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Molecular and biochemical characterization of viral and vector components required for cucumber necrosis… Kishore Kakani, Naga 2004

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Molecular and Biochemical Characterization of Viral and Vector Components Required for Cucumber Necrosis Virus Transmission By  Naga Kishore Kakani B.Sc. (Biology and Chemistry) Sri Venkateswara University, India, 1990 M.Sc. (Virology), Sri Venkateswara University, India, 1993 M.Tech. (Biotechnology), Jawaharlal Nehru Technological University, India, 1996  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF T H E REQUIREMENTS FOR T H E DEGREE OF DOCTOR OF PHILOSOPHY In T H E F A C U L T Y OF GRADUATE STUDIES (Plant Science)  We accept this thesis as conforming to the required standard  THE UNIVERSITY OF BRITISH COLUMBIA October 2004 ©Naga Kishore Kakani, 2004  Abstract Natural transmission of Cucumber necrosis virus (CNV) occurs via zoospores of the chytrid fungus Olpidium bornovanus. Transmission involves specific adsorption of virus particles onto the zoospore plasmalemma prior to infestation of cucumber roots by virus bound zoospores. In order to determine if specific regions of the CNV capsid are involved in transmission, several naturally occurring CNV transmission mutants were isolated and characterized. Analysis ofthe mutants showed that the CNV trimer cavity at the particle quasi three-fold axis plays an important role in transmission, and, moreover that the reduction in transmission is at least partially due to the reduced ability of mutants to bind to zoospores. In vitro virus/zoospore binding studies have shown that pre-treatment of zoospores with trypsin and sodium periodate each decrease CNV binding by approximately 80%, whereas no reduction in binding was found when zoospores were treated with phospholipase C. These studies suggest an important role for zoospore proteins and/or glycoprotein(s) in virus attachment. In virus overlay assays, CNV virions bound to specific-sized zoospore proteins, but CNV transmission mutants showed little or reduced binding. Several sugars were used to study their inhibitory potential on CNV binding to zoospores in vitro. It was found that a variety of mannosecontaining sugars inhibited CNV binding to zoospores whereas several others did not. These studies suggest that the putative zoospore receptor may be a mannose-containing glycoprotein. Many animal virus particles undergo conformational changes upon binding to their cellular receptors. CNV is an icosahedral virus and like many other isometric plant viruses, undergoes expansion in the presence of EDTA at an alkaline pH. In the case of  ii  CNV, we have demonstrated that during expansion, the internally located coat protein RNA binding domain (R) and arm domains translocate to the particle exterior, becoming protease sensitive. Protease digestion experiments of zoospore-bound virus have revealed that CNV undergoes conformational change upon binding to zoospores and that the conformationally altered virion resembles the swollen conformation. In addition, we have found that a CNV mutant defective in vector transmission is unable to undergo this conformational change. This is the first time that conformational change in a plant virus particle has been shown to be essential for vector transmission.  iii  Table of Contents Abstract  ii  Table of Contents  iv  List of Tables  viii  List of Figures  ix  List of Abbreviations  xi  Dedication  xv  Acknowledgements  xvi  1 CHAPTER ONE: Literature Review 1.1 Introduction 1.2 Plant virus transmission 1.2.1 Arthropod transmission 1.2.1.1 Aphid transmission 1.2.1.1. INoncirculative/Non-persistent 1.2.1.1.1.1 Capsid strategy 1.2.1.1.1.2 Helper strategy 1.2.1.1.2 Noncirculative/Semi-persistent (Helper strategy) 1.2.1.1.3 Circulative/Nonpropagative 1.2.2 Thrips transmission 1.2.3 Nematode transmission 1.2.4 Fungus transmission 1.2.4.1 Zoospore structure and the life cycle of Olpidium 1.2.4.2 Life cycle of plamodiophorids 1.2.4.3 Modes of fungus transmission 1.2.4.3.1 In vitro transmission 1.2 4.3.2 In vivo transmission 1.3 Virus-receptor interactions 1.3.1 Virus receptors 1.3.2 Secondary receptors 1.3.3 Conformational changes involved in virus entry 1.3.3.1 Enveloped viruses 1.3.3.2 Nonenveloped viruses  iv  1 1 4 6 6 8 7 8 10 12 16 17 20 21 22 23 24 27 30 31 32 33 34 36  1.4 Cucumber necrosis virus 1.5.1 Genomic organization 1.5.2 Functions of CNV encoded proteins 1.5.3 DI RNAs 1.5.4 Structural aspects of the CNV particle  39 40 40 42 43  1.5 Role of N-terminal arm in T=3 icosahedral plant viruses 1.6 Virus particle dynamics in T=3 icosahedral plant virus 1.7 Summary and thesis Objectives 1.8 References  44 46 49 64  2 C H A P T E R T W O : Identification of Specific Cucumber Necrosis Virus Coat Protein Amino Acids Affecting Fungus Transmission and Zoospore Attachment  81  2.1 Introduction 81 2.2 Materials and methods 83 2.2.1 Isolation of CNV transmission mutants 83 2.2.3 Virus purification 83 2.2.4 Fungus transmission assay 84 2.2.5 Cloning and sequence analysis of transmission mutants 85 2.2.6 In vitro transcription and inoculation of plants 85 2.2.7 In vitro binding assay 86 2.2.8 Homology modelling 86 2.3 Results 87 2.3.1 Isolation of transmission mutants from mechanically passaged CNV 87 2.3.2 Mutations in CNV transmission mutants map to either the CP shell or protruding domain 89 2.3.3 CNV transmission mutants show decreased binding to zoospores in vitro. 91 2.3.4 An artificial double mutant transmits and binds to zoospores at a lower efficiency than either ofthe individual mutants 92 2.4 Discussion 92 2.5 References 102  3 C H A P T E R T H R E E : Evidence that Binding of Cucumber Necrosis Virus to Vector Zoospores Involves Recognition of Oligosaccharide 105 3.1 Introduction 3.2 Materials and methods 3.2.1 Virus isolates and purification 3.2.2 Maintenance of fungal cultures  v  105 107 107 107  3.2.3 In vitro zoospore binding assay 3.2.4 Trypsin, periodate, and phospholipase C treatment of O. bornovanus zoospores 3.2.5 Virus overlay assays 3.2.6 Microtiter plate binding assays 3.2.7 Labelling of O. bornovanus zoospores with FITC-labelled lectins 3.3 Results 3.3.1 Binding of CNV to zoospores is saturable and specific 3.3.2 Periodate, trypsin, and phospholipase-C treatment of O. bornovanus zoospores 3.3.3 CNV binds to specific-size proteins in O. bornovanus zoospore extracts 3.3.4 CNV transmission mutants bind with reduced efficiency invirus overlay assays 3.3.5 CNV binding to O. bornovanus zoospores is competitively inhibited by several mannose-containing sugars 3.3.6 Surfaces of O. bornovanus zoospores contain fucose and mannose and/or glucose residues 3.4 Discussion 3.5 References  107 108 108 109 111 Ill Ill 112 113 114  114 116 116 128  4 CHAPTER FOUR: Evidence that Vector Transmission of a Plant Virus Requires Conformational Change in Virus Particles 131 4.1 Introduction 4.2 Materials and methods 4.2.1 Virus purification 4.2.2 Maintenance of O. bornovanus cultures 4.2.3 Agarose gel electrophoresis of purified virus 4.2.4 In vitro mutagenesis 4.2.5 In vitro transcription and inoculation of plants 4.2.6 Fungus transmission assays 4.2.7 In vitro binding assays 4.2.8 Production of polyclonal antiserum 4.2.9 In vitro swelling of virions 4.2.10 Limited proteolysis of native, swollen and zoospore-bound virus 4.2.11 Amino acid sequence analysis of trypsin-digested swollen virions 4.3. Results 4.3.1 Proteolytic digestion patterns of zoospore-bound CNV differ from those of native CNV 4.3.2 Effect of mutations of Pro73 and Pro85 on CNV particle formation 4.3.3 Transmission efficiency of Pro73Gly virions by O. bornovanus is reduced dramatically 4.3.4 Binding efficiency of Pro73Gly to zoospores is similar to that of vi  131 133 133 133 134 134 135 135 135 136 137 137 138 138 138 140 142  wild-type CNY 142 4.3.5 The swollen conformation of Pro73Gly is different from that of CNV.. ..143 4.3.6 The N-terminal region of swollen CNV particles is accessible to trypsin digestion 144 4.3.7 Zoospore-bound Pro73Gly does not undergo conformational change 145 4.4 Discussion 145 4.5 References 159  5 CHAPTER FIVE: General Discussion 5.1 References  163 171  6 CHAPTER SIX: APPENDIX  173  6.1 Introduction 6.2 Materials and methods 6.2.1 Virus purification 6.2.2 Maintenance of O.bornovanus cultures 6.2.3 In vitro mutagenesis 6.2.4 In vitro transcription and inoculation of plants 6.2.5 Fungus transmission assay 6.3 Results and discussion 6.4 References  vii  173 174 174 174 174 175 175 175 182  List of Tables Table 1.1 Invertebrate vectors of plant virus genera and their mechanisms of transmission  5  Table 1.2 Viruses transmitted by Olpidium vectors  29  Table 1.3 Viruses transmitted by plasmodiophorid vectors  30  Table 1.4 Viral protein receptors and co-receptors  38  Table 1.5 Viral carbohydrate receptors  39  Table 2.1 Transmission and in vitro binding efficiencies of CNV mutants  88  Table 3.1 Sugars classified as inhibitors and noninhibitors in virus/zoospore binding assays  121  Table 4.1 Oligonucleotides used for constructing Pro73 and Pro85 mutants  136  Table 4.2 Properties of CNV Pro73 and Pro85 mutants  146  Table 6.1 Oligonucleotide primers used for constructing various CNV mutants  176  Table 6.2 Properties of CNV mutants  179  viii  List of Figures Figure 1.1 Schematic representation of the ultrastructure of the O. brassicae zoospore  23  Figure 1.2 Life cycle of O. brassicae  25  Figure 1.3 Genomic organization of CNV  48  Figure 1.4 Linear and three-dimensional structure of CNV subunit (C-type)  52  Figure 1.5 Ribbon diagrams of homology modelled CNV