischer and Lockhart (1974) Koenig and Lesemann (1985) Pape (1927) Lovisolo (1956) Li et al, (1992) Smith (1935) Tomato bushy stunt1 tomato, pepper, Europe, (TBSV) eggplant, lettuce, Mediterranean, spinach, tulip, apple, North and South piggyback, pear America Adapted in part from Martelli et al., (1988) and Russo et al., (1994); ^denotes type member of the genus. 2 4 Figure 2. Infection cycle of C N V : 1. virus is naturally acquired from soil and transmitted into root hair cells by the motile zoospores of Olpidium bornovanus; 2. virion is disassembled to release genomic RNA; 3. the replicase is translated from genomic R N A and copies of genomic R N A are synthesized; 4. additional viral proteins are translated from genomic R N A or subgenomic messenger RNAs; 5. progeny virions are assembled; 6. virions and/or genomic R N A move into neighbouring cells probably through the plasmodesmata connections between cells. Details of some of these steps are provided in the text. 2 5 During the life cycle of this fungus, motile zoospores are released from infected root cells into the surrounding soil. C N V attaches to the plasmalemma of the zoospore and gains entrance to the protoplasm of the zoospore by an unknown mechanism. The retraction of the flagellum by the zoospore during its encystment on a root cell may facilitate this process (Temmink and Campbell, 1969; Temmink et al, 1970; Stobbs et al., 1982). The coat protein of C N V is known to be involved in the specificity of transmission (McLean et al., 1994) and probably interacts with a specific factor(s) in the zoospore plasmalemma to facilitate attachment. Virus is transmitted to the root cell when the virus-containing encysted protoplasm is discharged into the root cell (Temmink and Campbell, 1969; Temmink et al., 1970; Stobbs et al., 1982). Viral RNA replication The general mechanism of genome replication in C N V is believed to be similar to that shared by other (+) ss R N A viruses as previously outlined. At least two subgenomic mRNAs are synthesized from (-) templates using internally-located promoters (Johnston and Rochon, 1990; Johnston and Rochon, 1995). Beyond these generalities, no details of the mechanism of C N V replication are known. The association of DI RNAs with tombusviruses has enabled the identification of putative cz's-acting sequences required for viral replication (White and Morris, 1994a; White and Morris, 1994c; Chang et al, 1995; Havelda et al, 1995; Havelda and Burgyan, 1995). Details of these studies wil l be discussed in subsequent chapters of this thesis. By analogy with the related tombusvirus CyRSV, the 92 kDa RdRp and the 33 kDa protein (see next section) are likely the only viral-encoded proteins which are absolutely required for C N V replication. Host factors associated with the C N V replication complex have yet to be identified. A l l tombusviruses cause the generation of multivesicular membranous bodies (MVBs) in the cytoplasm of infected plant cells which are likely the 26 intracellular site of viral replication (Martelli et al., 1988). Depending on the host, MVBs seem to be derived from either peroxisomes or mitochondria based on the ultrastructure of infected cells (Martelli et al., 1988). Glycolate oxidase, a peroxisomal enzyme, can also be detected in MVBs, further suggesting a peroxisomal origin (Martelli et al., 1984). The RdRps from all well-studied plant R N A viruses are generally known to be membrane-associated (David et al., 1992; de Graaff and Jaspars, 1994; Song and Simon, 1994). Recently, the viral-encoded replication proteins of TBSV (the 92 kDa RdRp and 33 kDa protein; see next section) were both demonstrated to be membrane-associated (Scholthof et al., 1995b), congruent with the idea of MVBs being the site of replication. Genomic organization and expression Like all tombusviruses, the C N V genome is a monopartite (+) ss R N A that is approximately 4.7 kb in size (Rochon and Tremaine, 1989). It is not known whether the 5' end of the C N V genome is capped; however, uncapped transcripts have been demonstrated to be infectious (Rochon and Johnston, 1991). The 3' end of C N V R N A is not polyadenylated (Rochon and Johnston, 1991) and the existence of secondary structures in the 3' end common to many plant R N A viruses (David et al., 1992; Duggal et al., 1994) has not yet been investigated. The C N V genome, shown in Figure 3, contains at least five distinct ORFs which encode protein products of 33-, 92-, 41-, 21- and 20 kDa (p33, p92, p41, p21, and p20, respectively; Rochon and Tremaine, 1989). A small ORF in the extreme 3' end of the genome may encode a sixth protein (Boyko and Karasev, 1992). p33, p41, p21 and p20 have all been observed to be translated in vitro in wheat germ extracts using C N V virion R N A as template (Johnston and Rochon, 1990). p33 and p92 are translated directly from the viral genome. p92, although not observed upon in vitro translation, is predicted to arise from readthrough of the amber termination codon of the ORF for p33 (Johnston and Rochon, 1990). p41 is translated from a separate 2.1 27 ORF5 ORF1 ORF2 ORF3 k \ V \ M Q R F 6 ? V////////X/////////////////A HI- 4.7 p33 p92 putative polymerase ORF4 p41 c o a t protein p21 movement protein p20 induction of necrotic symptoms -[]— 0.35 Figure 3. Organization and expression of the C N V genome. The boxed regions represent open reading frames (ORFs) with different reading frames indicated by-different shading patterns. The encoded protein products are diagrammed with arrows below their corresponding ORF; the known or putative function of proteins are indicated. Numbers at the far right indicate the sizes of the genomic and sub genomic RNAs in kb. 28 kb subgenomic mRNA (Johnston and Rochon, 1990). The ORF for p20 is completely-nested within the p21 ORF on a different reading frame (Rochon and Tremaine, 1989). Both proteins are translated from a single, bifunctional subgenomic m R N A which is 0.9 kb in size (Johnston and Rochon, 1990; Rochon and Johnston, 1991). The mechanism by which the internally-located A U G codon of the p20 ORF is accessed by plant ribosomes is likely leaky scanning (Johnston, 1995). In this mechanism, first proposed by Kozak, ribosomes bind to the 5' end of a mRNA and can scan past the first A U G codon if it is in a poor translational context initiating instead at a downstream A U G codon in a better translational context (Kozak, 1989). A 0.35 kb subgenomic R N A can be detected in C N V infections which encompasses ORF 6 (Johnston, 1995). It remains to be determined whether or not a functional protein is translated from this subgenomic R N A in C N V infections. Function of proteins The only C N V protein for which a function has been firmly established is the 41 kDa viral coat protein (Johnston and Rochon, 1990). The C N V virion is made up of 180 copies of this coat protein, arranged in a T=3 icosahedral symmetry (Tremaine, 1972). Based on analogy with the known crystal structure of the TBSV coat protein, the C N V coat protein forms three architecturally distinct domains: the amino-terminal, highly basic R N A binding domain; the shell domain which forms the surface of the viral particle; and the protruding domain which pairs with a neighbouring protruding domain to form projections from the surface. The shell and protruding domains form distinct jelly-roll barrels (Harrison et al, 1978), a basic core structure which is common to many icosahedral viruses (Branden and Tooze, 1991). Jelly-roll conformations have been found in a variety of proteins with known receptor binding functions (Branden and Tooze, 1991), lending further support to the notion that the C N V coat protein may attach to a specific receptor on O. bornovanus zoospores. 29 The C N V coat protein effectively protects the genomic R N A from degradation. C N V virions are stable in the presence of active plant ribonucleases for at least one hour (McLean et al, 1994; M . A. Robbins, personal communication). Tombusviruses in general are known for their hardiness: they retain infectivity for up to seven months in soil and for five weeks in concentrated plant sap; they can be isolated from rivers, lakes and drainage water; they can withstand heating at 90 -95°C in vitro; and TBSV can even pass unharmed through the alimentary tract of humans (Martelli et al, 1988; Tomlinson et al, 1982). The coat protein of C N V is known to be dispensable for many viral functions including systemic movement of the virus within the plant, induction of necrotic symptoms, viral R N A replication and generation of subgenomic mRNAs (McLean et al, 1993; Sit et al, 1995). The dispensable nature of the viral coat protein has also been observed in other tombusviruses (Dalmay et al, 1992; Scholthof et al, 1993). p92 contains the G-D-D motif which is known to be a signature of viral RdRps (Kamer and Argos, 1984) and has sequence similarity with other known or putative polymerases. It is therefore believed to form at least part of the viral-encoded replicase (Rochon and Tremaine, 1989). p33 does not contain any distinguishing motifs to help elucidate its possible function. According to the supergroup delineation for positive strand R N A viruses proposed by Koonin, (1991) based on sequence alignment of viral RdRps from animals and plants, C N V is a member of supergroup II, the "flavivirus-like" viruses (Koonin and Dolja, 1993). Other members of this group include other tombusviruses, carmoviruses, luteoviruses, dianthoviruses, necroviruses (all plant R N A viruses), pestiviruses and flaviviruses (animal R N A viruses) and certain R N A bacteriophage. Many members of supergroup II, including C N V , lack consensus methyltransferase (Gorbalenya and Koonin, 1989) and helicase domains (Lain et al, 1991) in their RdRps. With the exception of bacteriophage Q P , replication complexes from members of supergroup 30 II have not yet been characterized in terms of their composition and identity of host factors involved. In both CyRSV and TBSV, proteins analogous to p33 and p92 must both be expressed for viral replication; expression of the readthrough portion alone is insufficient to allow replication (Dalmay et al, 1993; Scholthof et al, 1995b). Furthermore, these proteins are likely the only viral-encoded proteins absolutely required for replication as evidenced by the fact that protoplasts prepared from transgenic N. benthamiana expressing the genes for the CyRSV p33 and p92 can replicate CyRSV DI RNAs (Kollar and Burgyan, 1994) and also by the fact that C N V / T B S V recombinants containing only the genes for C N V p33 and p92 can replicate in protoplasts (White and Morris, 1994b). Mutational, analysis of infectious transcripts generated from pK2/M5 has revealed possible functions for both p20 and p21 (Rochon and Johnston, 1991). Mutants of pK2/M5 which do not express the gene for p20 replicate in plants and infected plants show less necrosis than is typically associated with pK2/M5 infections (Rochon and Johnston, 1991). Similar symptoms are induced by related tombusviruses lacking the expression of the analogous gene (Dalmay et al, 1993; Scholthof et al, 1993), suggesting that this protein is involved either directly or indirectly in the induction of necrotic symptoms. Introduction of the pl9 gene from TBSV (analogous to the p20 gene from CNV) into a plant virus which normally does not induce necrotic symptoms results in necrotic symptoms in infected plants, providing further evidence for the role of this protein in the induction of necrotic symptoms (Scholthof et al, 1995a). Infections established with C N V mutants lacking expression of the gene for p20 have also been demonstrated to generate high levels of DI RNAs after a single high multiplicity of infection passage. A role for p20 in replication fidelity was also proposed on the basis of this observation (Rochon, 1991). 3 1 Mutants of pK2/M5 which do not express the gene for p21 do not accumulate to detectable levels in plants (Rochon and Johnston, 1991) but do replicate i n protoplasts (Johnston, 1995). Thus, p21 probably potentiates cell-to-cell movement of the virus (movement protein, see next section). A similar role has been proposed for the analogous proteins (p22) from both TBSV (Scholthof et al, 1993; Scholthof et al, 1995a) and CyRSV (Dalmay et al, 1993). p22 of TBSV has also been shown to induce necrotic symptoms in certain hosts (Scholthof et al, 1995a). Movement A fundamental difference between plant and animal cells is that plant cells are encased by a rigid cell wall. This structural barrier prevents plant viruses from moving into neighboring cells by either surface fusion with the cell surface or receptor-mediated endocytosis. Consequently, plant viruses have evolved at least two different mechanisms for moving from cell-to-cell (also known as short distance movement). Both mechanisms utilize specific viral-encoded proteins to facilitate cell-to-cell movement (referred to as movement proteins) and involve transport of viral particles or viral nucleic acid through plasmodesmata. Plasmodesmata are trans-cell wall channels between plant cells which provide a cell-to-cell continuity of cytoplasm, plasma membrane, and endoplasmic reticulum (via the desmotubule, an extension of the endoplasmic reticulum which runs through the channel; Lucas and Wolf, 1993). In the first mechanism, typified by TMV, non-encapsidated R N A genomes are thought to move through plasmodesmata which have been modified by the T M V movement protein to increase their size exclusion limit (Deom et al, 1992). The T M V movement protein also acts as a ss nucleic acid binding protein in vitro (Citovsky et al, 1990; Citovsky et al, 1993), raising the possibility that the viral genome is transported as a ribonucleoprotein complex. In the second mechanism, typified by CPMV, whole virus particles move through distinctly modified plasmodesmata (Goldbach et al, 32 1994). Electron microscopy of CPMV-infected plants has revealed tubular structures penetrating through plasmodesmata which are filled with virus particles presumably in transit to neighboring cells. It has been proposed that the desmotubule is somehow removed from the plasmodesmata to allow tubule formation (Goldbach et al, 1994). Both of the aforementioned mechanisms have been proposed to explain short distance movement of viruses in plants. Systemic infection by plant viruses further requires movement into, through, and out of the vascular tissue of the cell, usually the phloem (Leisner and Howell, 1993). This process is termed long distance movement to distinguish it from cell-to-cell movement. Many host factors affect long distance movement including leaf age, rate of plant development, plant anatomy, and direction of nutrient flow (Leisner and Howell, 1993). In terms of specific viral factors involved in long distance movement, movement proteins are clearly still a requirement. The mechanism of movement into and out of vascular tissue, however, is likely very different than the mechanisms discussed above for movement between non-vascular tissues (see Leisner and Howell, 1993). Wi th in vascular tissue, virus is thought to diffuse through the sieve plates connecting cells (which have a large enough pore size to obviate the need for a specific movement mechanism). In addition to movement protein, viral coat protein is absolutely required in the case of some viruses, even those which do not appear to require coat protein for short distance movement (e.g. TMV; Dawson et al, 1988). With other viruses, the coat protein appears to be dispensable, however, this can depend on the host/virus combination (e.g. red clover necrotic mosaic virus; Xiong et al, 1993) and both the rate and pattern of long distance movement can be altered in the absence of viral coat protein (e.g. CNV; Sit et al, 1995). Other viral proteins, whose effects are more subtle (e.g. BMV 2a protein (Traynor et al, 1991) and CyRSV p21 (Russo et al, 1994)) have also been implicated. Long distance movement is clearly a complex process requiring more studies to distinguish between direct and indirect effects. 3 3 Movement of C N V from cell-to-cell requires expression of the gene for p21 (Rochon and Johnston, 1991; Johnston, 1995). Coat protein is not absolutely required for short and long distance movement (McLean et al, 1993) but a delay in long distance movement and a different pattern of long distance movement are observed in the absence of functional coat protein (Sit et al, 1995). The dispensable nature of the coat protein with respect to movement may implicate a TMV-type of mechanism (described above) for cell-to-cell movement of C N V . Rationale of study The overall objective of this thesis was to further our understanding of the replication and accumulation of a simple, (+) ss R N A plant virus by studying DI RNAs of defined sequence. DI RNAs are valuable tools for studying viral replication for two main reasons. First, DI RNAs have lost much of the viral genome while retaining the cz's-acting sequences required for replication. This provides an excellent starting point for defining the minimal sequence requirements for replication and also allows these sequences to be studied independently of other viral regulatory sequences. Second, DI RNAs are generated and can evolve by the process of R N A recombination. Since R N A recombination is mediated by the viral replication complex, it can be considered to be an aberrant activity of the viral replication complex. Thus, studying the generation and evolution of DI RNAs can provide insights into the workings of the viral replication complex. As the association of DI RNAs with plant viruses was a relatively recent discovery at the commencement of the studies in this thesis, these studies also serve to expand the knowledge base on aspects of the basic biology of plant virus DI RNAs. Any biotechnological application of plant virus DI R N A s , such as the development of transgenic plants expressing DI RNAs, requires a solid foundation on the basic biology of DI RNAs in order to assure their success. These 3 4 studies first required the establishment of biologically active cDNA clones of C N V DI RNAs which would serve as a perpetual source of DI RNAs of defined sequence. This work constitutes the first chapter of this thesis. The second chapter explores a novel discovery to arise during the course of these studies; the identification of dimeric forms of a tombusvirus DI RNA. These studies are discussed in terms of the insights they provide into C N V R N A replication, accumulation and R N A recombination. A model for the generation of C N V DI R N A dimers is also proposed. 35 MATERIALS A N D METHODS The methods described in this chapter employed certain procedures that are routinely used in molecular biology. These procedures include large and small scale isolation of plasmid DNA, precipitation of nucleic acids, organic extraction of nucleic acids, agarose gel electrophoresis, visualization and photography of agarose gels, ligation of D N A fragments, transformation of Escherichia coli, and growth and manipulation of E. coli cultures. Details of these procedures can be found in the laboratory manual "Molecular Cloning" by Sambrook et al, (1989). The preparation and handling of the various solutions and reagents used for the methods described in this chapter, are also discussed in this manual. Restriction enzymes, D N A -modifying enzymes, and products employed to purify nucleic acid from agarose gels were all used according to the manufacturer's specifications. Materials Chemicals used for the preparation of buffers and various other solutions were obtained from: Sigma (St. Louis, MO); Fisher (Nepean, Ont.); and BDH (Toronto, Ont.). Bacterial culture media was obtained from Gibco (Burlington, Ont.). Nucleotides were obtained from Pharmacia (Baie d'Urfe, Que.); radio-active nucleotides were obtained from: Amersham (Oakville, Ont.); Dupont (Mississauga, Ont.); and ICN (Mississauga, Ont.). Restriction enzymes were obtained from: Bethesda Research Laboratories (Burlington, Ont.); Boehringer Mannheim (Laval, Que.); New England Biolabs (Mississauga, Ont.); Pharmacia (Baie d'Urfe, Que.); and Promega (Madison, WI). Several products were routinely used to purify nucleic acids from agarose gels: either Gene Clean (BiolOl, La Jolla, CA) or Qiaex (Qiagen, Chatsworth, CA) for D N A and RNAaid (BiolOl, La Jolla, CA) for RNA. Sources of 36 other specialty chemicals, enzymes and reagents are referenced where appropriate within the text. Sources of C N V DI RNAs Virion R N A extracted from a laboratory culture of C N V (CNV-Lc) was obtained from D. M . Rochon. This R N A extract harbored small amounts of a low molecular weight virus-related R N A species suspected to be DI RNAs (Rochon and Johnston, 1991). R N A extracts from plants in which low molecular weight R N A s had been generated de novo by serial high multiplicity of infection (moi) passaging of an infection established with infectious transcripts of C N V (Rochon, 1991) were also obtained from D. M . Rochon. Oligonucleotides used in this study Oligonucleotides used for the various analyses in this study are listed in Table 3. A l l oligonucleotides were synthesized by the Nucleic Acid and Protein Synthesis Laboratory, University of British Columbia (Vancouver, B. C) . Purification of oligonucleotides was carried out according to procedures recommended by this facility. Oligonucleotides were purified by two different methods. Initially, a Ci8 Sep-Pak Classic chromatography cartridge (Millipore, Nepean, Ont.) for solid phase extraction was prepared by first passing 10 ml of HPLC grade acetonitrile (BDH, Toronto, Ont.) through the cartridge, followed by 10 ml of sterile water. The crude oligonucleotide, resuspended in 1.5 ml of freshly-prepared 0.5 M ammonium acetate, was loaded on to the Sep-Pak cartridge using a 3 ml disposable syringe. The cartridge was washed with 10 ml of sterile water, then purged with air to displace all liquid. One ml of 40% acetonitrile was then passed through the cartridge and the eluant containing the oligonucleotide was collected; this step was repeated twice. TABLE 3. Synthetic oligonucleotides used in this study 37 Name Sequence^ Use(s) C N V 1 A C C C A G T C T T C A A A C C D N A and RNA sequencing, RT-PCR C N V 4 C C C A T A C G A T G A C G A G D N A sequencing C N V 8 AAATTCTCCAGGATTTCT D N A sequencing, construction of pDISS9 and pDILS21 C N V 9 GGGCTGCATTTCTGCA D N A sequencing C N V 29 T C T G G A T C C T A A T A C G A C T C A C T A construction of DI R N A cDNA T A G A A A T T C T C C A G G A T T T C T C clones C N V 32 AGACCCGGGCTGCATTTCTGCAA construction of DI R N A cDNA TGTTC clones, pDI42/Dl, pDISS9, pDILS21, and pDILS21 / A T A A T C N V 33 T A A C T T C C A G T A A A C G A C G A D N A sequencing, RT-PCR C N V 34 C T A T A C G T C C T A A A C T G C A T D N A sequencing C N V 38 T A C G C A G A C A A C A C G C R N A sequencing C N V 39 C C T A A T A C G A C T C T C T A G A G A A A construction of pDI42/Dl TTCTCCAG C N V 49 G A C T G C A G A C T C T C C A C A G construction of pDISS9 and pDILS21 C N V 50 A G A C T G C A G T A A G A C A G A C T C T T construction of pDISS9 C N V 57 A G A C T G C A G A T C A C G C C C T C A A A construction of pDILS21 A T A A A G A A A G C G A G T C A G A C A G ACTCTTCAGTCTGACT C N V 59 G G T G G A A T C T T G C G A A T T T A A C T construction of GTGCAGTTTAGGACG pDILS21/ATAAT J-All sequences shown 5' to 3' (left to right); underlining indicates a restriction site; boldface type indicates a promoter sequence; nucleotides in italics are mutations relative to the hybridizing D N A template. Specific features and applications of each oligonucleotide are detailed in the text. 38 The concentration of oligonucleotide in each of these three fractions was determined spectrophotometrically and then fractions containing oligonucleotide were dried under vacuum. The oligonucleotide was resuspended in sterile H 2 O and stored at -20°C. Earlier in these studies oligonucleotides larger than 40 nucleotides were first separated on 12% polyacrylamide gels containing 7.7 M urea. The gel was transferred to a sheet of plastic wrap and the desired oligonucleotide band was identified by ultraviolet shadowing. A piece of Parafilm (American National Can, Greenwich, CT) was placed beneath the gel and the gel was illuminated from above with short wave length ultraviolet light, allowing oligonucleotide bands to be visualized as dark bands or shadows against a fluorescent background. The oligonucleotide band was excised using a razor blade and incubated overnight in 0.5 M ammonium acetate at 37°C. The supernatant was removed from the gel slice and the gel slices were washed twice more with 0.5 M ammonium acetate. The wash supernatants were pooled with the original supernatant and this was passed through a 45 micron filter (Millipore) to remove any gel fragments. This solution was then passed through a Cl8 Sep-Pak as described above. Later on in this study, oligonucleotides were prepared using a rapid method described by Sawadogo and Van Dyke (1991). Crude oligos were resuspended in 100 ul of 30% N H 4 O H and then vortexed with 1 ml of n-butanol for 15 sec. The mixture was then centrifuged at 14, 000 rpm in a microcentrifuge to pellet the oligonucleotide. The supernatant was removed from the pellet and the pellet was dried under vacuum, resuspended and stored as outlined above. Near the end of this study, oligonucleotides were available in a purified, ready-to-use format. Plasmids used in this study Plasmids used in this study are listed in Table 4 or referenced within the text. A l l plasmids were propagated in E. coli D H 5 a and purified according to standard TABLE 4. Plasmid constructs unique to this study! Name Description pDI9, pDI12, pDI15, pDI16 pDI21, pDI42 pDI48 pDI42/Dl pDI9/21PE pDI9/21PT pDI9/21TE pDI21/9PE pDI21/9PT pDI21/9TE pDISS9 pDISS9/21TE pDILS21 pDILS21/9TE pDILS21 / ATAAT cDNA sequence of DI RNAs generated de novo in pK2/M5 infections cDNA sequence of DI RNAs associated with CNV-Lc head-to-tail repeat of DI R N A 42 cDNA sequence pDI9 containing a PflMl/ EcoKL fragment from pDI21 pDI9 containing a PflMl/Tfil fragment from pDI21 pDI9 containing a Tfil/EcoRI fragment from pDI21 pDI21 containing a PflMl/EcoRI fragment from pDI9 pDI21 containing a PflMl/Tfil fragment from pDI9 pDI21 containing a Tfil/EcoRI fragment from pDI9 deletion mutant of pDI9 lacking additional nucleotides at the beginning of region III pDISS9 containing a Tfil/EcoRI fragment from pDI21 mutant of pDI21 containing additional nucleotides at the beginning of region III pDILS21 containing a Tfil/EcoRl fragment from pDI9 mutant of pDILS21 lacking nts 341 through 344 l A l l constructs shown are pUC19-based (Yanisch-Perron et al., 1985); specific variations within the multiple cloning site or other portions of pUC19 are discussed in text. 40 procedures (Sambrook et al, 1989). Specific approaches for the construction of plasmids unique to this study are discussed in the following subsections. Construction of cDNA clones of CNV DI RNAs Complementary D N A clones of potential DI RNAs in CNV-Lc were obtained using reverse transcription followed by the polymerase chain reaction (RT-PCR) as diagrammed in Figure 4. The first strand of cDNA was generated by reverse transcription (RT) in a 20 | i l reaction volume using 200 U Moloney murine leukemia virus (MMLV) reverse transcriptase (Bethesda Research Laboratories) i n the presence of IX Taq D N A polymerase buffer (20 m M Tris-HCl, p H 8.4, 50 m M KC1; Bethesda Research Laboratories), 1 m M each of dATP, dCTP, dGTP, and dTTP, 10 m M MgCl2, and 20 U of RNAasin (Promega) or 15 U of R N A guard (Pharmacia). Approximately 350 ng of CNV-Lc virion R N A preparation or de nouo-generated R N A extracts were mixed with 50 pmoles of a synthetic oligonucleotide complementary to the 3' terminus of C N V R N A (CNV 32), heated at 90°C for 2 min, and chilled on ice. C N V 32 incorporated a Smal site to facilitate cloning of the RT-PCR products recovered (Table 1). The remaining components of RT reaction were added and the reaction mixture incubated for 30 min at 37°C. The reverse transcriptase was inactivated by heating the reaction mixture at 95°C for 5 min. The RT mixture was then combined with 50 pmol of a synthetic oligonucleotide complementary to the 3' terminus of first strand cDNA (CNV 29), 2 U of Taq polymerase (Bethesda Research Laboratories) and IX Taq buffer to a final volume of 80 uL. C N V 29 incorporated the bacteriophage T7 R N A polymerase promoter and a BamHl site to facilitate cloning of the RT-PCR products recovered (Table 1). The reaction mixtures were overlaid with mineral oil and subjected to 40 cycles of 25 sec at 95°C, 50 sec at 50°C, and 2 min at 72°C. PCR products were gel-purified, digested with BamHl/ Smal, and ligated into similarly digested pUC19 4 1 DI R N A 5' cDNA CNV 29 3' dsDNA 5'. 3' CNV 32 - 3* reverse transcription 5' ^ polymerase chain reaction 3' • 5' digest with BamHl and Smal I ligate into Bam HI / Sraal-digested pUC19 B BamHl Accl PflMl Tfil - ^ ^ 1 i I » I Smal 111 Figure 4. Construction of complementary D N A clones of C N V DI RNAs. A . Cloning strategy: dashed line represents RNA; solid lines represent D N A ; solid lines with small arrowheads represent oligonucleotides. Oligonucleotides C N V 29 and C N V 32 contain BamHl and Smal sites respectively, C N V 29 also contains the T7 promoter. B. Cloned C N V DI R N A sequences in pUC19: shaded box represents cDNA sequence shown 5' to 3'; arrow represents T7 R N A polymerase promoter; solid line represents multiple cloning site of pUC19; dashed line represents plasmid sequence outside the multiple cloning site. Restriction endonuclease sites used i n cloning as well as sites within the cDNA sequence used for the construction of chimeric and mutated DI RNAs are indicated. 42 vector. The ligation mixture was used to transform competent cells. Plasmid was extracted from transformants and screened by double-digestion with BamHI and Smal. Plasmids containing BamHI/Smal inserts were retained for sequencing as described in the "DNA sequencing " section below. Construction of a head to tail repeat of DI RNA cDNA sequence The complete pDI42 insert was amplified using PCR with oligonucleotides C N V 32 and C N V 37. C N V 37 was designed to introduce a Xbal site immediately upstream of the 5' end of DI R N A 42 sequence. The resulting PCR product was gel-purified and ligated into a T-tailed intermediate vector (pT7Blue; Novagen)[T-tailed vectors like pT7Blue take advantage of the fact that Taq polymerase leaves 3' A overhangs to greatly facilitate the cloning of PCR products (Holton and Graham, 1991; Marachuk et al, 1991]. A Xbal/Smal fragment encompassing DI R N A 42 cDNA sequence was excised from this intermediate vector, treated with mung bean nuclease, and ligated into Smal-digested pDI42. This ligation mixture was used to transform competent cells and transformants were screened by restriction analysis. To confirm the junction sequence of potential head to tail dimer clones, AccI fragments were directly sequenced as described in the "Direct sequencing of RT-PCR products and restriction fragments" section below. Construction of chimeric DI RNAs The construction of chimeric DI RNAs utilized unique restriction sites within the DI R N A sequence (see Figure 3B) and the EcoRI site in the multiple cloning site of the vector. To make Tfil a unique restriction site within pDI9 and pDI21, the Tfil sites had to first be removed from the vector. To do this, pUC19 was digested with Tfil, treated with mung bean nuclease, religated and used to transform competent cells. Transformants were screened for loss of ability to be digested with Tfil (pUC19AT/i'i). BamHI/EcoRI fragments from either pDI21 or pDI9 were then ligated 4 3 into BamHI/EcoRI-digested pUC19AT/z'I to yield pDI21 and pDI9 clones in which Tfil was a unique restriction site. To construct the various chimeric DI RNAs, the appropriate insert and vector combinations were prepared and ligated together. This ligation mixture was used to transform competent cells and transformants were screened by restriction endonuclease digestion. The chimeric nature of all constructs was confirmed by sequencing. Construction of short stem and long stem mutants. To construct a short stem (SS) mutant of DI R N A 9, two portions of DI R N A 9 cDNA sequence were amplified independently from pDI9 template using PCR and ligated into a modified pDI9 vector. To facilitate this three piece ligation (see Figure 5), the AccI site was first removed from the multiple cloning site of pDI9. To do this, pDI9 was digested with PstI and Xbal, treated with mung bean nuclease, religated and used to transform competent cells. Transformants were screened for loss of ability to be digested with Xbal and Pstl (pDI9APX). A portion of pDI9 encompassing most of region I and region II (5' portion) was amplified using C N V 8 and C N V 49. A second portion encompassing most of region Ilia/nib but excluding 27 of the 28 additional nucleotides which are unique to DI R N A 9 (3' portion) was amplified using C N V 50 and C N V 32. A Pstl site was incorporated into both C N V 49 and C N V 50 to allow the amplified halves to be religated in a three piece ligation. Each PCR product was ligated into a T-tailed intermediate vector. A n AccI/Pstl fragment from the 5' portion and a Pstl/Smal fragment from the 3' portion were prepared from these intermediate vectors and ligated into an Acd/Smal-digested pDI9APX vector. This ligation mixture was used to transform competent cells and transformants were screened by restriction endonuclease digestion. The pDISS9 construct was completely sequenced to confirm that PCR did not introduce any unintentional nucleotide changes into this construct. A similar strategy to that depicted in Figure 5 was adopted for the 44 C N V 8> C N V 50, II III C N V 49 : C N V 32 pDI9 p p Accl/Smal pDI9APX + J ligate M pDISS9 Figure 5. Strategy for construction of pDISS9. Shaded boxes represent DI R N A 9 cDNA sequence, horizontal arrows represent oligonucleotide primers used for PCR, and "V" represents sequence deleted from pDISS9 relative to pDI9. Restriction endonuclease sites are designated with letters: A = AccI; P = PstI; S = Smal. A similar approach was used for the construction of pDILS21. See text for details. ' - 4 5 construction of a long stem (LS) mutant of DI R N A 21. To facilitate this three piece ligation, the AccI site was removed from the multiple cloning site of pDI21 to yield pDI21APX in a manner similar to that described above for the construction of pDI9APX. A 5' portion of DI R N A 21 cDNA sequence was amplified using PCR with C N V 8 and C N V 49 and pDI21 template and a 3' portion of DI R N A 9 was amplified using PCR with C N V 32 and C N V 57 and pDI9 template. Repeated attempts to use C N V 57 and C N V 32 to amplify a product from pDI21 were unsuccessful. Consequently, pDI9 was used as a template for amplification of the 3' portion. C N V 57 contained the additional 28 nucleotides unique to pDI9 with an incorporated Pstl site to allow the amplified halves to be religated in a three piece ligation. Each PCR product was ligated into a T-tailed intermediate vector, an AccI/Pstl fragment from the 5' portion and a Pstl/Smal fragment from the 3' portion were prepared from these intermediate vectors and ligated into Accl/Smal-digested pDI21APX. This ligation mixture was used to transform competent cells and transformants were screened by restriction endonuclease digestion. The 3' portion of this construct was then restored to the sequence of pDI21 by replacing the Tfil/EcoRI fragment with a Tfil/EcoKl from pDI21. The final pDILS21 construct was completely sequenced to confirm that PCR did not introduce any additional nucleotide changes into this construct. To construct pDILS21/ATAAT, a mutant which harbors both of the major sequence differences seen between DI R N A 21 and DI R N A 9, a 3' portion of DI R N A 21 cDNA sequence was amplified using PCR with C N V 59 and C N V 32 and pDI21 template. C N V 59 contained the Tfil site (see Figure 4B) and was designed to specifically eliminate nucleotides 341 to 344 of DI R N A 9. The PCR product was ligated into a T-tailed vector, subcloned into pUC19AT/zT , then a Tfil/EcoKl fragment encompassing the modified pDILS21 3' end was prepared from this subclone and ligated into T/zI/EcoRI-digested pDILS21. The final pDILS21/ATAAT 46 construct was sequenced to confirm that PCR did not introduce any additional nucleotide changes. D N A sequencing Double-stranded plasmid D N A templates were sequenced using the dideoxy chain termination method of Sanger et al. (1977) and Sequenase (U. S. Biochemical Corporation, Cleveland, OH) using a protocol obtained from the suppliers which was based on the method of Toneguzzo et al. (1988). Approximately 2 ug of plasmid D N A obtained by alkaline lysis "mini-prep" method (Sambrook et al, 1989) and 10 ng of synthetic oligonucleotide primer were denatured in a 40 ul volume containing 200 mM NaOH and 0.4 m M EDTA by heating at 85 to 95°C for 5 min. Reactions were chilled on ice and adjusted to 200 m M ammonium acetate and precipitated with absolute ethanol. The pellet was washed with 70% ethanol and resuspended in 6 u l of H20 and 1.5 ul of 5X Sequenase buffer (IX is 40 m M Tris-HCl, p H 7.5, 20 m M MgCl2, 50 m M NaCI) and incubated at 37°C for 15 min to allow annealing of the primer to the denatured D N A template. To this solution was added 1 ul of 100 m M dithiothreitol (DTT), 2 ul labeling mix (IX labeling mix is 1.5 uM each of dCTP, dGTP, dTTP), 2 to 5 uCi of a - 3 2 P dATP (3000 Ci/mmol) and 3 U of Sequenase in a total volume of 13 ul. Reactions were allowed to stand at room temperature for 2 to 5 min and 3 ul volumes from this reaction mixture were added to tubes containing 2.5 uL of 80 uM each of dGTP, dCTP, dTTP, dATP, 50 m M NaCI and 8 uM of either dideoxy (dd) GTP, ddCTP, ddTTP or ddATP and incubated a further 5 to 20 min at 37°C. To each reaction tube was added 4 ul of mixture containing 95% formamide, 20 m M EDTA, 0.05% bromophenol blue, and 0.05% xylene cyanol. Sequencing reactions were denatured by heating at 75 to 85°C for 2 min before loading onto a sequencing gel (described below). 