CP subunits and their location on the particle icosahedral axis  53  Figure 1.6 Surface representation of CNV the j8-annulus  54  Figure 1.7 Structure of the CNV particle  55  Figure 2.1 Agarose gel electrophoresis of total leaf RNA extracts from plants infected with CNV transmission mutants Figure 2.2 Agarose gel electrophoresis of particles of CNV fungus transmission mutants  90 91  Figure 2.3 Location of mutated amino acids on the CNV subunit and trimer in CNV transmission mutants  99  Figure 2.4 Location of mutations in CNV fungus transmission mutants  93  Figure 3.1 Virus/zoospore binding assays  112  Figure 3.2 Effects of trypsin, sodium periodate and phospholipase C digestion on CNV binding to O. bornovanus zoospores Figure 3.3 Binding of CNV and CNV transmission mutants to total zoospore extracts using virus overlay assays  116  Figure 3.4 Virus overlay assays using CNV incubated with mannose, methyl a-D-mannoside or yeast mannan  118  Figure 3.5 Inhibition of CNV binding to O. bornovanus zoospores by several sugars  120  Figure 3.6 Labelling of zoospores with FITC-labelled Con-A and TPA  123  ix  114  Figure 4.1 Location of Pro73 and Pro85 on the CNV subunit and /3-annulus  133  Figure 4.2 Time-course of trypsin digestion of native, swollen and zoospore-bound CNV and Pro73Gly  142  Figure 4.3 Location of conserved proline residues in CP arm of several small spherical plant viruses  144  Figure 4.4 Summary of fungus transmission assays using Pro73Gly virions  148  Figure 4.5 Summary of Pro73Gly in vitro virus/zoospore binding assays  149  Figure 4.6 Agarose gel electrophoresis of native and swollen CNV and Pro73Gly  151  Figure 5.1 Model for CNV binding to O. bornovanus zoospores  170  Figure 6.1 Surface representation of CNV CP subunit (subunit-C) in different orientations showing location of mutated amino acids  174  Figure 6.2 Summary of transmission assays of CNV mutants  178  List of Abbreviations 3' 5' a A  A aa Ab Ala Arg ArMV Asn Asp ATF BYMV BNYVV BWYV bp BNYVV BRL BSA BYDV C Ca CaMV CCMV cDNA CLSV cm CMV CNV Con A CSBV CP C-terminal ++  CV.  CymRSV Cys D DAG DAS Dl  three prime five prime arm adenosine in the context of nucleotide sequence; alanine in the context of amino acid sequence angstrom amino acid antibody alanine arginine Arabis mosaic virus asparagine aspartic acid aphid transmission factor Barley yellow mosaic virus Beet necrotic yellow vein virus Beet western yellows virus base pair Beet necrotic yellow vein virus Bethesda Research Laboratories bovine serum albumin Barley yellow dwarf virus cytidine in the context of nucleotide sequence; cysteine in the context of amino acid sequence calcium Cauliflower mosaic virus Cowpea chlorotic mottle virus complementary DNA Cucumber leaf spot virus centimetre Cucumber mosaic virus Cucumber necrosis virus concanavalin A Cucumber soil borne virus coat protein carboxy-terminal cultivar Cymbidium ringspot virus cysteine aspartic acid aspartate-alanine-glycine motif direct antibody sandwich defective interfering  xi  DIC DNA ds DTT E EC EDTA ELISA ELBA ER F  differential interference contrast deoxyribonucleic acid double-stranded dithiothreitol glutamic acid effective concentration ethylenediaminetetraacetic acid enzyme-linked immunosorbent assay enzyme-linked binding assay endoplasmic reticulum phenylalanine  FHV  Flock house virus  Fig. FITC  figure fluorescein isothiocyanate  FMDV  Foot-and-mouth disease virus  g G  gram(s); genome in the cotext of viral RNA guanosine in the context of nucleotide sequence; glycine in the context of amino acid sequence galactose glycine-aspartate-aspartate glutamic acid glycine glycoprotein hinge haemagglutinin hydrochloric acid helper component protease  Gal GDD Glu Gly Gp or gp h HA HC1 HC-Pro HIV  Human immunodeficiency virus  hr I He ISVP K kb kDa Krf L  hour isoleucine isoleucine inter subvirion particle lysine in the context of amino acid sequence; thousand in the context of size kilobase kilodalton dissociation constant leucine  LBVV  Lettuce big vein virus  Lys [i m M MAb  lysine micro milli molar in the context of concentration; methionine in the context of protein sequence monoclonal antibody  MDMV  Maize dwarf mosaic virus  xii  ml mM min  millilitre millimolar minute(s)  MNSV  Melon necrotic spot virus  MP mRNA N NeuAc ng nt N-terminal oligo(s) ORF P  movement protein messenger RNA aspargine neuraminic acid nanogram nucleotide amino-terminal oligonucleotide(s) open reading frame proline in the context of protein sequence; protruding domain in the context of CNV CP  PAMV  Potato aucuba mosaic virus  PAT PAGE PBS PCR PDB  poorly aphid transmissible polyacrylamide gel electrophoresis phosphate buffered saline polymerase chain reaction protein data bank  PEBV  Pea early browning virus  PEG Phe  polyethylene glycol phenylalanine  PhMV  Physalis mottle virus  pK2/M5 PMSF  full-length infectious clone of CNV phenylmethylsulfonylfluoride  PMTV  Potato mop top virus  pmol Pro PTK PVDF PVR  picomoles proline proline-threonine-lysine polyvinylidene difluoride poliovirus receptor  PVY  Potato virus Y  Q R RAD RdRp RGD RNA(s) rpm RT RTD RT-PCR S  glutamine arginine RNA and arm domain in the context of antibody RNA dependent RNA polymerase arginine-glycine-aspartate motif ribonucleic acid(s) revolutions per minute readthrough readthrough domain reverse-transcriptase polymerase chain reaction serine in the context of protein sequence; shell domain in the context of CNV CP  xin  SBWMV SDS sec Ser sg SLRSV SqNV T TB TAE TBE TBRV TBS TBSV TBSV-Ch Thr TM TMV TNV TPA Tris Trp TRV TSWV TuMV TVMV Tyr U V Val VAP VP W WT Y ZYMV  Soil borne wheat mosaic virus  sodium dodecyl sulphate seconds serine subgenomic Strawberry latent ringspot virus  Squash necrosis virus thymidine in the context of nucleotide sequence; threonine in the context of protein sequence; triangulation number in the context of icosahedral virus structure Tris borate Tris acetate EDTA Tris borate EDTA Tomato black ring virus  Tris buffered saline Tomato bushy stunt virus  cherry strain of TBSV threonine transmembrane Tobacco mosaic virus Tobacco necrosis virus Tetragonolobus purpureas agglutinin  Tris-hydroxymethyl amino methane tryptophan Tobacco rattle virus Tomato spotted wilt virus Turnip mosaic virus Tobacco vein mottling virus  tyrosine uridine in the context of nucleotide sequence; units when referring to amount of enzyme volts in the context of electrophoresis; valine in the context of protein sequence valine viral attachment protein viral protein tryptophan wild-type tyrosine Zucchini yellow mosaic virus  xiv  Dedication To my parents for their love and support.  xv  Acknowledgements Firstly I would like to express my deep appreciation to my supervisor Dr D'Ann Rochon for her guidance and incredible patience during this important stage of my scientific career. I could not have imagined having a better advisor and teacher for my Ph.D study. Her sound advice, encouragement and support both personally and scientifically is greatly appreciated. I would like to thank the Natural Sciences and Engineering Council for financial assistance. Many thanks to my supervisory committee, Dr. Brian Ellis, Dr. Caroline Astell and Dr. Mary Berbee for their interest and valuable suggestions during the course of this study. I would like to thank the Pacific Agri-Food Research Centre, Agriculture and Agri-Food Canada for allowing me to use their lab facilities. I am also grateful to Michael Weis for his help with image analysis and microscopy. I wish to thank the members of Rochon lab: Dr. Marjorie Robbins for her input and initial training; Ron Reade for his valuable scientific and technical expertise pertaining to various aspects of this project; Steve Orban for his excellent technical assistance and proof reading. I would also like to thank Allison, Steve, Ron, Use, Liz, Neelima, Sreenivas, Joan, Les, Ray, Sandy, Suresh, Charlie, Feranando, Linda, Tammy, Sarah, Lauren and Pedro for their friendship. I would like to thank Rob Linning and John Bolton for introducing me to some excellent music. Many thanks to past and present members of the Rochon Lab for providing stimulating and fun environment in the lab and their tongue biting tolerance to my diverse and often times insanely loud music. I would like to thank Lynne Boyd and the library staff for their timely assistance. Lastly and most importantly, I must acknowledge the incalculable support of my family and Peggy Watson without whom this would not have been possible.  xvi  1 CHAPTER ONE LITERATURE REVIEW 1.1 Introduction Viruses, being obligate parasites, must depend on susceptible hosts for their survival in nature. Animal viruses spread in nature by utilizing the host cell surface molecules as their receptors. Plant viruses on the other hand, are unable to penetrate plant cells due to the presence of the plant cuticle and cell wall (83). Thus successful spread of the majority of the plant viruses is achieved through a vector that induces wounds in the plant host and also carries virus from one susceptible host to another. Most virus-host interactions are quite specific in nature. In animal viruses, the specificity lies in the ability ofthe virus to recognize a particular receptor on the host cell surface which primes the virus for subsequent steps in the infection process (14, 79, 140, 210). In plant viruses, however, the specificity is determined at least in part by the ability of the virus to recognize a component or components ofthe vector. The ability of a plant virus to recognize a specific vector is probably controlled via one or more receptors in or on the vector (163). Plant viruses are responsible for severe economic losses in agriculture worldwide (83). Although there is considerable information available on the biology of virus-vector relationships, very little is known about virus-vector interactions at the molecular level. Understanding the molecular mechanisms that govern the virus-vector recognition is an important part ofthe strategy to control plant virus diseases.  