47 The above protocol was used for sequencing all DI R N A cDNA clones which constituted the majority of the sequencing performed in this study. Later, sequencing reactions were performed to verify the sequence of various constructs made throughout this study and to analyze the sequence variability at the junctions of dimer RNAs. These later sequencing projects employed a slightly modified version of the above sequencing protocol described by Hsiao (1991). Approximately 2 ug of double stranded D N A template in a 5 ul volume was mixed with 1 ul of sequencing primer (10 ng/ul) and 1 ul of 1 N NaOH. This mixture was incubated at 37°C for 10 min, then 1 ul of 1 N HC1 was added to neutralize the mixture along with 2 ul of 5X Sequenase buffer. This mixture was then incubated at 37°C for 5 min to allow annealing of primer and template. The remaining components of the sequencing reaction were added as outlined above except that a-^^S dATP was used in place of cc-32p dATP. Labeling and termination reactions and denaturation of templates prior to electrophoresis were carried out as outlined above. Sequencing reactions were electrophoresed through 6% polyacrylamide gels, either 0.4 mm thick gels or wedge gels (0.2 to 0.6 mm gradation), containing 7.7 M urea at constant power (50-55 watts). Following electrophoresis, gels were transferred onto filter paper and dried under vacuum at 80°C for 1-2 hr and exposed to X-Omat (Kodak, Nepean, Ont.) or Hyper film (Amersham) overnight or longer if necessary at room temperature. Direct sequencing of RT-PCR products and restriction fragments A modified version of the dideoxy chain termination procedure adapted from Winship (1989) was used for directly sequencing the population of RT-PCR products generated from dimer RNAs and also for sequencing Accl fragments from potential head to tail dimer clones. RT-PCR products, which ranged in size from approximately 400-500 bp depending on the dimer template used, and Accl 48 fragments which were in the same size range were purified from agarose gels. For each sequencing reaction, approximately 200 ng of gel-purified template was mixed with 20 pmol of sequencing primer and 2 ul of 5X Sequenase buffer in a final volume of 10 ul. This mixture was heated at 100°C for 3 min to denature the template, chilled on ice, and then centrifuged briefly at 14, 000 rpm to pool the small volume at the bottom of the centrifuge tube. This mixture was then incubated at 37°C for 5 min to allow annealing of primer and template. The remaining components of the sequencing reaction were added as outlined in the previous section using a-35s dATP to label synthesized DNA. Labeling and termination reactions, denaturation of templates prior to electrophoresis, gel electrophoresis and gel manipulations were carried out as outlined in the previous section. Typically, these gels required at least a 2 day exposure to enable reading of the sequence. RNA sequencing R N A templates were sequenced using dideoxy terminators, oligonucleotide primers and reverse transcriptase following a protocol essentially similar to that described in the "Protocols and Applications Guide" published by Promega (Madison, WI). DI R N A dimer templates were purified from nondenaturing agarose gels (using RNAaid; see Materials) of total nucleic acid extracted from leaves coinfected with DI R N A 9; DI R N A monomer templates were purified from nondenaturing agarose gels of total nucleic acid extracted from leaves coinfected with DI R N A 42. Genomic length C N V R N A , purified from virions, was obtained from D. M . Rochon. Five hundred ng of gel-purified R N A templates or 2 ug of genomic length R N A were mixed with 25 ng of complementary oligonucleotide primer in a final volume of 6 ul. This mixture was heated at 67°C for 3 min, slow cooled over approximately 30 min to room temperature to allow annealing of primer and R N A template, and then centrifuged briefly at 14, 000 rpm to pool the small volume at the bottom of the centrifuge tube. To this mixture was added: 6 u l of 5X M M L V first strand buffer (IX is 50 m M Tris-HCl, p H 8.3, 75 m M KC1, 3 m M MgCl2), 40 U of RNAasin (Promega) or 30 U of R N A guard (Pharmacia), 2 uls of 100 m M DTT, 50 uCi of oc- 3 2P dATP (3000 Ci/mmol), and 200 U of M M L V or Superscript reverse transcriptase (Bethesda Research Laboratories) to a final volume of 21 ul. Four ul volumes of this mixture were immediately dispensed into tubes containing 2.5 ul of 250 uM each of dGTP, dCTP, and dTTP, and 12.5 uM of ddCTP, 12.5 uM ddGTP, 50 uM ddTTP or 1.0 uM ddATP in 34 m M Tris-HCl, p H 8.3, 50 m M NaCI, 6 m M MgCl2, 5 m M DTT and one tube containing 2.5 uls of sterile H2O to identify regions on the R N A template where RT stops independently of the incorporation of a chain terminator ("hard stops"). The five tubes were then incubated at 37°C for 15 min, then 1 ul of a chase mix consisting of 2 m M each of dATP, dCTP, dGTP, and dTTP in 34 m M Tris-HCl, p H 8.3, 50 m M NaCI, 6 m M MgCl2 was added to each tube and incubation at 37°C continued for an additional 15 min. Reactions were stopped by the addition of 4 ul of a mixture containing 95% formamide, 20 m M EDTA, 0.05% bromophenol blue, and 0.05% xylene cyanol. Denaturation of templates prior to electrophoresis, gel electrophoresis and gel manipulations were carried out as outlined previously in the "DNA sequencing" section. RT-PCR analysis of dimer RNA junctions In order to determine the junction sequence between the monomer units of a dimer, RT-PCR was conducted essentially as described in the "Construction of complementary D N A clones of C N V DI RNAs" section above utilizing oligonucleotide C N V 1 to generate cDNA and primers C N V 1 and C N V 33 for PCR amplification. These primers amplify a ds D N A product specifically from (+) dimer RNAs. RT-PCR products were amplified from total nucleic acid extracts prepared 50 from leaves of plants coinfected with DI R N A and helper virus (approximately 1 ug of total nucleic acid), cloned into T-tailed vector, and then the sequences surrounding the junction sites were determined. RT-PCR products amplified from DI R N A 9 coinfections (12 clones) and DI R N A 42 coinfections (9 clones) were both sequenced. In vitro transcription Bacteriophage T7 R N A polymerase run-off transcripts of a full length, c D N A clone of C N V (pK2/M5; Rochon and Johnston, 1991) or, for certain experiments, from a cDNA clone lacking most of the coding region for the C N V coat protein [CP (-); McLean et al, 1993) and individual DI R N A cDNA clones were prepared as follows. Five ug of Smal-linearized D N A template was transcribed in 40 m M Tris, p H 8.0,10 m M NaCI, 8 m M MgC12, 2 m M spermidine-(HCl)3, 10 m M DTT, 0.5 m M each of ATP, CTP, GTP, and UTP, 20 U of RNAasin (Promega) or 15 U R N A guard (Pharmacia), and 100 units of T7 R N A polymerase (Bethesda Research Laboratories), in a 50 ul reaction volume at 37°C for 1 hr. Prior to plant inoculations, transcription reaction mixtures were mixed with 0.1 volume of 100 m M NaP04, pH 7.0. For coinoculations, 50 ul of DI R N A transcripts were mixed with 50 ul of helper transcripts prior to coinoculation. For control infections, 50 ul of helper transcripts were similarly diluted with 50 ul of 10 m M NaPC>4, p H 7.0. Coinoculation of Nicotiana clevelandii Approximately 0.5 ug of transcripts from each DI R N A cDNA clone premixed with approximately 5 ug pK2/M5 helper transcripts in 100 ul of transcription salts/10 m M NaP04, p H 7.0 was used as inoculum. The molar ratio of DI R N A transcripts to helper transcripts in this inoculum was approximately 1:1. Four to 5 1 five week old N. clevelandii was dusted with Carborundum (Aldrich, Milwaukee, WI) and then 25 ul aliquots of inoculum was applied to four leaves and gently rubbed across the entire leaf surface with a gloved finger. Plants inoculated with pK2/M5 transcripts alone (WT infection) or buffer alone (mock infection) were used as controls. Coinoculation with CP(-) helper transcripts were performed in a similar manner. A l l plants were maintained in a greenhouse under a day/night regime of 22°C/17°C with a 14 hr photoperiod. Plants were monitored every 1-2 days for symptom development. Extraction of total nucleic acid from Nicotiana clevelandii leaves Typically, leaves were removed for total nucleic acid extraction 5-6 days post inoculation (dpi). Approximately 0.1-0.5 mg of leaf material was ground to a fine powder in liquid nitrogen, mixed vigorously with 400 ul phenol/chloroform/ octanol (25:24:1), 400 ul 10X TNE (100 m M Tris-HCl, p H 7.5, 100 m M NaCI, 10 m M EDTA) containing 0.1% sodium dodecyl sulfate (SDS) and 5% P-mercaptoethanol and centrifuged for 2 min at 14, 000 rpm in a microcentrifuge. The aqueous phase was collected and re-extracted with phenol/chloroform/octanol and then with chloroform/octanol (24:1). The aqueous phase was combined with 0.1 volumes of 2 M Na acetate, p H 5.8 and precipitated with 2 volumes of absolute ethanol. The pellet was washed with 70% ethanol, dried briefly under vacuum and resuspended in 100 ul of sterile H2O. Typically, 1-2 ul of resuspended nucleic acids were analyzed by agarose gel electrophoresis. Northern blot analysis R N A purified as described in the previous section was denatured in a solution containing 5 m M methyl mercuric hydroxide (Johnson Matthey, Ward 52 Hil l , MA) and separated by electrophoresis through an agarose gel containing 5 m M methyl mercuric hydroxide (Bailey and Davidson, 1976). Sample preparation and electrophoresis was carried out in a fume hood. After electrophoresis, (3-mercaptoethanol was used to sequester mercury from the gel. R N A was then transferred from the gel to a Zeta Probe GT membrane (BioRad, Mississauga, Ont.) overnight under alkaline conditions (10 m M NaOH; Vrati et al., 1987) and the membrane was placed in a hybridization solution consisting of 250 m M Na2HP04, p H 7.2 and 7% SDS at 65°C. Prehybridization in this solution was carried out for at least 30 min prior to addition of probe. 32p_i a D e i e cl D N A probes were prepared by nick-translation (Sambrook et al, 1989) of 20 to 100 ng of linearized D N A in a reaction mixture consisting of 50 m M Tris-HCl, p H 7.5, 10 m M MgCl2, 0.1 m M DTT, 2.5 ug bovine serum albumin, 0.6 m M each of dCTP, dGTP, and dTTP, 50 uCi of a -32p dATP (3000 Ci/mmol), 250 pg DNAse I (Bethesda Research Laboratories), and 15 U of D N A polymerase I (Pharmacia). Incorporation of radioactivity into the probe was monitored by trichloroacetic acid precipitation and liquid scintilation counting, the reaction was stopped with the addition of 1 ul of 20% SDS and the probe was passed over a Sephadex G-50 (Pharmacia) spin column (Sambrook et al, 1989). The probe was denatured with 0.05 volume of 2M NaOH and boiling for 5 minutes before adding it to the hybridization solution. Hybridization was carried out for 16 to 22 hrs at 65°C, and the membrane was washed twice for 30 min in 20 m M Na2HP04, p H 7.2, 5% SDS and then twice for 30 min in 20 m M Na2HP04, p H 7.2, 1% SDS at 65°C. The washed membrane was exposed to X-Omat (Kodak) or Hyper film (Amersham) at room temperature. 53 CHAPTER 1. Sequence and biological properties of cloned CNV DI RNAs A crucial starting point for detailed molecular studies on C N V DI RNAs was to establish complementary D N A (cDNA) clones of C N V DI RNAs from which biologically active R N A transcripts could be synthesized by in vitro transcription. Biological activity implies that the R N A derived from a clone is able to accumulate in the presence of helper virus in vivo as well as to interfere with helper virus accumulation. This interference can be evidenced in plant virus infections by attenuation of symptoms and/or a reduction in viral R N A accumulation. Results Sources of DI RNAs for cloning Two different sources of C N V DI RNAs were available at the onset of this study for the construction of biologically active clones. It was previously reported that R N A extracts of plants infected with a laboratory culture of C N V (CNV-Lc) contained high levels of a low molecular weight (-400 nucleotide) viral-related RNA. Plants infected with CNV-Lc displayed attenuated symptoms in comparison to plants inoculated with synthetic wild-type (WT) C N V transcripts generated from a full-length cDNA clone of C N V (pK2/M5; Rochon and Johnston, 1991). This suggested that the 400 nucleotide RNAs associated with CNV-Lc infection were likely DI RNAs and so CNV-Lc was used as one source of DI RNAs. DI RNAs have been generated de novo from two other tombusviruses, tomato bushy stunt virus (TBSV) and cymbidium ringspot virus (CyRSV), by high multiplicity of infection (moi) serial passaging of viral infections (Burgyan et al., 1991; Knorr et al., 1991). DI RNAs were generated de novo by high moi serial passaging of an infection established with transcripts from pK2/M5 (WT transcripts) in Nicotiana clevelandii plants (Rochon, 1991). Use of synthetic WT transcripts for 54 inoculation insured that the starting material was DI RNA-free. After 14 serial high moi passages of this infection, large amounts of low molecular weight RNAs were detected in two of the six plants examined (Figure 6, lanes 2 and 3). These two plants also displayed attenuated symptoms and a persistent infection typical of CNV-Lc-infected plants. This was distinct from the severe systemic necrosis observed in the remaining four plants. A l l of the aforementioned work was carried out by D. M . Rochon prior to the start of the studies for this thesis. Leaf R N A extracts from the two plants carrying large amounts of low molecular weight RNAs were used as a second source of C N V DI RNAs. Cloning and sequence analysis of DI RNAs Previous sequence analysis of the DI RNAs associated with the related tombusviruses TBSV and CyRSV indicated that the 5' and 3' termini were derived from genomic R N A (Hillman et al, 1987, Burgyan et al, 1989). Consequently, cDNA clones of potential DI RNAs in CNV-Lc and the small RNAs generated de novo were obtained using reverse transcription followed by polymerase chain reaction (RT-PCR) using synthetic oligonucleotide primers specific for the 5' and 3' termini of C N V genomic RNA. This cloning strategy is depicted in Figure 4 of the preceding Materials and Methods chapter. The 3' oligonucleotide used, C N V 32, (all oligonucleotides referred to in this chapter are listed in Table 3 of the preceding Materials and Methods chapter) was complementary to the last 21 nucleotides of C N V R N A and contained a full Smal site at the 5' end for subsequent run-off transcription of the cloned sequences. The 5' oligonucleotide, C N V 29, contained sequences homologous to C N V R N A nucleotides 2 through 21 and also included the bacteriophage T7 R N A polymerase promoter immediately upstream of the C N V 5' sequence so that the cDNA clones obtained could be used to synthesize R N A with precise 5' termini. This oligonucleotide also incorporated a BamHl site upstream of the T7 promoter to facilitate cloning. PCR products were electrophoretically 55 separated on agarose gels, gel-purified, digested with BamHI and Smal, and ligated into similarly digested pUC19 vector. Due to the cloning strategy used, transcripts from the recovered C N V DI R N A cDNA clones contained two introduced changes relative to C N V . The 5'-most nucleotide of C N V was eliminated in oligonucleotide C N V 29 in order to place the T7 promoter immediately upstream of a G (defined here as C N V nucleotide 2 although the identity of this nucleotide was unknown at the commencement of these studies). Using the introduced Smal site to linearize plasmid templates for run-off transcription also introduced an extra C at the 3' terminus of transcripts. Four cDNA clones from CNV-Lc small RNAs and five cDNA clones from the de nopo-generated small RNAs (one from plant #2 and four from plant #3; lanes 2 and 3 respectively of Figure 6) were subjected to dideoxynucleotide sequencing using ds plasmid D N A templates, oligonucleotides primers and Sequenase (Figure 7). The complete D N A sequence of seven of the nine recovered clones in comparison to the published C N V sequence (Rochon and Tremaine, 1989) and the sequence of pK2/M5 is given in Figure 8. Two of the recovered clones had sequences identical to those of pDI21 and pDI12 and are consequently not reported in Figure 8. A simplified version of the overall organization of representative DI RNAs relative to the C N V genome is given in Figure 9. Sequence alignments clearly demonstrated that both the small RNAs from CNV-Lc virion R N A and those generated de novo by high moi passaging were linear deletion mutants of C N V genomic R N A ranging in size from 304 to 622 nucleotides. Each DI R N A sequence examined contained sequences derived from the C N V 5' untranslated and 3' terminal regions (regions I and region III, respectively; Figure 9) and a small portion of the open reading frame (ORF) for the putative polymerase (region II; Figure 9). At their 5' ends, all clones maintained sequences up to and immediately upstream of the C N V p33 start codon and seven of the nine clones retained the actual start codon. The largest ORF beginning with the 56 Figure 6. C N V DI RNAs generated de novo from wild-type inoculum. Total nucleic acid was extracted from N. clevelandii leaves after 14 high multiplicity of infection serial passages of an infection established with pK2/M5 transcripts and electrophoresed on a non-denaturing 1% agarose gel containing TAE buffer. Each lane corresponds to total nucleic acid from an independent passage series. Positions of C N V genomic R N A and DI RNAs are shown on the right. Positions of some prominent ribosomal RNAs (rRNAs) are shown on the right: the 25S and 18S rRNAs correspond to the large and small subunits of cytoplasmic rRNAs, respectively; the 16S rRNA corresponds to the small subunit of chloroplast r R N A ; the 14S rRNA is one specific breakdown product of the large subunit of chloroplast rRNA. Photograph provided courtesy of D. M . Rochon. 57 o 33 *• ; 1 I *• * z; ' , « . — « s 1 T 34 33 5 * ^ •< "1 u ^ 34 y 1 ~\ =100 bp Figure 7. Strategy used for sequencing C N V DI R N A cDNA clones. The top strategy was used for the largest pDI clone (pDI15); the bottom strategy was used for the other smaller pDI clones. The D N A template is shown 5' to 3': dark bars represent cloned DI R N A cDNA; stippled bars represent bacteriophage the T7 R N A polymerase promoter; thin lines represent the pUC19 multiple cloning site. Arrows pointing right represent sense oligonucleotide primers; arrows pointing left represent antisense oligonucleotide primers; R=M13 reverse primer and U=M13 -20 primer (universal primer); numbers refer to CNV-specific primers; length of sequence read is indicated by the thin horizontal line terminating with a vertical line after each primer. The sequence of the CNV-specific sequencing primers used can be found i n Table 3 of the Materials and Methods chapter. Scale in base pairs (bp) is indicated at the bottom right. 5 8 Figure 8. Comparison of cloned C N V DI RNA sequences with C N V . Portions of the published C N V sequence or infectious transcript sequence (pK2/M5) not represented in any DI RNAs are indicated within these sequence in parentheses. Note that in order to facilitate the construction of pK2/M5, the 5' and 3' ends of C N V were reamplified from infections established in N. clevelandii (see Rochon and Johnston, 1991). Consequently, sequence differences exist in comparison to the published sequence of C N V given in Rochon and Tremaine, (1989). DI R N A sequences in order of largest to smallest are indicated below the pK2/M5 sequence. Dashes indicate sequence identity with respect to C N V ; nucleotides are shown only when they differ from CNV. Asterisks indicate deletions relative to either C N V or another DI RNA. Underlining indicates the A U G codon which initiates synthesis of p33 and p92. Total lengths of C N V and pK2/M5 genomic R N A and each DI R N A are indicated in brackets at the end of each sequence. Numbers at the beginning of each line correspond to the first nucleotide in each line. In each DI R N A clone, the 5' terminal base is missing and 3' terminal C is added during the cloning procedure (see text). 5 CNV 1 NNAAATTCTCCAGGATTTCTCAACCTTGGTTGTGCTATCTGGTGACTTGCGCGTGTTGTCTGCGTAGAGAATTTCTCTCCTTAACCAAAAGGGGTTTGAAGACTG PK2/M5 1 *G T C * pDI15 1 *G T * pDI12 pDI16 pD19 PDI42 PDI21 pDI48 1 *G-1 *G-1 *G-1 *G-1 *G-1 *G-CNV 106 GGTCTACCGCTTGCGGG * ATAAATTGTAACTTCCAGTAAACGACGACATG.G (1163) TGGACGTCTTTCCCCGTTCAGGAAAGCGGTCTGCGAGAAGGTCGGGG PK2/M5 104 -G A- (1163)-PDI15 104 G-pDI12 PDI16 pD19 pDI42 pDI21 104 104 104 104 104 pDI48 104 -G -G -G -G-C—G--G -G CNV 1365 TTGCCCACCGTTTGGGGTATGATGGATTTCTATCATACTACAGTGGTGCGAAGCTCCGTACTTACACGCGGGCTGTGGAGAGTCTGCATATCAC (2869) PK2/M51364 (2869) pDI15 200 A •***._. T 1 ****** PDI12 201 A ****** pDI16 201 A T ****** pDI9 175 pD142 pDI21 177 177 pDI48 177 CNV 4327 GCCTTCGAAAATAAAGAAAGCGAGTAAGACAGACTCTTCAGTCTGACTTGGTGGAATCTTGCGAATTTAACTGTTACTCTTCATGGGTTCCTTCCCATACGATGA PK2/M54326 282 289 pDI15 PDI12 PDI16 289 pDI9 269 pDI42 266 pDI21 266 pDI48 266 * * * * * * * * CNV 4432 CGAGTCAGGTCGGGCCCTATCTTAGGTTTGGTCACCTAGGGGACGGGGATATGGAAATCACTTTCGCTTGCTGTCAGTCTAGTGGAAACACTTAGCTTGCAATGT PK2/M54431 A G pDI15 362 *** A *****_<; C— pDI12 - ** pDI16 - ** pDI9 - ** PDI42 - ** pDI21 - ** pDI48 - ** * * * * * * * * * * * * * * * * * * * * * * * * * * * * * * CNV 4 537 GGGTGTATGCCTGGATAAGTCGTATGGATGCTGGCCATGATGAATTGGATGCAGTTT* *AGGACGTATAGTGGAAATCTTGCCAGACACGGTTGATCTCACCCTC pK2/M54536 ** pDI15 pDI12 pDI16 pDI9 PDI42 pDI21 pDI4 8 459 368 348 342 314 314 TT CNV 4640 CGGGGGG*CTATAGAGATCGCTGGAA*CACTACCGGACAACCGGAACATTGCAGAAATGCAGCC* (4701) PK2/M54639 G G C (4703) 558 G G C ( 622) 454 G *—G C ( 517) 403 G G C ( 467) 393 G G C ( 459) pDI15 pDI12 pDI16 pDI9 pDI4 2 PDI21 PDI48 371 G G C ( 435) 369 291 -C ( 433) -C ( 304) 60 1 155 1318 1458 V77m II a-4327 4701 Hla/IIIb W77A DI RNA 15 F ^ ^ l DI RNA 12 DI RNA 16 de novo -generated DI RNAs from plant #3 M M ^ DIRNA9 -"SNSSSM DI RNA 21,42 DI R N A 48 ] DI R N A from plant #2 DI RNAs associated with CNV-Lc ' Figure 9. Structure of cloned DI RNAs relative to the C N V genome. Boxes along the C N V genome represent the five confirmed open reading frames of C N V (see Figure 3); hatched blocks immediately beneath the C N V genome represent the largest limits of the three distinct regions derived from C N V by the various DI RNAs. DI RNAs with distinct overall organizations are indicated in the bottom half with broken lines between the hatched blocks indicating deletions within each region. 61 retained C N V p33 start codon (which would remain in the original translational context) encoded 23 amino acids (pDI15). At the 3' end, there was some variability in the sequences retained from the C N V translated and untranslated portions. Only one clone (pDI9) contained some sequence from the 3' portion of the p20 ORF; all clones contained some sequence from the 3' portion of the p21 ORF. One clone retained nearly all of the 3' untranslated region (pDI15) while the remaining clones contained an internal deletion in the 3' untranslated region ranging in size from 310 nucleotides (pDI48) to 120 nucleotides (pDI12). Thus region III can be further delineated into region Ilia (upstream of the internal deletion) and region IHb (downstream of the internal deletion). Region Ilia and region IHb do not have defined borders; rather, there is some variability in the 5' and 3'- borders of region Ilia and in the 5' border of region Illb (see Figures 8 and 9). Clone pDI48 is most likely an artifact of the RT-PCR procedure since this clone lacked most of region III yet retained the last 15 nucleotides exactly complementary to the 3' oligonucleotide used for RT-PCR. There were very few substitutions in DI R N A sequences relative to C N V genomic R N A or pK2/M5 R N A (see Figure 8) and none of the DI R N A s sequenced contained internal duplications of sequences as has been observed with TBSV DI RNAs (Hillman et al, 1987) and C N V DI RNAs sequenced in another laboratory (Chang et al, 1995). Biological activity of transcripts derived from cloned DI RNAs Each of the DI RNAs in Figure 8 was tested on N. clevelandii hosts for their ability to accumulate and/or attenuate symptoms. Plants were infected with equal amounts (-0.5 ug) of T7 R N A polymerase-derived transcripts of each DI RNA cDNA clone alone or coinfected with T7 R N A polymerase-derived transcripts of pK2/M5 (-5 ug). Plants were monitored for symptom attenuation for up to three weeks (by which time controls infected with WT transcripts alone had desiccated and died; see example in Figure 10C). Six days post-inoculation total leaf R N A was extracted from 62 one originally infected leaf and subjected to Northern blot analysis using 3 2P-labeled nick-translated probes. Probes corresponded to the 5'-terminal 40 nucleotides (5' probe) and the 3'-terminal 540 nucleotides (3' probe) of C N V genomic RNA. A l l of the transcripts generated from the DI R N A cDNA clones were able to attenuate symptoms when coinfected with WT transcripts with the exception of pDI48. Plants coinfected with transcripts, from this clone displayed severe necrotic symptoms similar to those generated in WT transcript-infected plants. A typical example of the symptom attenuation observed in the presence of DI R N A transcripts derived from DI R N A cDNA clones is shown in Figure 10. A l l plants which were infected only with transcripts generated from DI R N A cDNA clones appeared as mock-infected controls (not shown). Northern blot analysis was used to determine if DI R N A transcripts were capable of accumulating in plants. Figure 11, lanes 3 through 9, show that in each coinfection experiment, it was possible to detect with both 5'- and 3'-specific probes a R N A species corresponding to the molecular weight of the DI R N A transcript used for coinfection. As expected, the synthetic DI R N A transcripts did not accumulate in infections performed in the absence of helper virus (Figure 11, lanes 10 through 16). R N A species which were estimated to be twice the size (dimer-sized) of the DI R N A transcript used in the coinfections and were not detected in WT-infected plants also hybridized to the 5' and 3'-specific C N V probes used and, in some cases, were the main R N A species detected. The identity and characterization of these dimer-sized RNAs constitute the second chapter of this thesis. Certain coinfections were observed to accumulate a large amount of dimer-sized R N A (e.g. DI R N A 9; Figure 11, lane 6) while other DI R N A coinfections accumulated an amount detectable only by Northern blot analysis (e.g. DI R N A 42; Figure 11, lane 7). This phenomenon is also explored in the second chapter of this thesis. Throughout the course of the experiments in this thesis, coinfections with these DI RNAs were repeated at least three times and numerous 63 Figure 10. Symptom attenuation caused by C N V DI RNAs in Nicotiana clevelandii. Panels A, B, and C: N. clevelandii infected with WT C N V at 1, 2, and 3 weeks post inoculation respectively. Panels D, E, and F: N. clevelandii coinfected with W T C N V and DI R N A 42 at 1, 2, and 3 weeks post inoculation respectively. 64 1 2 3 4 5 6 7 8 9 10 1112 13 14 15 16 17 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 Figure 11. Northern blot analysis of coinfected Nicotiana clevelandii. Leaf total nucleic acid was extracted from plants six days post inoculation. R N A was denatured and electrophoresed through 1% agarose gels containing 5 mM methyl mercuric hydroxide and transferred to a membrane The membrane was hybridized to a nick-translated probe. At the top is a membrane which was probed with a 5' specific probe; below is the same membrane which was probed with a 3' specific probe. Lane 1 contains ~1 ug of total nucleic acids from a plant infected with W T transcripts; lanes 2 through 8 contains ~1 ug of total nucleic acids from plants coinfected with WT transcripts and transcripts from pDIs 15, 12, 16, 9, 42, 21, and 48 respectively. Lane 9 contains ~1 ug total nucleic acid from a mock-infected plant and lanes 10 through 16 contains ~1 ug total nucleic acids from plants infected singly with transcripts from pDIs 15,12, 16, 9, 42, 21, and 48 respectively. Lane 17 contains 0.1 ug purified CNV-Lc virion RNA. Sizes given in kb on the left of each blot correspond from top to bottom to full length C N V genomic R N A (which was evident in all extracts in which WT transcripts had been used for inoculation upon longer exposure) as well as the coat protein subgenomic R N A and the bifunctional subgenomic R N A (both detectable by the 3' probe only). Arrows distinguish the R N A species corresponding to the molecular weight of the DI R N A transcript used for infection; brackets outline the dimer-sized R N A species generated during coinfection. 6 5 times in the case of DI RNAs 9, 21, and 42. The R N A accumulation patterns shown in Figure 11 were reproducible and are therefore a hallmark of the particular DI R N A used in the coinfection. The overall levels of R N A accumulation were also reproducible and reflect differences in the ability of individual DI RNAs to accumulate. The results described above confirm that transcripts derived from the C N V DI R N A cDNA clones, with the exception of pDI48, are biologically active. Involvement of coat protein in DI RNA accumulation It has been demonstrated previously that a C N V mutant in which most of the coding region for the viral coat protein had been removed [CP(-); McLean et ah, (1993)] accumulates and spreads efficiently in absence of coat protein (McLean et ah, 1993). In addition, studies carried out in the laboratory by D. M . Rochon demonstrated that C N V R N A extracted from virions purified from coinfected plants contained proportionately much less DI R N A than found in total nucleic acid extracted from coinfected leaves. This indicated that C N V DI RNAs were not efficiently encapsidated. The involvement of the C N V coat protein in DI R N A accumulation was further investigated by coinfecting DI R N A transcripts with transcripts from CP(-). DI RNAs were found to accumulate just as well using CP(-) as helper virus as they do using WT helper virus (Figure 12), indicating that coat protein is not required for DI R N A accumulation. Plants coinfected with DI R N A transcripts and CP(-) helper transcripts also showed attenuated symptoms indistinguishable from those of coinfections with WT helper. 66 genomic R N A CP(-) R N A DI R N A Figure 12. Analysis of DI R N A accumulation in DI R N A coinfections using wild-type C N V or CP(-) transcripts as helper. Total nucleic acid was extracted from Nicotiana clevelandii leaves five days post inoculation and electrophoresed through a 1% non-denaturing agarose gel containing TBE buffer. Lane 1 contains extract from a mock-infected plant, lane 2 from a plant infected with WT transcripts (the genomic R N A band was visible on the original photograph of the gel but is only barely visible in this reproduction) and lane 3 from a plant infected with CP(-) synthetic transcripts. Lanes 4, 6 and 8 contain extracts from plants coinfected with WT C N V R N A and DI RNAs 42, 9 or 15, respectively. Lanes 5, 7 and 9 were coinfected with CP(-) R N A and DI RNAs 42, 9 or 15, respectively. Positions of WT, CP(-) and DI RNAs are shown on the right. The prominent rRNA bands seen in all samples are defined in Figure 6. 67 Discussion This study confirms that CNV-Lc carries DI RNAs. In addition, it confirms that the small RNAs which spontaneously arise during serial high moi passages of WT C N V infections (Rochon, 1991) are in fact DI RNAs. These DI RNAs fit the classical criteria of a DI in that they contain viral-derived sequences, they are able to accumulate only in the presence of helper virus and they interfere with helper virus accumulation as indicated by the dramatic symptom attenuation seen i n coinfections. DI RNAs accumulate to very high levels in coinfected N. clevelandii. They are often more abundant than even ribosomal RNAs and can constitute as much as 50% of the nucleic acid in plant total nucleic acid extracts (for example DI RNA 42 coinfections, see Figure 12, lanes 4 and 5). The DI RNAs associated with C N V are a mosaic of three or four non-contiguous portions of the viral genome. These retained sequence blocks must contain essential cz's-acting sequences required for replication as the C N V DI R N A s characterized in this study are able to accumulate in the presence of helper virus. The same overall structure was found in C N V DI RNAs sequenced independently in another laboratory (Chang et al., 1995) and in DI RNAs associated with two other tombusviruses (Hillman, 1987; Burgyan et al, 1989). A n extensive deletion analysis performed on C N V DI R N A clones constructed in another laboratory, has further defined sequences within the retained sequence blocks which are either dispensable or crucial for accumulation (Chang et al, 1995). Similar analyses have also been carried out on the DI RNAs associated with TBSV (Chang et al, 1995) and CyRSV (Havelda et al, 1995). The general location of crucial versus dispensable sequences (in terms of replication) within each of these retained sequence blocks is similar in the case of all three tombusvirus DI RNAs with one exception. Deletion of the 5' portion of the 3'-terminal region (designated CI in the case of CyRSV DI RNAs; III in the case of TBSV DI RNAs; and III or Ilia in the case 68 of C N V DI RNAs) resulted in loss of ability to be replicated in the case of CyRSV Dl RNAs. In the case of TBSV and C N V DI RNAs, deletion of this same region did not affect the ability of the DI R N A to be replicated. The reason for this apparent contradiction remains to be resolved. Of particular interest in the studies of Chang et al, (1995) was the deletion analysis of region II. Region II was found to be necessary for accumulation of C N V DI RNAs; this requirement is also shared by both TBSV and CyRSV DI RNAs (Chang et al, 1995; Havelda et al, 1995). Limited deletions within the center of this region could be tolerated. This region is predicted to form a stem loop structure in both C N V (Chang et al, 1995, own unpublished observations) and TBSV DI RNAs (Chang et al, 1995) with the stem comprising those sequences which are crucial for accumulation and the loop containing the dispensable sequences. A detailed study involving serial passaging of TBSV DI RNAs in cucumber protoplasts demonstrated that more highly evolved DI R N A s contained a duplication of region II (White and Morris, 1994a; White and Morris, 1994c). Increased replication competence in DI RNAs with duplications of region II allowed them to outcompete DI RNAs lacking this duplication. The necessity of part of this region for accumulation and its duplication in DI RNAs with enhanced replication competence implicate region II as a ds-acting element recognized by the viral replication complex. Whether this is a sequence-specific element or a structure-specific element remains to be determined. One clone, pDI48, was recovered which was not able to be replicated by helper virus and did not attenuate symptom formation. Although examination of the sequence of this clone suggests it was likely derived as a cloning artifact, it still shows that there are sequences in region III corresponding to between nucleotides 4380 and 4400 and/or 4587 and 4688 in the C N V sequence which are absolutely required for C N V DI R N A to efficiently accumulate in the presence of helper virus. In the aforementioned study on C N V DI R N A cz's-acting sequences, deletion of the 3' terminus from nucleotide 4586 to 4701 was found to be detrimental to DI R N A 69 accumulation. Deletions which encompassed the sequences in region III between nucleotides 4380 and 4400 did not affect accumulation. Consequently, the lack of nucleotides 4587 through 4688 appears to be responsible for the inability of transcripts from pDI48 to accumulate. This 101 nucleotide stretch, therefore, contains important cz's-acting sequences for C N V replication. In addition, the last 13 nucleotides of C N V (nucleotides 4689 through 4701) are insufficient to promote synthesis of (-) RNA. The de novo generated DI RNAs have fewer deletions relative to the laboratory culture DI RNAs in region III suggesting that smaller DI RNAs may arise from larger precursors during the course of DI R N A evolution. The evolution of smaller DI RNAs from larger precursors has also been suggested in the case of CyRSV (Burgyan et al, 1991). In the previously mentioned study on TBSV DI R N A evolution, it was found that larger DI RNAs can indeed evolve into smaller ones, with further deletions occurring in region III (White and Morris, 1994a): Comparison of DI RNAs generated de novo from WT clones and those generated from C N V p20 mutant infections (Rochon, 1991) show that these DI RNAs are highly similar in sequence and overall primary structure. This suggests that the rapid appearance of DI RNAs in p20 mutant infections is not due to gross peculiarities in the DI R N A species generated which render them better able to accumulate. Instead, the C N V p20 protein may normally be involved in the C N V replication complex, possibly to aid in the resolution of secondary structure i n template R N A (Rochon, 1991). Absence of this protein may result in a replication complex which is more likely to undergo the dissociation events implicated in DI R N A formation, leading to the appearance of DI RNAs after just one high moi passage. A similar phenomenon is seen in the case of influenza virus where mutations in the NS2 non-structural protein resulted in DI R N A generation after a single high moi passage (Odagiri and Tobita, 1990). A role for the NS2 protein i n R N A replication was also proposed based on these observations. 