1  Cucumber necrosis virus (CNV) is a small icosahedral virus belonging to the family Tombusviridae, in the genus Tombusvirus. Vector transmission of CNV is facilitated by zoospores of the chytrid fungus Olpidium bornovanus (46). It is believed that CNV particles are adsorbed to the surface of the motile zoospores following independent release of virus and zoospores from roots of infected plants. Bound virus enters root cells upon zoospore encystment (34, 46). Previous work from our lab has shown that the CNV coat protein is important for fungus transmission (121). A single amino acid (aa) substitution in the shell domain (Section 1.5.4 and Figure 2.3) of the coat protein (CP) resulted in lowered transmission efficiency of CNV by O. bornovanus. In vitro binding assays demonstrated that this mutant bound zoospores less efficiently than wild type (WT) CNV indicating the role of specific regions on the CP shell domain in fungus transmission as well as zoospore attachment (153). In addition, it was shown that binding of CNV to zoospores is saturable suggesting that a specific number of recognition molecules exist on the zoospore surface for CNV attachment. Further in vitro binding studies demonstrated that binding between viruses and their respective vector zoospores was specific and reflective of their association in nature (154). Chytrid zoospores are known to contain a prominent cell coat. Cytochemical studies demonstrated that the cell coat is made predominantly of carbohydrates, especially glucosyl and/or mannosyl residues (141). Lectin binding studies from our lab suggested that glucose and/or mannose and fucose sugars exist on the surface of O. bornovanus zoospores (157) Plant virus-vector interactions are highly specific in nature (66, 67, 68, 139, 163). The specificity of the transmission process suggests the presence of specific recognition  2  molecules (receptors) on the vector surface for viruses (39, 163, 197). Plant virus-vector interactions may in some ways resemble animal virus/host interactions. Several animal viruses undergo extensive conformational changes as a part of their host cell entry process to expose the otherwise hidden fusion peptide (enveloped viruses) or membrane penetration proteins (nonenveloped viruses). These conformational changes are induced either by binding to the receptor or in endosomes at acidic pH (41, 48, 51, 140). Prior to the work described in this thesis there were no reports describing: 1) specific regions of a plant virus capsid involved in vector recognition or attachment; 2) the nature and biochemical characteristics of putative vector receptors for a plant virus; and 3) conformational changes induced in a plant virus capsid upon vector attachment or the importance of conformational rearrangements in vector transmission. This literature review is organized into five sections and provides background knowledge for the main areas of research covered in this thesis. The first section describes general aspects of plant virus transmission by arthropods, nematodes and fungi. The second section deals with primary and secondary cellular receptors for animal viruses with a special emphasis on the role of conformational changes in animal virus entry. In the third section, biological and molecular aspects of CNV are described. In the fourth and fifth sections, a brief overview of T=3 icosahedral plant viruses will be provided with a special focus on virus particle dynamics and the role of the CP Nterminal arm.  3  1.2 Plant virus transmission As mentioned earlier, there are no reported examples of plant virus entry by utilizing host cell surface molecules as their receptors. Infection of a new plant can only be initiated by entering the host cell via a wound. Plant viruses are transmitted in nature either mechanically by vegetative propagation, infected sap, seed or by specific association with specific invertebrate and fungal vectors (Table 1.1) (for a collection of recent reviews on vector transmission of plant viruses see 139, 68, 163). Vector transmission is a highly specific process in which the specificity is determined at different levels such as the type, family, genus, species and biotype of both the vector and the virus (83). Successful vector transmission requires: 1) acquisition - a specific association between the virus and the vector; 2) retention - the ability of the virus to remain infectious during vector association; and 3) inoculation - dissociation of the virus from the vector and subsequent infection of a new host. Understanding the concepts and terminology of plant virus vector transmission is valuable in epidemiological studies since it conveys an important property of the vector's ability to retain the virus. Watson and Roberts (204) proposed the first system of classification, which was based on early biological studies on aphid and leafhopper transmission. They classified viruses into two groups, non-persistent and persistent, based on the duration in which the virus remains inoculative in the vector (125). As our understanding of the biological and molecular aspects of vector transmission of plant viruses increased over the years, the classification of plant virus transmission has been revised and some new terms have been added. In this literature review, I have followed the terminology used by Gray and Banerjee (66) and Gray and Rochon (67). This  4  Table 1.1. Invertebrate vectors of plant virus genera and their mechanisms of transmission  1  Transmission Mechanism  Virus genera  Vector(s)  Noncirculative, nonpersistent  Fabavirus Potyvirus Carlavirus Cucumovirus Alfamovirus Machlomovirus Macluravirus  Aphids Aphids Aphids Aphids Aphid Thrips and beetles Aphids  Noncirculative, semipersistent  Caulimovirus Badnavirus Closterovirus Nepovirus Sequivirus Tobravirus Trichovirus Waikavirus  Aphids Mealybugs, leafhoppers Aphids, whiteflies, mealybugs Nematodes Aphids Nematodes Aphids, whiteflies, mites Aphids, leafhoppers  Circulative, nonpropagative  Enamovirus Geminivirus Luteovirus Nanavirus Umbravirus Bromovirus Comovirus Sobemovirus Tymovirus Rymovirus  Aphids Aphids, whiteflies, leafhoppers, treehoppers Aphids Aphids Aphids Beetles Beetles Beetles Beetles Mites  Bunyaviridae Marafivirus Reoviridae Rhabdoviridae Tenuivirus  Thrips Leafhopper Leafhoppers, planthoppers Aphids, leafhoppers, planthoppers Planthoppers  Circulative, propagative  This table was adapted and modified from reference # 66  5  terminology is based upon two important criteria: 1) the ability o f t h e virus to cross the vector cell membrane and subsequent internalization and; 2) virus retention time in the vector. The following are the different types o f transmission mechanisms observed i n viruses transmitted by invertebrate vectors.  Non-circulative  transmission  (externally  borne): In this type o f transmission, the  virus does not cross the vector cell membrane. Instead, it is carried externally on the surface o f the vector such as mouthparts. Non-circulative transmission can be conveniently divided into two types, semi- persistent and non-persistent. In both types the virus is associated specifically to either the cuticular lining ofthe mouthparts or the foregut o f the vector. The acquisition feeding time is very short (a few seconds to minutes), and the acquired virus is transmitted immediately b y the vector. In semipersistent transmission, the virus remains infectious for slightly longer periods in the vector.  Circulative  transmission  (internally  borne):  In circulative transmission, the virus  can cross the vector cell membrane and becomes internalized. A l l circulative viruses are retained b y the vector even after moulting. Acquisition and inoculation periods are usually longer (hours to days). Circulative transmission can be further divided into propagative  (replicative) and non-propagative  (non-replicative),  o f the virus to replicate i n the vector body upon internalization.  6  based upon the ability  The mechanisms of virus transmission by fungi and the terminology used are slightly different from virus transmission by invertebrates and will be discussed in the section 1.2.4.3.  1.2.1 Arthropod transmission Arthropods are the most important group of plant virus vectors, both in terms of the number of plant viruses transmitted and the economic importance of the disease caused. Arthropod vectors (197) transmit more than 70% of the viruses infecting plants. Approximately 99% of all arthropod vectors are insects (72). Among 29 orders that are described in the sub-phylum Insecta, the orders Homoptera, Coleoptera (beetles) and Thysanoptera (thrips) contain the important plant virus vectors. Over 70% of all insect vectors belong to the order Homoptera, which includes aphids, leafhoppers, whiteflies and mealybugs (83, 125). Mouthparts of these insects are of the piercing and sucking type, which is well-suited for the plant virus transmission. In addition, many of these insects are tissue-specific feeders often feeding on phloem, where many plant viruses are located (125).  