70 The strategy used for the construction of cDNA clones of C N V DI R N A s employed CNV-specific primers and, consequently, selected only those molecules with extreme 5' and 3' ends resembling those of C N V . The intent of this part of the study was to establish a set of cDNA clones of C N V DI RNAs as opposed to extensively screening for all the various DI RNAs which may exist in the population. It is possible that C N V DI RNAs with different 5' and/or 3' ends existed in the source templates used for cloning but were overlooked by this cloning procedure. Precedents exist for DI RNAs with a 5' end distinct from that of the helper virus. Sindbis virus DI RNAs with cellular t R N A a s P sequences at the 5' end have been described (Monroe and Schlesinger, 1983) and a turnip crinkle virus (TCV) DI R N A with a 5' end resembling a TCV satellite R N A has also been described (Li et al, 1989). Unlike the definition of a DI R N A as outlined in the Introduction, C N V DI RNAs do not require viral coat protein for accumulation. This is reflective of the fact that many plant viruses, C N V included, do not require the viral coat protein i n order to accumulate and spread from cell-to-cell in an infected plant (Stanley and Townsend, 1986, Dawson et al, 1988; Gardiner et al, 1988; Lazarowitz et al, 1989; Petty and Jackson, 1990; Dalmay et al, 1992; McLean et al, 1993; Scholthof et al, 1993; Xiong et al, 1993). The DI RNAs associated with both TBSV and CyRSV also do not appear to be efficiently encapsidated (Hillman et al, 1987; Burgyan et al, 1992) and TBSV DI RNAs also accumulate to high levels in the absence of coat protein (White and Morris, 1994a). The observation that C N V DI R N A accumulation occurs efficiently in the absence of coat protein also suggests that factors other than encapsidation or interaction with the C N V coat protein contribute to the stability of these molecules in planta. Several different factors could be involved including inherent secondary or tertiary structure of the DI RNA, the ability of DI RNAs to bind ribosomes, and/or interaction of DI RNAs with host- or virus-encoded proteins. C N V DI RNAs have 7 1 the potential to fold into highly structured molecules with free energy values ranging from -132.3 to -192.8 kcal/mol (unpublished observations). Also, several C N V DI RNAs retain the A U G codon that initiates synthesis of the C N V 5'-proximal ORF raising the likelihood that C N V DI RNAs can bind ribosomes. In the case of defective R N A associated with clover yellow mosaic virus, the process of translation appears to contribute to stability in plants (White et al, 1992). This is also the case for mouse hepatitis coronavirus DI RNAs (de Groot et al, 1992; van der Most, et al, 1995). C N V p21 has been shown to be the viral movement protein (Rochon and Johnston, 1991; Johnston, 1995). As other movement proteins have been demonstrated to be ss nucleic acid binding proteins (Citovsky et al, 1990), it is possible that the C N V p21 is also a ss nucleic acid binding protein which could act to stabilize DI RNAs. Host proteins are known to be integral components of the replicase complexes of (+) ss R N A viruses (Blumenthal and Carmichael, 1979; Quadt and Jaspars, 1990; Kajitani and Ishihama, 1991; Quadt et al, 1993), so it is reasonable to suggest the involvement of host proteins in DI R N A stability instead of or i n addition to viral proteins. Competition for encapsidation proteins by DIs has been suggested as one means for their interfering capability (Perrault, 1981; Holland, 1991; Roux et al, 1991). The observation that C N V DI RNAs are still able to attenuate symptom development in the absence of coat protein suggests that competition for coat protein does not contribute significantly to the interference effect. 72 CHAPTER 2. Identification and characterization of C N V DI RNA dimers Dimer-sized RNAs were observed in coinfections of C N V DI R N A transcripts and WT transcripts generated from pK2/M5. Their presence in all coinfections (with the exception of pDI48 transcripts which were unable to replicate) suggested they could be intermediates of DI R N A replication. The level of accumulation of dimer-sized RNA in coinfections also seemed to depend on the DI R N A used in the coinfection. Elucidation of the nature of these RNAs and investigations into the influence of primary sequence on the accumulation of these RNAs were therefore carried out in order to gain insights into how they are generated and whether or not these structures are relevant in terms of DI R N A replication. Dimer-sized forms of tombusvirus DI RNAs had not been described nor investigated in any manner prior to the work reported in this chapter. Results Analysis of overall structure and size of dimer-sized RNAs Dimer-sized R N A species were not observed in blots where synthetic transcripts alone were electrophoresed demonstrating that they were not migration artifacts due to insufficient denaturation (data not shown). In addition, they did not represent ds forms of the DI R N A as control experiments demonstrated that the electrophoresis conditions used were sufficient to denature ds C N V R N A (data not shown). Total nucleic acid extracts from coinfections with prominent dimer-sized RNAs (coinfections with DI R N A 9 and DI R N A 12) were denatured with glyoxal and dimethyl sulfoxide (Sambrook et ah, 1989) and electrophoresed on the same agarose gel as non-denatured samples in a sodium phosphate buffer system at p H 7.0. Under such conditions, the denaturation of nucleic acids by glyoxalation is irreversible; consequently, denatured and non-denatured samples can be compared 73 on the same agarose gel (McMaster and Carmichael, 1977). After electrophoresis, the gel was stained with acridine orange to distinguish double-stranded (ds) nucleic acids from ss nucleic acids. Under ultraviolet illumination, acridine-stained ds nucleic acids wil l fluoresce green while acridine-stained ss nucleic acids (including highly structured ribosomal RNAs) wil l fluoresce red (McMaster and Carmichael, 1977). Dimer-sized RNAs fluoresced red under both denaturing and non-denaturing conditions (not shown), suggesting that the dimer-sized RNAs were not copy back RNAs (see Figure 1) which would be expected to fluoresce red under denaturing conditions and green under nondenaturing conditions as they are predominately ds molecules. Instead, these RNAs appeared to be ss with a molecular weight twice that of the DI R N A used for coinfection. To confirm the molecular weight of the dimer-sized RNAs found in DI R N A 42 and DI R N A 9 coinfections, total nucleic acid extracted from leaves coinfected with WT transcripts and either of these two DI RNAs were analyzed following denaturing agarose gel electrophoresis and Northern blotting. Figure 13 shows that the dimer-sized R N A band observed in DI R N A 42 coinfections (lane 3) comigrates with synthetic transcripts derived from a constructed cDNA clone corresponding to a head-to tail repeat of DI R N A 42 sequence (lane 5). In addition, the prominent DI R N A 9 dimer-sized R N A band (lane 6), which is predicted to be 918 nucleotides i n length, migrated at a position above the 872 nucleotide synthetic dimer transcript (lane 5) and very similar to the 931 nucleotide synthetic transcript included as a size marker (lane 7). Therefore, dimer-sized RNAs found in both DI R N A 42 and 9 coinfections have electrophoretic migration patterns which are consistent with them being duplications of the complete monomeric-length RNA. They w i l l consequently be referred to as dimers or dimer RNAs for the remainder of this thesis. 1 2 3 4 5 6 7 8 -genomic R N A DI R N A dimers -^j. DI R N A monomers Figure 13. Northern blot analysis of dimer-sized RNAs generated during coinfections. RNAs were denatured with methylmercuric hydroxide and electrophoresed through a 1.2% agarose gel containing 5 m M methylmercuric hydroxide and transferred to a membrane. The membrane was probed with a nick-translated probe corresponding to C N V nucleotides 21-68. Lanes 1 through 4 correspond to total nucleic acid extracts prepared five days post inoculation from plants inoculated with: (1) buffer (mock-infection); (2) WT transcripts; (3) W T transcripts plus DI R N A 42 monomer transcripts (435 nucleotides); and (4) W T transcripts plus DI R N A 42 dimer transcripts. Lane 6 corresponds to a total nucleic acid extract prepared six days post inoculation from plants infected with W T transcripts plus DI R N A 9 monomer transcripts (459 nucleotides). Lanes 5 and 7 correspond respectively to gel-purified synthetic DI R N A 42 dimer transcripts (872 nucleotides) and transcripts prepared from HmcII-linearized pK2/M5 (931 nucleotides), included as molecular size markers. Both of these size markers were mixed with total nucleic acid extract from mock-infected plants prior to denaturation and electrophoresis. Lane 8 corresponds to 2 M LiCl-insoluble nucleic acids prepared from the 14th passage of an infection established with infectious transcripts which had generated DI RNAs de novo (see also Figure 6, lane 2). Note that smaller RNAs, including monomeric-length DI R N A 9, are underrepresented in this sample because they fractionate primarily into the 2 M LiCl-soluble material (Hillman et al, 1987). 75 Analysis of the cDNA sequence generated from dimer RNAs One structure which would be consistent with the properties of the dimer RNAs established in the preceding experiments was a linear, head-to-tail repeat of DI R N A sequence. Dideoxynucleotide sequencing of dimer R N A using reverse transcriptase and an oligonucleotide complementary to C N V nucleotides 93 through 108 (CNV 1; all oligonucleotides referred to in this chapter can be found i n Table 3 in the Materials and Methods chapter) was carried out in an effort to determine if the dimer RNAs were in fact linear, head-to-tail repeats of DI R N A sequence. Dimer RNAs were purified from a non-denaturing agarose gel of total nucleic acid extracted from leaves coinfected with DI R N A 9, which produces high levels of dimer RNAs. If the dimer RNAs were linear, (+), head-to-tail dimers the sequence should be complementary to the 5' end of DI R N A 9, followed by termination in all four lanes (stop), followed by sequence complementary to the 3' end of DI R N A 9. Figure 14A shows that this was the sequence pattern obtained using DI R N A 9 dimers as templates. The sequence beyond the stop appeared to be the complement of the 3' end of DI R N A 9; a more definitive demonstration of this was accomplished using RT-PCR (see next section). Control sequencing reactions using the same primer on gel-purified monomer-sized R N A templates from DI R N A 42 coinfections (Figure 14A and 14B) and C N V virion R N A templates (Figure 14B) yielded only 5'-complementary sequence and a stop but not 3'-complementary sequence. Similar results were also seen using an oligonucleotide complementary to C N V nucleotides 51 through 66 (CNV 38) in these reactions (data not shown). These R N A sequencing results, along with the size of the RNAs determined as described above, are consistent with linear, head-to-tail repeats of DI R N A sequence. 7 6 Figure 14. Analysis of cDNA sequence generated from dimeric- and monomeric-length DI RNA. Both sets of reactions used an oligonucleotide primer complementary to C N V nucleotides 93 to 108 (CNV 1). Dimeric- and monomeric-length DI R N A templates were gel-purified from total leaf nucleic acid extracts obtained from DI R N A 9 or DI R N A 42 coinfections, respectively; genomic-length C N V R N A was prepared from virions (Rochon and Johnston, 1991). Lanes marked O denote reactions without any added dideoxynucleotides, whereas lanes marked A , C, G or T correspond to the dideoxynucleotide used in the chain termination reactions. Panel A shows reactions with monomeric- and dimeric-length DI R N A templates. Panel B shows shorter exposures of an independent reaction with monomeric-length DI R N A and also genomic-length C N V RNA. 77 » - H H Z Z 78 Analysis of dimer junction sequences The exact junction sequence between monomer units was obscured by the strong stop in the R N A sequencing reactions (see Figure 14A) and even upon shorter exposures of reactions with DI R N A monomer template (see Figure 14B) the identity of the 5' base could not be determined unambiguously. In order to determine the junction sequence and definitively demonstrate the presence of 3' complementary sequence, an RT-PCR strategy, outlined at the top of Figure 15, was developed utilizing primers which would amplify a ds D N A product specifically from (+) dimer RNAs. RT-PCR products were amplified from total nucleic acid extracts prepared from leaves of coinfected plants, cloned and then the sequences surrounding the junction sites were determined. RT-PCR products amplified from DI R N A 9 coinfections and DI R N A 42 coinfections were both sequenced and a summary of the sequences obtained from each DI R N A dimer is shown in the lower half of Figure 15. Approximately half of the cloned sequences from both DI R N A 42 and 9 dimers contained the sequence 5 ' . . .CCCAGAAA.. .3 ' (see Figure 15) at the junction site. This sequence also represents the consensus junction sequence obtained when the RT-PCR products of these dimers were sequenced as a population. The remaining junction sequences consisted of either deletions or insertions relative to this consensus junction sequence. With the exception of one clone, these alterations were all found on the right hand side of the junction. The 5' and 3' terminal sequences of DI R N A 42 as deduced from the cDNA clone sequence are indicated in Figure 15. The N corresponds to an unidentified extra 5' terminal base which was revealed from a shorter autoradiographic exposure of R N A sequencing reactions using both DI R N A 42 monomer template and C N V vi r ion RNA template shown (Figure 14B). This unidentified base probably corresponds to the A (in boldface) present in the consensus junction sequence described above. It would therefore appear that about 50% of the dimer junction sequences correspond 79 5" 3' I reverse Y transcriptase ^ Taq polymerase RT-PCR product DI RNA 42: 3' 5" . CAGAAATGCAGCCCNGAAATTCTCCAGGATTTCTCAACC. . CAGAAATGCAGCCCAGAAATTCTCCAGGATTTCTCAACC. . CAGAAATGCAGCCC-GAAATTCTCCAGGATTTCTCAACC. . CAGAAATGCAGCC-AGAAATTCTCCAGGATTTCTCAACC. . CAGAAATGCAGCCCGAGAAATTCTCCAGGATTTCTCAACC (4/9) (1/9) (1/9) (3/9) DIRNA9: 3' 5' CAGAAATGCAGCCCNGAAATTCTCCAGGATTTCTCAACC... . CAGAAATGCAGCCCAGAAATTCTCCAGGATTTCTCAACC. . . (6/12) . CAGAAATGCAGCCCGAGAAATTCTCCAGGATTTCTCAACC. . (2/12) , CAGAAATGCAGCCCAGAGAAATTCTCCAGGATTTCTCAACC. (1/12) . CAGAAATGCAGCCC CC . . . (3/12) Figure 15. Sequence analysis of dimer R N A junctions. The upper portion depicts the RT-PCR strategy designed to amplify ds D N A encompassing dimer junctions. Dimer R N A is shown schematically using adjoining arrows. The short arrows correspond to the approximate location of the oligonucleotide primers used. C N V 1 was used to generate first-strand complementary D N A and subsequent complementary strands in the PCR; C N V 33 was used to generate second-strand D N A and subsequent sense strands. The lower portion depicts cloned junction sequences derived from dimers of DI RNAs 42 and 9. The top sequence in each alignment represents a precise fusion between the 3' and 5' ends of DI R N A 42 monomer based on the sequence of in vitro transcripts derived from Smal-linearized templates and shorter exposures of the monomer R N A sequencing reaction depicted in Figure 14B. The numbers in parentheses refer to the occurrence of each sequence. Nucleotides in boldface are not present in the in vitro transcripts and dashes indicate nucleotides which are deleted in comparison to the precise fusion. 