1.2.1.1 Aphid transmission Aphids belong to the order Homoptera and are the most important group of plant virus vectors. Approximately 20% of the 4000 aphid species described are known to be vectors of plant viruses (125). The following are the three different stages of the aphid transmission cycle (83, 201): Acquisition phase. This refers to the phase in which the  vector aphid feeds on an infected plant to acquire a virus for subsequent transmission; Latent period. This is the period of time in which the virus is associated with the vector aphid before it can be transmitted; Retention time. This is the length of time during which the viruliferous aphid can transmit the virus to a susceptible host. The aphid transmission section of this literature review is described under three subcategories: 1) non-circulative and non-persistent; 2) non-circulative and semipersistent; and 3) circulative and non-propagative.  1.2.1.1.1 Non-circulative, non-persistent aphid transmission Most of the aphid transmitted plant viruses are transmitted in a non-persistent manner (83). In this type of transmission the virus is acquired rapidly by aphids after feeding on an infected plant for a very short time. The acquired vector transmits these viruses immediately, since the aphid rapidly loses (usually within hours) its ability to transmit the infectious virus (201). According to Pirone and Blanc, (137), the nonpersistent viruses can be divided into two types based on the molecular mechanisms involved: viruses following a capsid strategy and viruses a following helper component strategy.  1.2.1.1.1.1 Capsid strategy Several groups of viruses including cucumoviruses, alfamoviruses and carlaviruses do not need a helper component for successful aphid transmission. These viruses are grouped under the non-circulative, non-persistent viruses using the capsid strategy, since the virus particles sufficient for vector transmission. Much of the  8  information on the molecular mechanisms behind this type o f transmission comes from the work on cucumovirus transmission. Cucumber mosaic virus ( C M V ) is transmitted by a wide variety o f aphids (134). W i t h the help o f transcapsidation studies using a poorly transmissible C M V - 6 and a highly transmissible C M V - T , Gera et al., (60), showed that the coat protein contains the determinants for aphid transmission. Further transcapsidation studies by Chen and Francki, (43) showed that Tobacco mosaic virus R N A is aphid transmissible only i f it is encpasidated in vitro by an aphid transmissible Tomato aspermy virus strain V coat protein, confirming the role o f the C P as the sole determinant i n aphid transmission. A m i n o acid comparison o f the coat proteins ofthe highly transmissible F n y isolate o f C M V and the poorly transmissible M - C M V revealed that aa 129 and 162 are important for the aphid transmission (132). Repeated mechanical passage resulted i n a mutant that is transmissible by only one o f the two aphids that usually transmits C M V , indicating that different regions o f the viral coat protein influence the transmission o f C M V by different aphids (133). Recently L i u et al () reported that a conserved capsid surface domain o f C M V consisting o f a negatively charged /3H-/3I loop is important for aphid transmission. Mutations introduced i n this loop affected aphid transmission without grossly affecting virion structure or virus infectivity. L i u et al. predicted that the change i n the surface charge caused b y the mutations i n the /3H-/3I could be responsible for the transmission deficiency.  9  1.2.1.1.1.2 Helper strategy Viruses belonging to the genera Potyvirus, Caulimovirus,  and Waikavirus are  known to require a virus encoded helper component in addition to coat protein for successful vector transmission (65, 66, 67). Helper component is a non-structural protein that is believed to mediate the virus-vector interactions by forming a molecular bridge between the virus particle and its putative receptor in the aphid mouthparts (137). Much of our understanding of the molecular basis for helper dependent aphid transmission has come from poty- and caulimoviruses. The following section briefly describes the helper dependent aphid transmission of potyviruses. The potyviruses are filamentous particles made up of nearly 2000 subunits of the coat protein (145). In many potyviruses the coat protein has been implicated in aphid transmission. An N-terminal segment (30-90 aa in length) of the coat protein that is exposed on the particle surface, has been shown to be important for aphid transmission. In several potyviruses, mild treatment of virus particles with trypsin results in loss of this exposed N-terminal segment and a corresponding loss of aphid transmission (145). Based upon amino acid sequence comparisons of several aphid transmissible and non-aphid transmissible potyviruses, it was suggested that a conserved aspartate-alanine-glycine (DAG) motif located in the N-terminal region of the coat protein is involved in aphid transmission (116, 128, 136). Atreya et al., (8), provided direct evidence for the involvement of the DAG motif in aphid transmission. These studies showed that a substitution of Gly to Lys in the DAG motif of Tobacco vein mottling virus (TVMV) completely abolished aphid transmissibility. There are several reports indicating the pivotal role of the DAG motif in aphid transmission of potyviruses (8, 145). Recently it  10  has been reported that theflankingaa sequences of the DAG motif also play an important role in aphid transmission (9, 10). In vitro protein overlay studies have shown that in TVMV the DAG motif interacts with the viral encoded helper component (HC-Pro) (19). Govier and Kassanis, (64) first reported the role of the helper component in (HCPro) potyvirus transmission. They showed that aphid non-transmissible Potato virus-C (PVY ) and Potato acuba mosaic virus (PAMV, now classified as Potexvirus) are C  transmitted only when aphids are fed previously on aphid transmissible PVY-infected plant extracts. They also concluded that the helper component might act as a bridge between the aphids and the virus particles. Cell-free translation experiments provided the evidence that HC-Pro is of viral origin (80). Potyvirus helper component is a papain-like protease and Carrington et al., (37), showed thefirstindication of its proteolytic activity. Based upon amino acid sequence comparisons of the helper components of aphid transmissible and non-aphid transmissible isolates of PVY and PVY , it was suggested C  that a lysine-isoleucine-threonine-cysteine (KITC) motif may be important for helper activity (192). Later, the role of the KITC motif in the helper activity was reinforced by site-directed mutagenesis experiments (145). Based on in vitro protein overlay experiments and immunogold labelling studies it has been proposed that this KITC motif might be involved in binding to the aphid stylets (20, 131). Comparison of wild type (WT) and PAT (poorly aphid transmissible) strains of Zucchini yellow mosaic virus (ZYMV) revealed that a highly conserved proline-threonine-lysine (PTK) motif is also important for helper activity (99, 145). It has been shown that this PTK motif binds to the ZYMV CP in dot blot experiments, suggesting that the PTK motif may be a part of the virion-binding domain of HC-Pro (131).  11  1.2.1.1.2 Non-circulative, semi-persistent aphid transmission (Helper strategy) Semi-persistent transmission is very similar to non-persistent transmission except that the acquisition, inoculation and retention times are slightly longer. There is no conclusive evidence that molecular mechanisms involved in semi-persistent transmission are different from non-persistent transmission (21). Much of the information on semipersistent transmission comes from aphid transmission of caulimoviruses (for recent reviews see, 21, 70). Cauliflower mosaic virus (CaMV) is transmitted by the aphids Myzus persicae  and Brevicoryne brassicae. Like potyviruses, aphid transmission of CaMV involves helper components. Pirone and Megahead, (135), first demonstrated the requirement of the helper component. They showed that the purified virus was unable to be transmitted by aphids. Lung and Pirone (111) confirmed the presence of a helper component by transmitting the purified non-transmissible Campbell strain of CaMV using aphids that were fed first on a transmissible cabbage strain CaMV-infected plant. This helper component is called the aphid transmission factor (ATF). Several mutagenesis and gene replacement studies showed that the CaMV gene-II product (P2 protein) is the aphid transmission factor (7, 208). Later it was shown that the lack of aphid transmissibility of Campbell and CM1841 isolates of CaMV was due to a single aa substitution in P2 at residue 94 from glycine to arginine (71). Blanc et al (18) unequivocally demonstrated the role of P2 as the helper factor by transmitting several non-transmissible CaMV isolates using recombinant P2 protein produced in the baculovirus expression system. In addition, they showed that the aphids that were incubated with P2 still could not transmit purified CaMV particles, suggesting that still an additional transmission factor may be 12  needed. The CaMV gene III product (P3 protein) was identified as an additional aphid transmission factor (100). It was shown that the P2 incubated aphids were able to transmit purified CaMV only when they were also pre-incubated with bacterially expressed P3 protein. The presence of two transmission factors has prompted the idea that the bridge between the CaMV and the aphid mouthparts could be made of both P2 and the P3 proteins. A large C-terminal domain spanning aa 61-110 of P3 protein was shown to contain determinants for virion binding (101). In addition, the N-terminal region of P3 was shown to contain a binding domain for P2. The current proposed model for the aphid transmission of CaMV is: 1) P2 protein binds to the putative receptor in the aphid alimentary canal; and 2) the N-terminal region of the P3-virion complex that is formed in infected plants binds to the C-terminal region of P2; thus completing the bridge (21, 70).  