80 to a precise fusion of monomer units while the remainder contain insertions or deletions of a few or several nucleotides. Biological activity of synthetic dimer transcripts It is possible that the dimer RNAs generated during DI R N A coinfections i n plants represent obligatory intermediates in the formation of monomers. Alternatively, the dimer RNAs may represent byproducts of replication which have no further specific role in the DI R N A replication process. A cDNA clone corresponding to a head-to-tail repeat of DI R N A 42 sequence and from which T7 R N A polymerase-derived synthetic DI R N A 42 dimers could be prepared was constructed to assess the possibility that dimers act as intermediates in the formation of monomers during C N V DI RNA replication. C N V 39 and C N V 32 were used as primers in a PCR reaction to amplify the complete monomeric-length sequence from pDI42. C N V 39 is a mutagenic primer which introduces an Xbal site immediately upstream of the 5' end of the DI R N A 42 sequence. The Xbal site was included in the primer to enable excision of PCR products from the pT7Blue intermediate vector used to facilitate the cloning procedure. C N V 32 corresponds to the complement of the 3' terminus of DI R N A 42 and contains a Smal site used to linearize the template for run-off transcription. The Xbal/Smal fragment encompassing DI R N A 42 sequence was excised from pT7Blue, treated with mung bean nuclease and then ligated into Smal-digested pDI42. Sequencing of AccI restriction fragments from two dimer clones revealed that the A in the center of the dimer junction was missing in both clones, probably as a result of mung bean nuclease treatment (Hammond and D'Allessio, 1986). T7 R N A polymerase-derived synthetic dimer transcripts generated from one of these dimer cDNA clones (pDI42-Dl) were coinfected with synthetic W T transcripts onto N. clevelandii. Total nucleic acid extracts prepared from leaves five days post inoculation revealed that the corresponding DI R N A 42 monomer had 8 1 accumulated along with a small amount of dimer R N A (Figure 13, lane 4). This R N A accumulation profile was similar to the profile obtained when DI R N A 42 monomer transcripts were used for coinfection (Figure 13, lane 3). Analysis of transcription products yielded from the dimer clone by Northern blotting failed to reveal evidence of monomeric-length R N A (data not shown) suggesting that small amounts of monomeric-length RNA, generated by incomplete transcription of dimers and/or degradation of dimer transcripts, are not responsible for the accumulation of monomers in coinfected plants. In an effort to rule out the presence of undetectable amounts of monomeric-length R N A in the inoculation mix, dimer transcripts were diluted 10 fold and 100 fold prior to coinfection. Dimer transcripts were still highly infectious, yielding monomers even when diluted 100 fold; this was also the case for similarly-diluted monomer transcripts (Figure 16). The observation that synthetic dimer transcripts can be used as a template for the formation of monomers in coinfected plants might suggest that dimers represent obligate intermediates in the formation of monomers during DI R N A replication in plants. Alternatively, the role of the dimer in DI R N A replication may not be specific. For example, the initiation of DI R N A replication may be somewhat insensitive to 3' terminal extensions on the template molecule enabling the efficient generation of a monomer from a synthetic dimer template. Biological activity of monomers with 3' terminal extensions To investigate the effects of 3' terminal extensions on DI R N A accumulation, coinfections were performed with transcripts generated from Sspl-linearized pDI42 (Ssp transcripts). This introduces an extra 597 plasmid-derived, non-DI R N A nucleotides onto the 3' end of DI R N A 42 monomer transcripts (see Figure 17A). Analysis of total nucleic acid extracts from coinfections with Ssp transcripts (Figure 17B) revealed that monomers equivalent in size to DI R N A 42 monomers accumulated. This suggests that an internally-located initiation site for synthesis of 82 1 2 3 4 5 6 7 8 Figure 16. DI R N A monomer accumulation in coinfections with dilutions of monomeric and dimeric-length DI R N A 42 transcripts. DI R N A transcripts (approximately 0.5 ug) were diluted in in vitro transcription mix minus D N A template and T7 RNA polymerase (see Materials and Methods chapter for composition of this mix); undiluted and diluted DI R N A transcripts were all combined with the same amount of WT transcripts (approximately 5 ug)and used to inoculate N. clevelandii. Total nucleic acid was extracted six days post inoculation and electrophoresed through a 1% non-denaturing agarose gel containing TBE buffer. Lane 1 contains extract from a WT-infected plant; lanes 2, 3, and 4 contain extracts from coinfections with undiluted monomeric-length DI R N A 42 transcripts, l:10-diluted transcripts or l:100-diluted transcripts, respectively; lanes 5, 6, and 7 contain extracts from coinfections with undiluted dimeric-length DI R N A 42 transcripts, l:10-diluted transcripts or l:100-diluted transcripts, respectively; lane 8 contains extract from a mock-infected plant. Positions of C N V genomic R N A and DI R N A 42 monomer are shown on the right. The prominent rRNA bands seen i n all samples are defined in Figure 6. 83 A Smal I II Ilia Illb 5* I II Ilia Illb I (597 nts) 3' genomic R N A DI R N A monomer Figure 17. Monomer accumulation in coinfections with transcripts from Sspl-linearized pDI42. A . Diagrammatic representation of R N A transcripts generated from Smal- and Sspl-linearized pDI42 templates: boxes represent DI R N A 42 sequence; dashed line represent sequence transcribed from pUC19. Restriction endonuclease used to linearize plasmid templates for in vitro transcription are indicated above each transcript. B. Biological activity of Ssp transcripts i n coinfections. Coinfections were carried out with either DI RNA 42 transcripts or Ssp transcripts. Total leaf nucleic acid was extracted six days post inoculation and electrophoresed through a 1% non-denaturing agarose gel containing TBE buffer. Lane 1 contains extract from a WT-infected plant; lane 2 contains extract from a coinfection with DI R N A 42 transcripts; lane 3 contains extract from a coinfection with Ssp transcripts; lane 4 contains extract from a mock-infected plant. Positions of C N V genomic R N A and DI R N A 42 monomer are shown on the right. The prominent rRNA bands seen in all samples are defined in Figure 6. 84 (-) R N A can be recognized by the C N V replication complex. Removal of the 3' terminal extension by degradation of Ssp transcripts, however, cannot be unequivocally ruled out as a possibility in these experiments. Therefore, internal initiation on (+) dimer R N A template is at least one mechanism by which DI R N A monomers can be generated from dimers. Analysis of dimer accumulation in chimeric and mutated DI RNAs DI R N A 9 and DI R N A 21 are highly related in terms of primary sequence and size (Figure 18) yet they consistently show dramatically different R N A accumulation patterns; DI R N A 9 ^ accumulates primarily as a dimer whereas DI R N A 21 accumulates primarily as a monomer (see Figures 11 and 13). The overall level of DI R N A accumulation is also greater in DI R N A 9 coinfections as compared to DI R N A 21 coinfections. To begin investigating the role of primary sequence in the ability of a DI RNA to accumulate as dimers, chimeric and mutated DI R N A c D N A clones were made between pDI9 and pDI21 (see Figure 18 and Figure 19). DI R N A accumulation patterns of at least three independent coinfections with in vitro transcripts derived from each chimeric and mutated DI R N A cDNA clone were examined. A representative example of these DI R N A accumulation patterns is shown in Figure 20. The limitations of the in vivo assay system used for these investigations do not allow the results to be interpreted in a precisely quantitative manner. However, as pointed out in the preceding chapter, both the pattern and overall level of DI R N A accumulation in coinfections have been found to be very reproducible. Consequently, qualitative differences in the pattern and overall level of DI R N A accumulation in coinfections with chimeric and mutated DI RNAs have been interpreted in relation to the primary sequence of DI RNAs. In the first set of chimeric DI R N A cDNA clones, PflMl/EcoRI fragments from pDI9 and pDI21 were exchanged to create pDI9/21PE and pDI21/9PE (Figure 19). The 85 pDI9 pDI21 PDISS9 PDISS9/21TE PDILS21 PDILS21/9TE PDILS21/ATAAT GAAATTCTCCAGGATTTCTCAACCTTGGTTGTGTTATCTGGTGACTTGCGCGTGTTGTCTGCGTAGAGAATTTCTCTCCTTGACCAAAGGGGTTTGAAGAC pDI9 pDI21 PDISS9 PDISS9/21TE PDILS21 PDILS21/9TE pDILS 21 / ATAAT TGGGTCTACCGCTTGCGGGGATAAATTGTAACTTCCAGTAAACGACGACAT**AGCGGTCTGCGAGAAGGTCGGGGTTGCCCACCGTTTGGGGTATGATGG pDI9 pDI21 PDISS9 PDISS9/21TE PDILS21 PDILS21/9TE PDILS21/ATAAT I I Ia-» ATTTCTATCATACTACAGTGGTGCGAAGCTCCGTACTTACACGCGGGCTGTGGAGAGTCTGCATATCACGCCCTCAAAATAAAGAAAGCGAGTAAGACAGA -G*-A-** -G*-A-*» pDI9 pDI21 pDISS9 PDISS9/21TE PDILS21 pDILS21/9TE PDILS21/ATAAT CTCTTCAGTCTGACTTGGTGGAATCTTGCGAATTTAACTGT * * * *GCAGTTTAGGACGTACAGTGGAAATCTTGCCAGACACGGTTGATCTCACCCTCCGG : TAAT T TAAT T TAAT T •• pDI9 GGGGGCTATAGAGATCGCTGGAAGCACTACCGGACAACCGGAACATTGCAGAAATGCAGCCC (459) PDI21 A — (433) PDISS9 • — (431) PDISS9/21TE A --. (435) PDILS21 A (462) PDILS21/9TE (458) pDILS 21 / ATAAT A (458) Figure 18. Comparison of D N A sequences of DI R N A 9, DI R N A 21 and mutated DI RNAs. Dashes indicate sequence identity with the DI R N A 9; nucleotide differences are shown only when they differ; italicized nucleotides represent major sequence differences; asterisks indicate deletions. Restriction endonuclease sites used in the construction of chimeric DI RNAs and the beginning of each distinct region of the DI RNAs (nucleotide immediately beneath the arrow represents the first nucleotide of each region) are indicated above the sequences. Total length of each DI R N A in nucleotides is indicated in brackets at the end of each sequence. 8 6 Figure 19. Chimeric and mutated C N V DI RNAs. Dark boxes represent sequence derived from pDI9; white boxes represent sequence derived from pDI21; deleted sequence is indicated by a"V" between sequence blocks; dark horizontal bars within pDI21 and pDI9 sequence blocks indicate the positions of the major primary sequence difference between these DI RNAs; dashed line indicates sequence found 3' to the DI R N A sequence. Restriction endonuclease sites used in the construction of the depicted chimeric DI RNAs are indicated; the EcoRI site used is from the multiple cloning site of the plasmid and is found 3' to the DI R N A sequence. 87 PARENTS pDI21 I n Ilia • Illb . . . . t T L-pDI9 PflMl Tfil EcoRI 11 — M a Illb CHIMERICS -pDI9/21PE pDI21/9PE pDI9/21PT pDI21/9PT pDI9/21TE L-pDI21/9TE MUTANTS & CHIMERIC MUTANTS I—pDISS9 pDISS9/ 21TE pDILS21 pDILS21/ ATAAT pDILS21/ 9TE 88 Figure 20. R N A accumulation in coinfections with chimeric and mutated C N V DI RNAs. A . Total nucleic acid extracts from various coinfections. For each coinfection, total nucleic acid was extracted from all coinfected leaves at six days post inoculation and separated on 1.5% denaturing agarose gels containing 5 m M methyl mercuric hydroxide, and transferred to a membrane. The membrane was hybridized to a nick-translated probe specific for regions I and II of DI R N A sequence. Prior to running the samples on denaturing gels, the samples were run on nondenaturing gels, stained with ethidium bromide and visually inspected to ensure that the levels of ribosomal RNAs were equivalent between samples. Lanes 1 and 2 contain extracts from a mock and WT infection, respectively. Remaining lanes from left to right contain extracts from coinfections with DI RNAs 21, 9, 9/21PE, 21/9PE, 9/21PT, 21/9PT,9/21TE,21/9TE,SS9, SS9/21TE, LS21, LS21/ATAAT, LS21/9TE. Positions of DI R N A 9 monomers and dimers are indicated at the right. Asterisk indicates an intermediate R N A whose primary structure is shown in B. B. The primary structure of an intermediate R N A which accumulates to high levels in DI R N A SS9 coinfections (indicated with an asterisk in lane 11 of A). 89 A E—1 < ^ Cu O H Cu E—1 E—1 r-i „ ZT; ^ c ^ r ^ i c T N O N C N i a N r ^ O N r N i o o c / ^ . — 1 __]_] genomic R N A DI R N A dimer DI R N A monomer 1 2 3 4 5 6 7 8 9 10 11 1 2 1 3 1 4 1 5 B I n Ilia Illb n Ilia Illb 90 EcoRI site used was from the plasmid multiple cloning site and is found 3' to the DI R N A sequence. Transcripts from pDI9/21PE accumulated primarily as a monomer (Figure 20A, lane 5), similar to the situation with DI R N A 21 (in Figure 20, compare lanes 3 and 5). Transcripts from the reciprocal construct (pDI21/9PE) accumulated primarily as a dimer (lane 6), similar to the situation with DI R N A 9 (in Figure 20, compare lanes 4 and 6). Thus, the presence of the sequences found in the PflMl/EcoRI fragment of pDI9 correlate with the characteristic dimer accumulation pattern of DI R N A 9. In addition, the presence of this sequence appears to correlate with an increased overall level of DI R N A accumulation. Whether the overall level of DI R N A accumulation and the level of dimer accumulation are somehow related remains to be determined. To facilitate the comparison of dimer R N A levels in coinfections with differing overall levels of DI R N A accumulation, dimer R N A levels wil l be considered relative to the levels of the corresponding monomer R N A (i.e. the ratio of dimer R N A to monomer RNA) in the following analysis. In an attempt to further narrow down primary sequence differences which may influence dimer accumulation, a second set of chimerics was constructed by exchanging PflMl/Tfil fragments and Tfil/EcoRl fragments (Figure 19). These chimeric DI R N A cDNA clones separate the two major primary sequence differences observed between DI R N A 9 and DI R N A 21; the additional nucleotides found at the beginning of region Ilia in DI R N A 9 and the additional nucleotides found at the beginning of region Illb in DI R N A 21 (see Figures 18 and 19). Coinfections with transcripts from DI R N A constructs carrying the PflMl/Tfil fragment of DI R N A 9 (pDI21/9PT; Figure 20, lane 8 and pDI9/21TE; Figure 20, lane 9) both showed an increase in the ratio of dimer R N A to monomer R N A in comparison to coinfections with transcripts from pDI21 (in Figure 20, compare lanes 8 and 9 with lane 3). The overall level of DI R N A accumulation was also found to be increased in both cases. Coinfections with transcripts from DI R N A constructs carrying the Tfil/EcoRI fragment of DI R N A 9 (pDI21/9TE; lane 10 and pDI9/21PT; lane 7) both 9 1 showed a marginal increase in the ratio of dimer R N A to monomer R N A i n comparison to coinfections with transcripts from pDI21 (in Figure 20, compare lanes 10 and 7 with lane 3). Thus, the presence of sequences found in both the PflMl/Tfil and the Tfil/EcoRI fragments of pDI9 appear to correlate with increased dimer accumulation. Sequences within the PflMl/Tfil fragment of pDI9 also may have an additional effect of increasing the overall levels of DI R N A accumulation. The influence of the major primary sequence differences on dimer accumulation was further investigated by introducing mutations into either pDI9 or pDI21 (Figure 19) and examining the DI R N A accumulation patterns of these mutated DI RNAs in coinfections. In the first of these mutants, all but one of 28 additional nucleotides at the beginning of region Ilia (which are unique to DI R N A 9) were removed from pDI9 to yield pDISS9. This mutant was designated "SS" for "short stem" as the region encompassing the 28 additional nucleotides at the beginning of region Ilia is predicted to form a long stem loop structure in DI R N A 9 (predicted stem loop structures are shown in Figure 23 of the Discussion section of this chapter) which is predicted to be shortened in DI R N A 21 and in this mutant. There are two additional changed nucleotides and one additional deleted nucleotide as a result of the strategy used to introduce this deletion into pDI9 (Figure 18). These three nucleotide changes reside in the loop portion of the shortened stem loop (see Figure 23). Coinfections with transcripts from pDISS9 showed an increase in the ratio of dimer R N A to monomer R N A in comparison to coinfections with transcripts from pDI21 (in Figure 20, compare lanes 11 and lane 3) as did coinfections with transcripts from pDISS9 containing the Tfil/EcoRI fragment of DI R N A 21 (pDISS9/21TE; in Figure 20, compare lanes 12 and lane 3). These observations suggest that sequences in the PflMl/Tfil fragment of DI R N A 9 which were not eliminated in pDISS9 and/or the few differences which distinguish DI R N A SS9 from DI R N A 21 (two substitutions and one deletion) may additionally influence dimer accumulation. 92 DI R N A SS9 and DI R N A SS9/21TE coinfections also accumulated a large amount of an R N A which was intermediate in size between monomer and dimer DI R N A (approximately 700 nucleotides; see lanes 11 and 12 of Figure 20). Similar-sized bands can also be seen at lower levels in coinfections with DI RNAs 21/9PT, 9/21TE, and LS21/ATAAT (described below) in lanes 8, 9, and 14, respectively of Figure 20. This intermediate R N A was cloned from total nucleic acid extracts of DI R N A SS9 and sequenced so that its primary structure could be determined. Assistance with the cloning of this intermediate R N A and determination of its sequence was provided by H . G. Damude. This intermediate was determined to contain a complete duplication of regions II and ITJa/nib, making it 710 nucleotides in size (Figure 20B). Possible explanations for the origin of this intermediate R N A will be discussed. The 28 nucleotides at the beginning of region Ilia were also introduced into pDI21 to yield pDILS21 (Figure 19), with "LS" designating "long stem" to assess the role of this sequence in dimer accumulation. There are two changed nucleotides relative to DI R N A 9 in this region as a result of the strategy used to introduce these additional nucleotides into DI R N A 9 (Figure 18). The overall structure of the predicted stem loop structure in DI RNA 9 is conserved in DI R N A LS21 (see Figure 23). Coinfections with transcripts from pDILS21 showed a very slight increase in the ratio of dimer R N A to monomer R N A as compared to coinfections with transcripts from pDI21 (in Figure 20, compare lanes 13 and 3). This increase was not as dramatic as that observed in coinfections with transcripts from either pDI21/9PT or pDI9/21TE which both contain the entire PflMl/Tfil fragment of DI RNA 9 (in Figure 20, compare lane 13 with lane 8 or lane 9). This observation suggests that sequences in the PflMl/Tfil fragment of DI R N A 9 which were not introduced into pDILS21 and/or the specific differences introduced into this region (two substitutions) also contribute to dimer accumulation. 