1.2.1.1.3 Circulative, non-propagative aphid transmission Viruses that are transmitted in a circulative manner are internalized by the vector. The translocation of virus across the cell membranes inside the vector is very important for successful transmission. Circulative, non-propagative transmission of luteoviruses by aphids is one of the well-studied virus/vector interactions both at the biological and molecular level. The following section describes recent advances in luteovirus-aphid interactions.  Vector determinants: The general circulative pathway is similar for both propagative and non-propagative viruses. Virus is acquired via the food canal and then  13  released into the haemocoel, after passing through the foregut, midgut and hindgut. Eventually the virus must be associated with the salivary glands to be transmitted to the plant host (66). Based on ultrastructural studies, it has been proposed that receptormediated endocytosis and exocytosis may be involved in the virus crossing the barriers of the aphid gut and salivary gland epithelial linings (197). It has been reported that the determinant for vector specificity does not lie at the gut barrier, since different luteoviruses are acquired into the haemocoel irrespective of whether the carrying aphid is a vector or not (61, 66). The haemocoel acts as a reservoir in which acquired viruses remain infective without undergoing replication. Following acquisition and entry into the haemocoel, the virus diffuses through the haemolymph until it encounters the aphid's salivary glands (26). Luteoviruses are exclusively associated with the accessory salivary gland (66, 68). The specificity is believed to be determined at the level of the accessory salivary gland (62). Recently Li et al., (103) have identified the possible receptors for Barley yellow dwarf virus (BYDV) in its vector aphid Sitobean avenae. With the help of virus overlay assays they showed that BYDV binds to two proteins (33 kDa and 50 kDa) from the head tissues of the vector aphid Sitobean avenae; but not from the non-vector aphid Rhopalosiphum padi. Anti-idiotypic antibody produced against a monoclonal antibody (MAV-4) of BYDV-MAV was able to bind several proteins including the 33 and 50 kDa proteins from the head tissue of Sitobean avenae. These data support the presence of receptor or receptor-like molecules in the aphid salivary gland for the virus.  14  Viral determinants: Rochow (164) showed the involvement of the coat protein in aphid transmission using virus transcapsidation studies on MAV and RPV isolates of BYDV. The aphid R. padi transmits the RPV isolate but not the MAV isolate of BYDV. During co-infection in a single plant, virus containing the MAV genome encapsidated in RPV coat protein could be transmitted by R. padi. MAV replication was detected in the new host plant, but the virus purified for this infection (MAV genome encapsidated by MAV coat protein) could not be transmitted by R. padi. The importance of the coat protein in luteovirus transmission also comesfromstudies of transmission of umbraviruses. Umbraviruses do not encode a coat protein (182) and are transmitted by aphids only if plants are co-infected by a luteovirus which provide the coat protein necessary for transmission. The type of luteovirus providing the coat protein determines which aphid will transmit the umbravirus (125). The 24 kDa major coat protein of luteoviruses is encoded by ORF-3. Sequence analysis showed that there is a larger ORF (ORF-5) following ORF-3. Due to occasional translational suppression of the coat protein stop codon, ORF-5 is expressed, resulting in a 74 kDa readthrough (RT) protein. The RT protein is exposed on the particle surface and is not necessary for particle assembly (42, 92, 151), providing evidence for the requirement ofthe RT protein in aphid transmission. Moreover, a mutant of a PAV isolate of BYDV that does not have RT protein, is not aphid transmissible. Sequence comparisons of the RT proteins of several luteoviruses revealed that the N-terminal region is highly conserved, whereas the C-terminus is variable. Purified preparations of several luteoviruses that are aphid transmissible revealed that the RT protein is present in a truncated form with its C-terminal variable region being proteolytically cleaved in vivo.  15  These studies indicate that the C-terminal region of the BYDV RT protein is not important for aphid transmission (202). In addition, mutant BYDV particles that do not contain the N-terminal region of the RT protein are not aphid transmissible (53, 68). The N-terminal region of the RT protein contains important determinants for the aphid transmission. Several point mutations in the N-terminal region of the RT protein of Beet western yellows virus (BWYV) resulted in reduced transmission (25). Moreover, it was shown that aphid transmission of the RPV isolate of BYDV could be prevented by mixing the purified virus with the antibodies raised against the N-terminal region of the RT protein before aphid feeding, whereas antibodies raised against C-terminal region RT protein did not prevent transmission (119). Virions that did not contain the RT protein were able to be taken up by aphids and were observed in haemocoel, indicating that the coat protein contains sufficient determinants to cross the aphid gut barriers to reach the haemocoel (42, 66,148). It also suggests that the role of the RT protein probably lies in facilitating the virus entry from the haemolymph to the accessory salivary gland (66).  Role of endosymbiotic bacteria in aphid transmission: Aphids harbour  endosymbiotic bacteria that belong to the genus Buchnera in their haemocoel. The bacteria provide essential amino acids that are not synthesized by the aphids. In addition, they also produce large quantities of symbionin, a chaperonin protein, which is a homologue of E.coli chaperonin Gro-EL (15). The chaperonins are a class of proteins that play an important role in protein folding and translocation across membrane barriers (See 66). It has been shown that symbionin binds to purified luteoviruses and the RT proteins in vitro (54). In addition the N-terminal half of the RT protein of BWYV was shown to  16  bind to symbionin. (196). It has been suggested that the RT protein on the viral capsid is involved in binding to symbionin and thereby helping the virus to move from the aphid haemolymph to the accessory salivary gland. However, the role of symbionin in vector specificity has not yet been proven, since luteoviruses bind to symbionin from both vector and non-vector aphid species with relatively similar affinities (196).  1.2.2 Thrips transmission Viruses in the genus Tospovirus are transmitted by thrips in a circulative and propagative manner. Tomato spotted wUt virus (TS WV) is the type member of this genus and much of the information (molecular and biological) on transmission comes from TSWV and its vector Franklineilla  occidaentalis. At least 10 species of thrips have been  reported to transmit tospoviruses (194). TSWV and its vector, F. occidaentalis have a unique relationship, in that the larvae but not the adults must acquire the virus in order to be transmitted. Virus acquisition capacity rapidly declines as immature larva develop into adults (195). Four potential barriers have been recognized for the virus to cross in its thrips vector: 1) entry into the midgut; 2) escape from the midgut; 3) entry into the salivary gland; and 4) escape from the salivary gland (194). TSWV has been reported to be associated with cellular membranes both in its thrips vector and the plant host wherein the viral glycoproteins, GP1 and GP2 are involved. Immunolocalization experiments have shown that TSWV is associated with the Golgi complex in thrips and with the endoplamic reticulum (ER) in the plant cell (58, 193). Several lines of evidence show that GP1 and GP2 are the viral attachment proteins in the thrips body. Electron microscopic  17  studies revealed that the TSWV GP1 binds to the apical membrane in the midgut of the thrips larvae. The GP2 contains an arginine-glycine-aspartic acid (RGD) motif, which is a highly conserved motif among several animal virus attachment proteins and known to play an important role in recognizing viral receptors (197). Recently, with the help of gel overlay assays, Bandla et al, (12) showed that both GP1 and GP2 bind to a 50 kDa protein in total thrips protein extracts. Viral GPs did not recognize the same band in similar assays using non-vector and adult thrips, which reflects the virus-vector association in nature. Anti-idiotypic antibodies to the viral GPs selectively bind to 50 kDa protein in western blots, indicating the specificity ofthe interaction. Electron microscopic studies showed the association of anti-idiotypic antibodies of viral GPs with the larval midgut membrane. This data suggests that the GP1 and GP2 are viral attachment proteins. In addition, it also suggests that the 50 kDa protein is a potential receptor in thrips larva for TSWV. Viral GP2 also binds to a 94 kDa protein, which is abundant in the thrips body, but is not in the midgut. Moreover this 94 kDa protein was recognized in both vector and non-vector thrips (96). It has been suggested that this protein may be involved in circulation ofthe virus in the thrips vector.  1.2.3 Nematode transmission Some nematodes are root-infecting ectoparasites of plants. Plant viruses belonging to Nepo- and Tobravirus genera are transmitted by nematodes. Currently 12 nepoviruses and three tobraviruses are transmitted by nematodes belonging to the families Longidoridae  and Trichodoridae respectively (114). Species within the genus  Longidorus, Paralongidorus  and Xiphinema of the family Longidoridae  18  and species  within the genus Paratrichodorus  and Trichodorus ofthe family Trichodoridae are  identified as vectors of plant viruses. Virus transmission by nematodes is ofthe noncirculative and semi-persistent type, since: 1) virus is retained in the vector for several weeks; 2) viruses do not replicate in the vector; and 3) virus is not retained after moulting (114, 115). Based on specificity studies, Brown and Weischer (30) proposed two concepts in nematode transmission: 1) exclusivity, in which the vector nematode species transmits only one virus or one serologically distinct virus strain; and 2) complementarity, in which the vector nematode species transmits more than one virus or serologically distinct strains of a virus. Successful transmission of virus by nematodes depends on five phases. They are acquisition, adsorption, retention, release and transfer of the virus. The specificity of the virus and nematode interaction probably lie at the adsorption, retention and release phases of the transmission process (114).  Viral determinants: Tobra- and nepoviruses are positive-sense, single stranded RNA viruses with two genomic RNAs, a larger RNA1 and a smaller RNA2, which are encapsidated into separate virus particles (113). Tobravirus particles are rod-shaped and nepovirus particles are spherical. Pseudo-recombination experiments involving viral genomic RNAs from differentially transmissible isolates of the same virus have shown that RNA2 of both tobra- and nepoviruses contain important determinants for vector transmission (73, 138). RNA2 of nepovirus encodes a coat protein along with two other proteins called 2A and 2B. Gene replacement studies between Grapevine fanleaf virus, transmitted by X. index, and Arabis mosaic virus (ArMV), transmitted by X. diversicaudatum have shown  19  that 2A and 2B proteins are not necessary for transmission, indicating the CP as a sole determinant of vector specificity (16). RNA2 of tobraviruses also produces a coat protein along with three other proteins, 2B, 2C and a small 9 kDa protein. The C-terminal 20 to 30 aa of the coat protein is exposed on the surface of the particle and is relatively unstructured (22). Deletion of 15 aa in this region has resulted in loss of transmission in Pea early browning virus (PBEV) isolate Ppa56 and Tobacco rattle virus isolate PpK20. Replacement of the coat protein gene of a poorly transmissible SP5 isolate of PEBV with the coat protein gene of the highly transmissible Ppk20 isolate of TRV, did not result in high transmissibility, indicating the role of other proteins encoded by RNA2 in vector transmission apart from the coat protein. Deletion of the 2C gene reduced transmission in PEBV, but not in TRV (81). Deletion of 2B from both PEBV and TRV resulted in loss of vector transmission (81, 112). In addition, it has been shown that the 2B protein can act in trans so that a TRV transmission mutant lacking the 2B gene can be transmitted when co-inoculated with WT TRV (198). Yeast two-hybrid and immuno-gold labelling studies have shown that the 2B protein interacts with the coat protein C-terminus, probably acting as a helper component (200). Moreover, a single aa substitution in the PEBV 2B gene completely abolished vector transmission. A predicted coiled-coil region of 2B protein is speculated to be involved in the interaction with CP (199). It has been suggested that the 2B protein might form a bridge between TRV and the oesophageal lining of nematode mouthparts. Also, the specificity of tobravirus transmission could be determined by the 2B protein (115).  20  Vector determinants: The site of virus retention in the vector nematode mouthparts varies either from the inner surface of the odontostyle, the region between odontostyle and cuticular lining of the guiding sheath or the oesophageal tract depending on the vector species. (114). A discontinuous layer of carbohydrate staining material was observed on the oesophageal tract of X. diversicaudatum and  Paratrichodorus  pachydermus (29). Adsorption of ArMV and Strawberry latent ringspot virus (SLRSV)  particles was found only in the presence of carbohydrate lining material in the oesophageal tract of X. diversicaudatum  (155). These studies provide some preliminary  evidence for the involvement of specific recognition molecules in nematode transmission. It has been suggested that the nepovirus CP may have lectin-like properties in recognizing carbohydrates (29).  1.2.4 Fungus transmission The fungi that are known to transmit plant viruses are root-infecting, zoosporic, obligate parasites on plants (34). Two species belonging to Phylum Chydridiomycota (Olpidium bornovanus and Olpidium brassicae) and three species belonging to the plasmodiophorids (Polymyxa graminis, P.betae and Spongospora subterranea) are  recognized as vectors of several plant viruses (Table 1.2 and 1.3) (4, 34,163). Plasmodiophorids were considered as fungi for a long time. But recently, based on molecular studies, their taxonomic status has been changed. They are now considered to be protozoans (24, 203). Although chytrids and plasmodiophorids are taxonomically different, they share several characteristics with regard to their life cycles: 1) both are  21  zoosporic and obligate parasites of plants; 2) survival and infection occurs in nature via resting spores; 3) both are holocarpic (i.e., the entire thallus is converted into a zoosporangium) (34). Because of the similarities in their life cycles and their relation to virus transmission, several reviews discussed chytrids and plasmodiophorids as fungal vectors of plant viruses (2, 4, 34). However, they are now referred to as fungal and plasmodiophorid vectors of plant viruses.  1.2.4.1 Zoospore structure and the life cycle of Olpidium spp. O. bornovanus and O. brassicae are obligate, root-infecting parasites, which are extremely common in nature. These are symptomless parasites on the host but, are economically important because of their ability to act as vectors for several important plant viruses (163). Much of the information on the structure ofthe Olpidium zoospore and life cycle (Figure 1.1 and 1.2) comes from the work on O. brassicae by Temmink and Campbell (191). O. brassicae zoospores are small, about 2-3 um in diameter, with an oval shaped body surrounded by a plasmalemma (188). Zoospores of O. bornovanus are also oval shaped but are 6-7 um, slightly larger than that of O. brassicae. Possession of a single posterior, whiplash flagellum is a typical characteristic of Olpidium species. A membranous sheath that is continuous with the plasma membrane of the zoospore body also covers the flagellum. Zoospores contain a prominent nucleus, several mitochondria, vacuoles, multivesicular bodies and lipid globules. Endoplasmic reticulum is distributed throughout the zoospore body as short tubules, and ribosomes are present in high number (189). Several chytrid zoospores are known to contain a  22  Figure 1.1. Schematic representation o f the ultrastructure o f the O.brassicae  zoospore*.  Abbreviations used: A , axoneme; A S , axonemal sheath; E R , endoplasmic reticulum; K , kinetosome; L , lipid globules; M , mitochondrion; M V B , multivesicular body; N , nucleus; R, rhizoplast; R V , rhizoplast vesicle; V , vacuole; Z E , zoospore ectoplast. * This diagram was adapted from reference # 190.  23  prominent cell coat (45). The cell coat is present on the flagellum as well as on the zoospore body. Cytochemical studies have demonstrated that the cell coat is predominantly made of carbohydrates, especially glucosyl and/or mannosyl residues (141). Primary zoospores are producedfromresting spores upon favourable environmental conditions. These primary zoospores swim to the host plant root hairs, where they encyst. During encystment the flagellum is projected awayfromthe host surface. Chytrid zoospores become sticky during encystment, indicating a change in the composition of cell surface material (141). Upon encystment, the zoospore retracts its flagellum and in several chytrids, it has been suggested that the axoneme coils and is released into the zoospore cytoplasm. The axonemal sheath wraps around the zoospore body and ultimately fuses with the plasma membrane (141). In the case of O. brassicae, Temmink, (191) proposed a "reeling in" mode of flagellar retraction in which, the axonemal sheath along with the axoneme is released into the zoospore cytoplasm. Once theflagellumis retracted, a cyst wall appears around the zoospore body and a papillum develops in between the host cell and the cyst. Then, a vacuole appears in the cyst and the zoospore cytoplasm along with surrounding ectoplast move into the host cell via the papillum (190). The thallus increases in size as it undergoes mitotic divisions becoming a mature zoosporangium. A cell wall appears around the mature thallus and it is therefore believed that virus transmission to the root cell must occur before the thallus wall is formed (188). As thalli mature, zoospores are formed inside and are released to the outside through exit tubes upon contact with moisture. During unfavourable conditions the thalli may develop into resting spores.  