93 In an effort to convert DI R N A LS21 to DI R N A 9 in terms of dimer accumulation, the additional four nucleotides at the beginning of region nib were specifically removed from pDILS21 to yield pDILS21/ATAAT (Figure 19). Coinfections with transcripts from pDILS21/ATAAT showed an increase in the ratio of dimer R N A to monomer R N A as compared to coinfections with transcripts from pDILS21 (in Figure 20, compare lanes 13 and 14). Thus, it would appear that the lack of the TAAT sequence in the Tfil/EcoRI fragment of DI R N A 9 is a major factor i n the ability of this fragment to influence dimer accumulation. This is further reinforced by the similarity in the ratio of dimer R N A to monomer R N A seen i n pDILS21 / ATA AT coinfections and in coinfections with transcripts of a derivative of pDILS21 which contains the Tfil/EcoRI fragment of DI R N A 9 (pDILS21/9TE; i n Figure 20, compare lanes 14 and 15). The effect of introducing just the T A A T deletion into pDI21 was not addressed in any of the constructs shown in Figure 19, however, based on the above results, coinfections with such a mutant would be predicted to appear similar to those of pDI21/9TE. DI R N A LS21/ATAAT coinfections also accumulated an R N A which migrates approximately halfway between the monomer and the dimer and slightly lower than the intermediate band of pDISS9. Its approximate size was 650 nucleotides and it is likely a DI R N A containing a smaller duplication (for example, a duplication of region Ilia /Illb would result in a DI R N A with an approximate size of 650 nucleotides). There is another higher molecular weight R N A which accumulated in DI R N A LS21/ATAAT coinfections and also in SS9/21TE coinfections. The nature of this R N A was not investigated. At this point in the studies on C N V DI R N A dimers, nothing is known regarding why dimers preferentially accumulate in certain DI R N A coinfections. It is possible that dimers preferentially accumulate because certain DI R N A monomers have a greater tendency to generate dimers or because of defects in the ability of monomers to be generated from certain dimers. A third possibility is that dimers 9 4 accumulate because dimers replicate better than the corresponding monomer. It is important to bear in mind that this third possibility would invalidate using the ratio of dimer R N A to monomer R N A in the preceding analysis. 95 Discussion This work establishes that dimeric forms of tombusvirus DI RNAs can be generated during coinfections. These dimers consist of both precise head-to-tail repeats of DI R N A sequence and imprecise repeats with non-templated bases or small deletions occurring on the right hand side of junctions between monomer units. Individual C N V DI RNAs varied in the amount of dimer which accumulated in coinfections. Analysis of dimer accumulation in chimeric and mutated C N V DI RNAs identified two distinct tracts of sequence which correlate with increased dimer accumulation; one corresponding to the PflMl/Tfil fragment of pDI9 and the other corresponding to the Tfil/EcoRI fragment of pDI9. In the case of the Tfil/EcoRI fragment, one crucial area was further narrowed down to four nucleotides at the junctions between regions Ilia and Illb. The amount of DI R N A dimers in coinfections appeared to depend on the DI R N A used in the coinfection. The template for the production of the DI R N A 9 cDNA clone (which accumulates a high level of dimer in coinfections) was obtained from a plant in which DI R N A had been generated de novo (see Figure 4, lane 2). Agarose gel electrophoresis and northern blot analysis of nucleic acid extracted from this plant revealed that the primary DI R N A component was -900 nucleotides i n length (see Figure 13, lane 8). RT-PCR with this large DI R N A invariably resulted i n the amplification of an -450 nucleotide product. In addition, R N A sequence analysis of this -900 nucleotide R N A yielded a cDNA sequence pattern similar to that depicted for the dimer template in Figure 14A (data not shown). Thus, it seems likely that the 900 nucleotide R N A corresponds to a DI R N A dimer which was generated de novo and whose monomeric form is represented by the DI R N A 9 cDNA clone. Dimers have subsequently been identified in CyRSV DI R N A coinfections (Dalmay et al, 1995; Havelda et al, 1995). Also, RNAs referred to as "new DI R N A s " 96 with sizes consistent with those of DI R N A dimers have been observed by others i n both C N V and TBSV DI R N A coinfections (Chang et al, 1995). Characterization of CyRSV DI R N A dimers revealed several similarities with the DI R N A dimers described in these studies: they are head-to-tail repeats of unit length RNA; the majority of junctions (more than 75%) consisted of precise fusions of 5' and 3' termini; deletions were found on the right hand side of the junctions; and non-templated nucleotides were also found within the junctions, although not as frequently (two of 30 cloned junctions versus six of 21 clones in this study; Dalmay et al, 1995). DI R N A dimers of similar overall structure have also been described i n association with the related carmovirus TCV (Cascone et al, 1990; Carpenter et al, 1991). TCV DI R N A dimers also contains non-templated nucleotides within junctions, however, no deletions were found within the five junctions analysed. Dimeric forms of the R N A 2 segment of Flockhouse nodavirus have also been tentatively identified (Ball, 1994). Two different mechanisms currently exist to explain the generation of R N A multimers from R N A monomers: (1) rolling circle replication (Figure 21) and (2) R N A recombination (Figure 22). Replication-independent intermolecular ligation of monomer units could be hypothesized as a third possible mechanism. N o evidence for replication-independent intermolecular ligation has been found to date from laboratories studying R N A recombination (Simon and Bujarski, 1994). In addition, heterodimers were not observed when two different CyRSV DI R N A s were used in coinfections (Dalmay et al, 1995). By analogy with these studies, replication-independent intermolecular ligation is not likely to be the mechanism by which C N V DI R N A dimers are generated. Hallmarks of rolling circle replication, implicated as the mechanism of replication of some satellite RNAs, virusoids, and viroids (Branch and Robertson, 1984; Bruening et al, 1988; Bruening et al, 1991; Symons, 1992; Diener, 1993), are the existence of circular R N A intermediates and the ability of at least (+) multimers to 9 7 Figure 21. Rolling circle R N A replication. Starting from the top left, linear (+) R N A monomers ligate to form circular RNAs which are then tandemly copied to yield (-) head-to-tail multimers. These multimers can be copied to yield (+) multimers or, alternatively, (-) monomers can be released from (-) multimers by autocatalytic cleavage or by host-derived factors, circularize and then copied to yield (+) multimers. To complete the cycle, (+) monomers are released from (+) multimers by autocatalytic cleavage or by host-derived factors. Adapted with permission from Bruening et al, (1988). For details on this mechanism, see Branch and Robertson, (1984), Bruening et al, (1988), Bruening et al, (1991), Symons, (1992), and Diener, (1993). 98 Figure 22. Generation of turnip crinkle virus satellite R N A and DI R N A multimers by R N A recombination. Reprinted with permission from Carpenter et al. (1991a). After synthesizing of a unit length of (+) R N A (a complete unit in case 5b or a premature termination product in case 5a), the replicase complex initiates another round of synthesis prior to releasing the nascent (+) RNA. For details on this mechanism see Cascone et al, (1990) and Carpenter et al. (1991a). 99 be autocatalytically processed into monomers. A ladder-like distribution of R N A multimers with three, four, five and more monomer units can often be visualized by electrophoresis and Northern blotting (for example, see Passmore and Bruening, 1993). Circular forms of C N V DI RNAs were not detectable when DI R N A 42 monomers purified from non-denaturing agarose gels (where circular and linear forms of R N A are expected to comigrate; see Schumacher et al, 1983 and Linthorst and Kaper, 1984) were electrophoresed on denaturing acrylamide gels (which resolve circular and linear forms; see Schumacher et al, 1983 and Linthorst and Kaper, 1984). In addition, the cDNA sequence yielded from DI R N A 42 monomer template purified from non-denaturing agarose gels terminated in a strong stop (see Figure 14A and 14B), indicating that this R N A was linear. Autocatalytic cleavage of DI R N A 42 dimer transcripts of either polarity into monomers was also not detectable under in vitro transcription conditions (see Materials and Methods chapter for a description of in vitro transcription conditions; data not shown). It is important to recall that the dimer transcripts used in these analyses lacked the A or the U in the center of the junctions of (+) or (-) transcripts, respectively (see p. 80 of preceding Results section) and this may have a deleterious effect on autocatalytic cleavage if it does occur. A ladder of R N A multimers was also not seen in total nucleic acid extracts of coinfections (see Figures 11, 13, 18, and 21). As these distinguishing features of rolling circle replication do not seem to be associated with C N V DI RNAs, it does not seem likely that this is the mechanism by which dimers are generated. In addition, the formation of DI RNAs containing internal duplications (the intermediate R N A seen in DI R N A SS9 coinfections) is not adequately explained by rolling circle replication. The capacity of tombusviruses to undergo R N A recombination, as evidenced by their ability to generate DI RNAs and by direct experimental evidence (see White and Morris, 1994b), coupled with the imprecise nature of the junctions between monomer units is more reflective of a recombination-mediated mechanism of 100 dimer generation. This mechanism was first proposed by Carpenter et al. (1991a) to explain the generation of dimeric TCV satellite RNAs and DI RNAs. This mechanism was proposed as an alternative to rolling circle replication, as circular intermediates and autocatalytic processing of multimers were not identified in this system (Simon et al, 1988; Cascone et al, 1990). In this mechanism, dimers are generated by the replication complex initiating another round of replication prior to release of the completed nascent strand (Figure 22; Cascone et al, 1990; Carpenter et al. 1991a). This recombination event is believed to take place primarily during the synthesis of (+) R N A from (-) RNA. To explain the presence of deletions found only at the left hand side of the junctions of satellite R N A dimers, the model presented in Figure 22 predicts that replication can terminate prematurely prior to reinitiation (step 5a). A similar recombination-mediated mechanism has also been proposed for the formation of CyRSV DI R N A dimers (Dalmay et al, 1995). Reinitiation in this system is believed to take place on the same template as only homodimers were observed when two different DI RNAs were used for coinfection. A recombination-mediated model for the generation of C N V DI R N A dimers w i l l be presented and discussed further in the next chapter of this thesis. Other studies have also demonstrated that primary sequence differences can influence the accumulation of dimers. A n area analogous to the junction of regions Ilia and Illb has been implicated in the case of CyRSV DI RNAs (Havelda et al, 1995). Deletion of region III of C N V and TBSV DI RNAs (analogous to region Ilia in this study) also resulted in the appearance of new DI RNAs in coinfections which are presumably dimers (Chang et al, 1995). In the case of TCV satellite RNAs, a short tract of sequence near the 5' end (nucleotides 79 to 100) was identified as a region which influenced dimer accumulation (Simon et al, 1988; Carpenter et al, 1991). Size of the DI R N A template alone does not appear to influence DI R N A accumulation pattern. Consider, for example, that DI R N A 21 and DI R N A SS9 differ in size by only two nucleotides yet DI R N A SS9 accumulates much more 101 dimer than DI R N A 21 and also accumulates an intermediate R N A which is not seen in DI R N A 21 coinfections. In the studies of Dalmay et al, (1995), when deletions were made within a CyRSV DI R N A with a complete block C (analogous to region III), an increase in dimer accumulation was observed. Insertion of sequence from an unrelated virus into a CyRSV DI R N A harboring an internal deletion within block C had the opposite effect with a decrease in dimer accumulation observed as the size of the insert increased. These results led to the conclusion that size of the monomer molecule is a major factor in the accumulation of dimers. The results of this study are not consistent with this conclusion. A n unforeseen effect of changes in DI R N A sequence was the accumulation of intermediate sized DI RNAs containing an internal duplication of regions II and Illa/IIIb. It is plausible that these DI RNAs are also products of the same recombination mechanism proposed for the generation of DI R N A dimer. This possibility wil l be discussed further (next chapter). The identification of two distinct regions which contribute to the high level of dimer accumulation seen in coinfections with DI R N A 9 prompted the reexamination of the R N A accumulation patterns in coinfections with other DI RNAs characterized in this study in context with the primary sequence in these regions. The bottom panel of Figure 11, shows that coinfections with DI RNAs 12 and 16 (lanes 3 and 4, respectively) contain prominent levels of higher molecular weight RNAs. Two higher molecular weight bands can be seen in DI R N A 12 coinfections; based on size estimates, the top band likely corresponds to a complete head-to-tail repeat of DI R N A 12 sequence while the other band may correspond to a DI R N A with an internal partial duplication or a dimer with a deletion. The ratio of dimer to monomer in DI R N A 12 coinfections is slightly reduced in comparison to DI R N A 9 coinfections. The junction between regions II and Ilia in DI R N A 12 is different from DI R N A 9 and there are two additional changed nucleotides relative to DI R N A 9 within the sequence corresponding to the PflMl/Tfil fragment (see 102 Figure 8). This may account in part for the reduced level of dimer accumulation i n comparison to DI RNA 9. The junction between regions Ilia and Illb in DI R N A 12 is different from both DI R N A 21 and DI R N A 9, making it difficult to ascertain the influence (if any) of this region on dinier accumulation. It is possible that other sequences, which are specific to DI R N A 12, influence dimer accumulation. In DI R N A 16 coinfections, the higher molecular weight R N A migrates slightly faster than the lower of the two higher molecular weight bands in DI R N A 12 coinfections. Thus, the prominent form of DI R N A 16 in coinfections likely contains a partial duplication, a situation somewhat analogous to that of DI R N A SS9 coinfections. However, this intermediate R N A is much more prominent relative to the levels of DI R N A monomer and dimer in DI R N A 16 coinfections than in DI R N A SS9 coinfections. It is again difficult to ascertain which regions of a DI R N A could be responsible for the accumulation of these intermediate R N A based on primary sequence comparison of DI R N A SS9 versus DI R N A 16. However, it is noted that, like DI R N A SS9, DI R N A 16 maintains the last of the 28 additional nucleotides found in DI R N A 9 at the beginning of region Ilia. The observation that subtle changes in the primary sequence of a DI R N A can considerably alter the R N A accumulation pattern coupled with the identification of two different areas of a DI R N A which may somehow interact to promote dimer accumulation, suggest that secondary (or higher) structure differences between individual C N V DI RNAs could be responsible for differences in dimer accumulation. This would make predictions about R N A accumulation patterns based solely on comparison of primary sequence very difficult. In this regard, it is of interest to note that the junction between region II and region Ilia and region Ilia and Illb adopt different predicted secondary structures in DI R N A 21 and DI R N A 9. In both regions, the predicted stem loop structure is longer in DI R N A 9 than in DI R N A 21. A n example of the predicted stem loop structures surrounding the junction of regions II and Ilia are shown in Figure 24. Secondary structure 103 differences could manifest themselves in many different ways. The interaction of proteins with the R N A template could be affected; for example, as suggested i n Carpenter et al, (1991), the replicase/nascent strand dissociation kinetics could be altered resulting in the preferential accumulation of dimers. Alterations in the secondary structure could also serve to bring the 5' and 3' ends into greater proximity, thereby increasing the chances of reinitiation prior to release on the nascent strand as suggested in Carpenter et al, (1991) and Dalmay et al, (1995). These suggestions are all based on the assumption that dimers accumulate in certain coinfections because of a greater ability of certain DI R N A monomers to generate dimers. While this is an attractive suggestion which fits well with the data obtained in this study and in the studies of others, the possibility that dimers accumulate because certain dimers are better templates for replication or because of defects in the ability of monomers to be liberated from certain dimers cannot yet be discounted. Finally, the biological activity of Ssp transcripts coupled with the observation that coinfections with synthetic DI R N A 42 dimer transcripts yield DI R N A 42 monomers suggests that internally-located replication initiation sites can be recognized by the C N V replication complex. This may be envisioned as one possible mechanism by which DI R N A monomers can be generated from a DI R N A dimer. The generation of monomers from input dimer transcripts in coinfections was also observed in the case of CyRSV DI RNAs (Dalmay et al, 1995). The results of these studies and the studies of Dalmay et al, (1995) were both inconclusive with respect to whether dimers represent obligate intermediates in DI R N A replication. There is an inherent imprecision in the process which generates dimers as evidenced by the heterogeneity in junctions between monomer units. Dimers can be generated during coinfections probably, as proposed here, as byproducts of R N A recombination; if this happens, they are able to serve as templates for the generation of monomers. The possibility that the ability of DI RNAs to form dimers may impart certain advantages to a DI R N A in terms of replication should not, however, 104 be overlooked. A n interesting potential consequence of the DI R N A dimer structure in terms of replication is discussed further in the next chapter. 