24  Root cell penetration  zoosporangia  Attachment and encystment on roots  1  zoospore release  zoospores  resting spores  Figure 1.2. Life cycle o f O. brassicae (adapted from reference #191)  25  1.2.4.2 Life cycle of plasmodiophorids The plasmodiophorid life cycle consists of two phases: 1) the primary or sporangial phase, which results in zoosporangia and the release of secondary zoospores; and 2) the secondary or sporogenic Plasmodium, which produces thick-walled resting spores (24, 95). All plasmodiophorids zoospores are biflagellate with anteriorly located whiplash flagella (6). The resting spores of plasmodiophorids are called sporosori. Upon favourable conditions, these sporosori germinate and release primary zoospores. The primary zoospores swim to the host root hair or epidermal cells where they encyst. A tubular structure called "Rohr" that contains a dagger-like body called a "Stachel" is formed within the cyst. Later the contents of zoospores along with the Rohr and the Satchel are injected into the host cell. Inside the cell, the contents of primary zoospores undergo cruciform nuclear divisions to form a multinucleate Plasmodium. This Plasmodium can develop into either ofthe two phases described above. During the sporangial phase, the nucleus undergoes several cycles of non-cruciform mitotic divisions resulting in an aggregate of zoosporangia. Later these zoosporangia develop exit tubes to release secondary zoospores either to the outside of the root or into adjacent root cells. During the sporogenic phase, the nucleus undergoes non-cruciform mitotic divisions before forming unicellular thick-walled resting spores.  1.2.4.3 Modes of fungus transmission Campbell, (31) introduced terminology to fungus-transmitted viruses. It is slightly different from the terminology used for invertebrate transmitted viruses because ofthe differences in life cycles of fungi and invertebrates. The terminology was based upon  26  virus acquisition and location of virion relative to resting spores. The in vitro mode of transmission involves virus acquisition by vector outside the host and the virus is not located within the resting spore. In the in vivo mode of transmission, virus is acquired by vector within the infected plant and the virus is persistent in resting spores. Later the terms non-persistent (externally borne) and persistent (internally borne) were suggested to align the terminology with that of aphid transmission (2, 4, 187). Non-persistent transmission was applied viruses that were carried externally on the surface of the zoospores (in vitro). Whereas, the term persistent transmission was used when virus was acquired and carried internally (in vivo). In this thesis I have followed the terminology used by Campbell (31).  1.2.4.3.1 In vitro transmission Evidence for this type of transmission comesfromthe work on TNV transmission by O. brassicae (191). Virus can be transmitted by mixing purified virus from infected crude sap with virus-free zoospores. Virus-free zoospore cultures can be obtained by air drying the doubly-infected roots for several weeks or months during which time virus loses infectivity but resting spores retain the capacity to initiate fungus infection. Virusfree culture can also be obtained by treating roots containing resting spores with 20% trisodium phosphate (Na PC>4) or 5N HC1 prior to root inoculation. Viruses transmitted in 3  the in vitro manner lose infectivity following this treatment and were suggested to be outside rather than inside the spores (34). Studies showed that adding virus specific antisera to virus/zoospore suspensions immediately upon virus/zoospore mixing could prevent transmission of virus. If antiserum was added much later (after 10-15 minutes)  27  transmission of virus could not be prevented. This indicated that virus adsorption to zoospores is occurring within 5 to 10 minutes of mixing (34, 163). As mentioned earlier, two species of Olpidium (O. bornovanus and O. brassicae) have been shown to transmit viruses by the in vitro method (Table 1.2 and 1.3). All the viruses that are known to be transmitted by in vitro method are isometric particles belonging to the family Tombusviridae (Table 1.2; 63). Under natural conditions, virus and vector zoospores are released independently into the soil from the roots of infected plants. Virus particles are then adsorbed onto the zoospore plasmalemma and flagellar sheath. Upon flagellar retraction, virus is believed to enter the zoospore cytoplasm along with the flagellar sheath (181, 191).  Specificity: Virus and zoospore interactions are very specific and there is a direct correlation between the ability of the virus to adsorb to the zoospore surface and transmission efficiency. Electron microscopic studies (191) have shown that TNV, which is transmitted by O. brassicae zoospores, adsorbed to O. brassicae zoospore more efficiently than it did to non-vector O. bornovanus zoospores. Transmission of TNV by O. brassicae zoospores also depends on the isolate of fungus. Lettuce and tomato isolates of O. brassicae transmit TNV, whereas the mustard isolate does not. CNV adsorbed onto O. bornovanus zoospores more efficiently than that of non-vector O. brassicae zoospores (191). Moreover different O. bornovanus isolates differ in their ability to transmit Cucumber leaf spot virus (CLSV), Melon necrotic spot virus (MNSV), Cucumber soil borne virus (CSBV), Squash necrosis virus (SqNV) and CNV (33). These specificity  28  Table 1.2. Viruses transmitted by Olpidium vectors Virus  Acronym  Genus  Fungal vector  Acquisition mode  Tombusviridae Cucumber necrosis virus Cucumber leaf spot virus Cucumber soilborne virus Melon necrotic spot virus Squash necrosis virus Red clover necrotic mosaic virus Chenopodium necrosis virus Lisianthus necrosis virus Tobacco necrosis virus-A Tobacco necrosis virus-D  CNV CLSV CSBV MNSV SqNV RCNMV ChNV LNV TNV-A TNV-D  Tombusvirus Aureusvirus Carmovirus Carmovirus Carmovirus Dianthovirus Necrovirus Necrovirus Necrovirus Necrovirus  O. bornovanus 0. bornovanus 0. bornovanus 0. bornovanus O. bornovanus 0. bornovanus 0. brassicae 0. brassicae O. brassicae 0. brassicae  In In In In In In In In In In  vitro vitro vitro vitro vitro vitro vitro vitro vitro vitro  Viruses in unassigned families Miration lettuce virus Tulip mild mottle mosaic virus Freesia leaf necrosis virus Lettuce big vein virus Lettuce ring necrosis virus Tobacco stunt virus  MiLV TMMMV FLNV LBVV LRNV TSV  Ophiovirus Ophiovirus Varicosavirus Varicosavirus Varicosavirus Varicosavirus  0. brassicae O. brassicae O. brassicae 0. brassicae 0. brassicae 0. brassicae  In In In In In In  vivo vivo vivo vivo vivo vivo  This table was adapted from reference # 163  29  Table 1.3. Viruses transmitted by plasmodiophorid vectors  1  Genus  Fungal vector  Acquisition mode  Potyviridae Barley mild mosaic virus BaMMV Barley yellow mosaic virus BaYMV OMV Oat mosaic virus Rice necrosis mosaic virus RNMV Wheat spindle streak mosaic virus WSSMV WYMV Wheat yellow mosaic virus  Bymovirus Bymovirus Bymovirus Bymovirus Bymovirus Bymovirus  P. P. P. P. P. P.  graminis graminis graminis graminis graminis graminis  In In In In In In  vivo vivo vivo vivo vivo vivo  Viruses in unassigned families Beet necrotic yellow vein virus Beet soilborne mosaic virus Indian peanut clump virus Peanut clump virus Chinese wheat mosaic virus Oat golden stripe virus Rice stripe necrosis virus Soilborne cereal mosaic virus Soilborne wheat mosaic virus Sorghum chlorotic spot virus Potato mop top virus Beet soilborne virus Beet virus Q  BNYYV BSBMV IPCV PCV CWMV OGSV RSNV SBCMV SBWMV SrCSV PMTV BSBV BVQ  Benyvirus Benyvirus Pecluvirus Pecluvirus Furovirus Furovirus Furovirus Furovirus Furovirus Furovirus Pomovirus Pomovirus Pomovirus  P. betae P. betae P. graminis P. graminis P. graminis P. graminis P. graminis P. graminis P. graminis P. graminis S. subterranea P. betae P. betae  In In In In In In In In In In In In In  vivo vivo vivo vivo vivo vivo vivo vivo vivo vivo vivo vivo vivo  Unclassified viruses Aubian wheat mosaic virus Watercress yellow spot virus  AWMV WYSV  Tombusvirus?  P. graminis S. subterranean f.sp. nasturtii  In vivo Not known  Virus  Acronym  This table was adapted from reference # 163  30  studies strongly suggest the involvement of receptor or receptor-like molecule(s) on the zoospore surface (2, 34, 154, 163)  Viral determinants: Based on specificity studies on viruses that are transmitted via the in vitro fashion, it was speculated that specific receptors on Olpidium zoospores are involved in recognizing the viral coat protein (2, 191). MacLean et al., (121) first demonstrated the involvement of coat protein in the in vitro mode of transmission. Reciprocal exchanges of CP genes were made between infectious cDNA clones of the non-transmissible cherry strain of Tomato bushy stunt virus (TBSV-Ch) and transmissible CNV. Virions containing the TBSV-Ch genome encoding the CNV CP were found to be readily transmitted by O. bornovanus. However, transmission did not occur using virions containing the CNV genome encoding the TBSV-Ch CP. Robbins et al., (153) provided further evidence of the involvement of the coat protein in CNV transmission by O. bornovanus. A naturally occurring CNV transmission defective mutant (LL5) was isolated by repeated mechanical passage of CNV on Nicotiana clevelandii. Further sequence analyses showed that the loss of transmissibility was due to a single amino acid substitution (glutamic acid (E) to lysine (K)) in the CP shell domain (see Chapter 2). Further, with the help of in vitro virus/zoospore binding assays, it was concluded that the poor transmissibility of LL5 was at least partially due to its inability to bind to vector zoospores efficiently. Amino acid sequence comparisons of the CPs of several viruses of Tombusviridae have shown that viruses transmitted by O. bornovanus are more closely related to each other than they are to the genus to which they belong (163).  