105 A A U A A A U A A A A A A A C-G A A C-G U-A C A U-A C A W C C - G A G-C ^G-C T c - G c A A C " G C A C U - G C U - G A - U A - U u u c A A C A G-C A © C A G-C U-A A A C G C G G - C A > * U-A U-A U-A U-A C-G C-G C-G C-G U-A U-A U-A U-A G-C G-C G-C G-C A - U A - U A - U A - U G-C G-C G-C G-C A - U 5'" ^ 3' A - U 5'^  ^3' A - U A - U 5'" v3' D l RNA 21 DI RNA SS9 D I R N A 9 DI R N A LS21 Figure 23. Predicted stem loop structures of various DI RNAs at junctions between regions II and Ilia. The depicted structures for DI RNAs 21 and DI R N A 9 were determined using the algorithm of Zuker (Zuker and Stiegler, 1981; Zuker, 1989) on the entire DI R N A sequence; only the junction sequence around regions II and Ilia are shown. Circled bases represent nucleotide changes introduced during the construction of pDISS9 and pDILS21; arrows delineate the border of the 28 additional nucleotides found in DI R N A 9. 106 GENERAL DISCUSSION In this final chapter, the insights that tombusvirus DI R N A research has provided into tombusvirus replication thus far wil l be reviewed. Additional concepts regarding the generation and possible biological significance of dimer RNAs wil l also be presented. These concepts have arisen from observation of primary sequence and predicted secondary structures and have not been tested by the experiments in this thesis. They are presented to provide some direction for future work. Tombusvirus DI RNAs have been utilized to delineate the minimal cz's-acting sequences required for R N A replication. A l l tombusvirus DI RNAs consist of a mosaic of the same three main non-contiguous portions of the viral genome: the 5' untranslated region; a region derived from the p92 ORF; and a 3'-terminal region (designated A, B, and C, respectively in the case of CyRSV DI RNAs; I, II, and III/IV, respectively in the case of TBSV DI RNAs; and I, II, and either III/IV or ma/nib, respectively in the case of C N V Dl RNAs). The similarity in cz's-acting sequences retained by different tombusviruses may reflect a common replication strategy. This is further highlighted by the ability of C N V to support the replication of TBSV DI RNAs (White and Morris, 1994a; White and Morris, 1994c). Whether it is the primary sequence of these cz's-acting elements or a certain secondary (or higher) structure adopted by these cz's-acting elements which are recognized for replication remains to be determined. Very recently, the putative structure adopted by the last 77 nucleotides of CyRSV DI RNAs was subjected to mutational analysis (Zoltan and Burgyan, 1995). This region was predicted to form three stem loop structures interspaced with two non-base-paired regions. Mutations which deleted any of the three stem loops or disrupted their structure were found to be detrimental to DI R N A replication. Compensatory mutations which restored these structures also 107 restored the viability of the DI RNA. These results suggest that the proposed structure rather than the primary sequence is important for CyRSV replication. Similar structures can be adopted by related tombusviruses suggesting the general relevance of this structure in tombusvirus replication (Zoltan and Burgyan, 1995). In addition to the similarities in cz's-acting sequences required for DI R N A replication, tombusvirus DI RNAs can generate head-to-tail repeats of DI R N A sequence (this thesis; Dalmay et al, 1995). In the case of both CyRSV and CNV, these dimers can be used as templates for the generation of monomers, however, it has not been established if dimers represent obligate replicative intermediates. The studies in this thesis are consistent with the idea that C N V DI R N A dimers are formed by R N A recombination; this has also been proposed in the,'case of CyRSV DI R N A dimers (Dalmay et al, 1995). In the case of both C N V and CyRSV DI R N A dimers, small deletions of varying size are found on the right hand side of the junction. Two different models of a general R N A recombination mechanism can be invoked to account for this observation (Figure 24). In model 1, recombination would occur during synthesis of (+) R N A from (-) templates. The replicase complex makes a complete copy of a monomer and then, along with the nascent (+) R N A would reinitiate another round of replication prior to releasing the nascent strand. This reinitiation event can be precise, leading to a complete duplication of DI R N A sequences (case a) or imprecise, leading to deletions on the right hand side of the junction (case b) or an incomplete duplication of DI R N A sequence (the intermediate sized R N A seen i n DI R N A SS9 coinfections; case c). In model 1 the reinitiation event which would generate precise dimers (case a) would occur at a known promoter of R N A synthesis. In model 2, recombination would occur during synthesis of (-) R N A from (+) templates. Synthesis of (-) R N A can terminate prior to reaching the 5' end of the template (premature termination; cases b and c); this must be followed by a precise 1 0 8 Figure 24. Models for the generation of C N V DI R N A dimers. In both models, the shaded ball represents the viral replication complex, (+) R N A is represented by a solid line, (-) R N A is represented by a broken line, and junctions between monomer units are indicated with a vertical line. Three different options (a, b, and c) are indicated in each model to explain the different R N A products observed in this study. In model 1 a, b, and c correspond to different sites of reinitation whereas i n model 2 a corresponds to run-off at the end of the R N A template and b and c correspond to different sites of premature termination The final R N A product shown in each model is a complete, (+) dimer which would arise from option a in both cases. See text for details. 109 110 reinitiation event in order to conserve sequences at the left hand side of the junction. It is important to note that the products of premature termination would lack exact 3' termini and therefore may be unable to undergo further replication. These premature termination products can restore their 3' termini by undergoing another complete round of replication. It has been observed previously that the 5' and 3' termini of the C N V genome are roughly complementary (Rochon and Tremaine, 1989). In this respect, it is of interest to note that computer predictions of DI R N A structure using the computer algorithm of Zuker (Zuker and Stiegler, 1981; Zuker, 1989) show that the 5' and 3' termini of all C N V DI RNAs in this study can be brought into proximity by secondary structure. Two different predicted interactions between the 5' and 3' termini of (+) DI R N A are shown in Figure 25 A and 25B. The proximity of the 5' and 3' termini may facilitate the reinitiation event in either model. Thus, the replicase complex initiates another round of replication on the same template (i.e. a ds-recombination event). Congruent with this idea is the observation that when coinfections were performed with two different CyRSV DI RNAs, progeny DI R N A dimers were found to be exclusively homogeneous (Dalmay et al, 1995). Examination of the sequences surrounding the consensus junction sequence in C N V DI R N A dimers shows that an 11 nucleotide sequence at the C N V 3' terminus is imperfectly repeated three times at the C N V 5' terminus (Figure 26A). Similar repeats can be found in the DI RNAs or genomes of all definitive tombusviruses for which the complete sequence of the genome is known (Figure 26B). Sequences at the 3' termini of viral genomic RNAs have been shown to be required for replicase recognition (David et al, 1992; Duggal et al, 1994; Pogue et al, 1994). The presence of a reiterated putative C N V genomic R N A replicase recognition site on the right hand side of the DI R N A dimer junction site raises the 111 Figure 25. Some possible interactions of the 3' terminus of (+) DI R N A with other regions of DI RNA. Vertical or horizontal lines represent Watson-Crick base-pairing, vertical arrow indicates the deletion endpoint seen in DI R N A 9 dimers (see Figure 15 of Chapter 2) in B and junction between regions I and II in C, dashed arrow indicates the direction of R N A synthesis, dashed line indicates continuation of sequence, bases in italics are part of the repeated sequences (see Figure 26). A and B are computer-assisted predictions of the interaction between the 5' and 3' termini of (+) C N V DI RNA; in B the stem loop structure is formed between repeats I and II defined in Figure 26. C is a possible heteroduplex between the 3' termini of (+) DI R N A and a region immediately 3' of region II on (-) DI RNA. AGA A A UUC UC CA GGA UUUC U C A A W5" q ^ c a A C a b k k k b k c h h h B c c U A C G U-G U-A A-U A-U A-U G-C [+]5^A-U C A A C C U U G G U U G U G U U A U C U G i i i ,C c c iskc Guk G A 3' U n 3 ' U D G A A G G U C A U U U G C U G C U G U A £ y ki6AiiGcA6 kkku6 C A G C C c 113 Figure 26. Organization and comparison of repeated sequences found at the junction of C N V DI R N A dimers. A. Dimer R N A is shown schematically using adjoining arrows. The bars below the arrows show the approximate location of the repeated sequences. The repeats have been assigned Roman numerals based on their 5' to 3' order of appearance in C N V genomic RNA. The repeated sequences as they appear in a (+) dimer are compared below; rows with a single asterisk indicate that at least three of four nucleotide positions are identical, whereas rows with two asterisks indicate that the nucleotide is conserved in all four repeats. The numbering corresponds to C N V genomic R N A nucleotide positions as defined in Rochon and Tremaine, (1989). Note that repeat I spans the dimer R N A junction to include one nucleotide (CNV nucleotide 4701) from the 3' end of the monomer. Below this comparison is a comparison of repeat IV with the inverse complement of repeats I, II and III. B. Comparison of repeated sequences in different tombusviruses. CyRSV repeat sequences were derived from the published sequence of CyRSV DI R N A dimers (Dalmay et al, 1995) and de nouo-generated CyRSV DI RNAs (Burgyan et al, 1991); TBSV repeat sequences were derived from the published sequence of de nouo-generated TBSV DI RNAs (Knorr et al, 1991); and A M C V repeat sequences were derived from the published sequence of A M C V genomic R N A (Tavazza et al., 1994). Note that unlike in A , rows with one asterisk indicate that the nucleotide is conserved in all four tombusviruses. 114 i n * * * * * * * * * * * * * * * IV CAGAAAUGCAG 4689-4699 I CAGAAAUUCUC 4701-10 II CAGGAUUUCUC 11-21 I I I GAGAAUUUCUC 67-77 * * * * * * * * * * * * * *** IV CAGAAAUGCAG • , . c inverse complement of: I GAGAAUUUCUG 47 01-10 II GAGAAAUCCUG 11-21 I I I GAGAAAUUCUC 61-11 B I I i n IV CNV CyRSV TBSV AMCV -k k k k k k k ~k ic CAGAAAUUCUC CAGAAAUCCUC CNGAAAUUCUC CNGAAAUUCUC CAGGAUUUCUC CAGGACACCUC CAGGAUUUCUC CAGGAUUUCUC * * * * ***** GAGAAUUUCUC GAGGAUUUCUC GAGAAUUUCUC GGGAAUUUCUC *** * ****** CAGAAAUGCAG CAGCAAUGCAG CAGAAAUGCAG CAGCAAUGCAG 115 possibility that these sequences may sequester the viral replicase complex for synthesis of (-) DI R N A monomers from (+) DI R N A dimers. This sequestering of viral replicase may be a means by which DI RNAs can interfere with helper virus accumulation. These repeats may also function as signals which direct the reinitiation events implicated in recombination model 1. In fact, the ends of the large deletions found near the dimer junctions in both C N V (see Figure 15) and CyRSV coincide with these repeats: the deletion end is found immediately downstream of repeat II in the case of C N V DI R N A dimers and immediately downstream of repeat III in the case of CyRSV DI R N A dimers (Dalmay et ah, 1995). The inverse complement of repeats I, II, and III also align well with sequences found at the 3' termini of (+) RNA, further suggesting that they could act as replicase reinitiation sites (Figure 26A). It should also be noted that these repeats have the potential to adopt secondary structures which could cause the dissociation events implicated i n recombination model 2 (see example depicted in Figure 25B). Hairpin structures introduced into the 5' end of TBSV R N A donor templates have been observed to shift the recombination site to the 3' side of the hairpin (White and Morris, i n press). Based on these results, it was proposed that the hairpin may be acting as an obstacle, causing premature termination of R N A synthesis. These studies on a related tombusvirus provide a nice parallel to the possible situation depicted i n Figure 25B, where a hairpin structure formed between repeats I and II (also predicted to be a possible 5' end structure of (+) C N V DI RNAs using the computer algorithm of Zuker) causes premature termination resulting in the 23 nucleotide deletion seen on the right hand side of C N V DI R N A dimer junctions. Alternatively, the secondary structures adopted by these repeats (as opposed to the primary sequence) may direct reinitiation in a manner analogous to the R N A recombination mechanism proposed for TCV (Simon and Bujarski, 1994). 116 The prominent intermediate R N A seen in DI R N A SS9 coinfections is proposed to result from the same recombination-mediated mechanism as proposed for complete dimers (case c in both models 1 and 2). Recombination, in this case, is somehow redirected to the beginning of region II. Examination of sequences surrounding the junction of regions I and II failed to reveal a good match with the repeated sequences shown in Figure 26, indicating that the sequences similar to repeats I through IV do not govern either the reinitiation or premature termination events in the generation of this RNA. There also does not appear to be a good potential for the 5' end of (-) R N A to anneal back on itself immediately around the junction between regions I and II to facilitate a reinitiation event at region II. It was noticed that there is a potential for a heteroduplex between the 3' terminus of (+) R N A and a region immediately 3' of region II on (-) R N A (Figure 25C). Perhaps this could serve to bring these sequences into proximity, allowing synthesis to recommence at the beginning of region II. As stated in the previous chapter, bands analogous to this intermediate sized R N A can be seen at high levels in DI R N A SS9/21TE and DI R N A 16 coinfections and at lower levels in coinfections with other DI RNAs. Therefore, like dimers, the level of accumulation of this intermediate R N A seems to depend on the DI R N A used in the coinfection. It is generally assumed that the mechanism of DI R N A replication mirrors that of viral R N A as DI RNAs utilize the viral replication complex and derive their cz's-acting sequences from viral RNA. Are the above observations regarding the replication of tombusvirus DI RNAs valid for the replication of the tombusvirus genomic RNA? The identification of cz's-acting elements at the extreme 5' and 3' termini seems reasonable in light of what is currently known about replication i n well-characterized plant R N A viruses (David et al, 1992; Duggal et al, 1994; Pogue et al, 1994). The possible requirement of an additional internal sequence element is an intriguing aspect of tombusvirus replication which certainly requires further investigation. The cz's-acting elements identified in DI RNAs wi l l have to be studied 117 in the context of the entire viral genome in order to confirm or refute their involvement in viral replication. The finding that DI R N A dimers may serve as intermediates in the generation of monomers could point to the involvement of dimers in the replication of genomic RNA. Currently, there is no evidence that head-to-tail repeats of the entire C N V genome exist in infected tissue, however this may be due to a lack of sensitivity in our RT-PCR protocol which presently requires 1 ng of dimer R N A template for efficient amplification (unpublished observations). The suppression of viral R N A accumulation in DI R N A containing infections may be due instead, at least in part, to a different, more efficient dimer RNA-based DI R N A replication strategy. Continuation of studies on tombusvirus DI RNAs as models for tombusvirus replication would certainly benefit from purification of an active tombusvirus replication complex and the development of an in vitro replication system. This would allow the identification of the host component(s) of the replication complex and permit detailed biochemical analysis of the interaction of replication proteins with R N A template. Determining the exact role of the cz's-acting sequences involved in tombusvirus replication wil l require the analysis of structure(s) which may be adopted by these sequences. Finally, the phenomenon of dimer formation by tombusvirus DI RNAs may prove to be useful for working out the mechanism of R N A recombination in tombusviruses. Several potential contributing factors have been identified in this study including: two distinct regions within the DI R N A sequence, one of which was also independently implicated in CyRSV DI R N A s (Havelda et al, 1995); repeated 3' terminal sequences at the 5' terminus; and possibly the overall structure of the DI RNA. 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