31  Vector determinants: Robbins (154) provided some preliminary evidence for the presence of receptor or receptor-like molecules on the surface of O. bornovanus zoospores for CNV. These studies showed that virus/zoospore binding is specific and saturable suggesting typical receptor mediated interactions. In vitro binding studies showed that CNV, CLSV and MNSV, which are transmitted by O. bornovanus, bound to O. bornovanus zoospores more efficiently than TNV, which is transmitted by O. brassicae. Similarly, TNV bound to O. brassicae zoospore more efficiently than CNV, CLSV and MNSV (see Chapter 3). In vitro binding studies have also shown that CNV/zoospore interactions are time and pH dependent (154). In addition, FITC labelled lectin binding studies have shown the presence of glucose/mannose and fucose on the surface of O. bornovanus and O. brassicae zoospores. It was speculated that these sugars could be a part of the putative viral receptor on the zoospore surface (154).  Vims delivery to host plant: As mentioned earlier, it has been postulated that virus bound to the surface of zoospores enters the zoospore cytoplasm along with the flagellar sheath and axoneme during flagellar retraction as virus particles have been observed between the whorls of flagellar sheath membrane inside the zoospore cytoplasm (181). In order for virus to infect a root cell it must cross the plasma membrane of the young fungal thallus (before it develops a thick wall) (188). It is intriguing that, even though the virus can be found in the zoospore cytoplasm, it is unable to be incorporated into resting spores. Two possible reasons suggested by Rochon et al, (163) are: 1) during co-infection, virus may not be present in the same tissue as the resting spores develop and; 2) Virus may not be able to cross the membrane of the thallus from the root cell  32  cytoplasm. More experimental study is needed in this area of the in vitro mode of transmission.  1.2.4.3.2 In vivo transmission In the in vivo or internally-borne mode of transmission, virus is acquired during vector development inside the host plant. Virus is also carried inside the resting spore, and presumably also within zoospores (4, 34, 163). One Olpidium species (O. brassicae) and three plasmodiophorids (P. graminis, P. betae and S. subterranea) are reported to transmit plant viruses in the in vivo fashion. All viruses that are known to transmit in this mode are either rod-shaped or filamentous particles with multipartite genomes (Table 1.3; 163). Viruses belonging to Ophio- and Varicosavirus are transmitted by O. brassicae. Viruses belonging to the Bymo-, Beny-, Porno-, Furo and Pecluvirus  genera are  transmitted by plasmodiophorids ( Table 1.3; 63). Several lines of evidence are available to show the persistence of virus inside the resting spores. Lettuce big vein virus (LBVV) was shown to remain for long periods in air dried as well as chemically treated resting spores (31, 32). Persistence of several other viruses in their respective vector resting spores has been reported (4). Immuno-gold labelling studies have provided the evidence for the presence of Barley mild mosaic virus (BaMMV) in zoospores and zoosporangia of P. graminis (38). Similar results were obtained in Beet necrotic yellow vein virus (BNYVV)//*. betae infected plants (4). There are very few reports providing direct evidence for the presence of virus in resting spores, probably because of technical difficulties, such as the thick cell wall of resting spore and low number of viruses per spore (163). Filamentous virus-like bundles of Barley yellow  33  mosaic virus (BYMV) were observed in resting spores of P. graminis (39). Potato mop top virus (PMTV) -like particles were observed in fractured resting spores of its vector S. subterranea (123). Recently Driskel and Verchow (50) provided the evidence for the presence of the movement protein and viral RNA, but not the coat protein of Soil-borne wheat mosaic virus (SBWMV) in resting spores of P. graminis. It has been suggested that the viral RNA/movement protein complex may be the infectious component (50, 163).  Vector determinants: The role of viral encoded proteins in plasmodiophorid transmission has been well studied. BNYVV, which is transmitted by P. betae, has five RNA components. RNA4 and RNA2 have been shown to be important for fungus transmission (183, 184). RNA2 encodes the coat protein as well as a CP readthrough (RT) product. The RT domain is thought to play an important role in particle assembly and its C-terminus is probably exposed on the particle surface (171). Tamada and Kusume (184) have reported that serial passage of BNYVV in the absence of vector has resulted in deletions at the C-terminal end of the RT domain, which in turn, resulted in complete loss of fungus transmission, indicating the RT is essential for vector transmission. Further work has shown that a KTER motif located at the N-terminus of the RT domain is important for fungus transmission (185). PMTV also produces a CP RT which has been shown to be essential for its transmission by S. subterranea (147). Readthrough proteins of several other viruses were also shown to be important for fungus transmission (4, 163). A 39 kDa protein expressed by Peanut clump virus (PCV) RNA2 and a P2 protein encoded by bymovirus RNA-2 are reported to be essential for fungus transmission (117).  34  Unlike RT proteins, these two proteins are expressed independently from the coat protein. The P2 protein was also shown to be associated with inclusion bodies in infected tissues. It has been suggested that these two proteins might act as helper components bridging the CP and vector as in aphid transmission of poty and caulimoviruses (163). Adams et al, (3) have recently identified two trans-membrane domains (TM1 and TM2) in the CP-RT of beny-, furo- and porno viruses genera and in the P2 protein of bymo viruses. The TM2 is either absent or disrupted in naturally occurring deletion mutants that are not fungally transmitted. A strong role for these two trans-membrane domains has been suggested in translocation of virus from the host cell cytoplasm to fungal plasmodia (3).  1.3 Virus/receptor interactions Most animal, insect and bacterial viruses require attachment to the host cell surface for successful infection. This attachment is mediated by a viral attachment protein (VAP) (either envelope glycoprotein or one or more capsid proteins) and a host cell surface molecule (s) acting as virus receptor (14, 79, 118, 165, 210,). Similarly, many plant viruses may require cellular receptors on their vectors for successful transmission to susceptible hosts (163, 197). Recognition of a cellular receptor by a virus is the first and crucial step in the infection process. Binding to the host cell surface receptor usually results in either direct fusion between the viral lipid envelope and host cell plasma membrane, receptor-mediated endocytosis or conformational changes in the viral capsid, all of which lead to the transfer of the viral genome or viral nucleoprotein into the host cell (140, 210). The nature, number and expression of host cell receptors play an important role in determining host range and tissue tropism (102). Virus/receptor  35  interactions are often complex and are often followed by extensive structural rearrangements in either the VAP or the viral capsid. Understanding the structural details of virus/receptor interactions has considerable significance in terms of designing drugs that inhibit virus entry (14). Virus particles are known to attach to cell surfaces non-specifically through electrostatic interactions. Viruses can also adhere non-specifically to many substances including inert materials (186). In order to distinguish between receptor mediated attachment and non-specific interaction, Tardieu et al., (186) have proposed three standard criteria: 1) Binding of viruses to the host cell must be saturable, indicating a discrete number of binding sites; 2) virus binding to cells that do not contain specific binding component(s) should not be saturable; and 3) unlabelled binding of virus should be competitively inhibited by labelled virus or by closely related viruses (assuming they share the same receptor).  1.3.1 Virus receptors Viruses take advantage of a wide variety of cell surface molecules as their receptors. A virus receptor can be defined as the host cell surface component(s), that interacts specifically with the virus and facilitates viral entry and subsequent infection (79, 176). Several virus receptors have been recognized and characterized (Tablel.4 and 1.5). The receptors belong to different families of proteins, carbohydrates and lipids that are involved in important cellular functions such as signal transduction, cell adhesion, immune modulation and enzymatic activities (14). Viral receptors can be identified by various techniques. These include the use of specific monoclonal antibodies raised  36  against the cell surface molecules, gene transfer techniques, mutant cell lines, virus overlay assays and the use of carbohydrate digesting enzymes (210). A comparison of viruses and their cellular receptors reveals no apparent correlation between the virus family and receptor structure and function (14). For example, viruses belonging to Picornaviridae  share common structural features in their  capsids, but they use different molecules such as integrins, glycoproteins of the immunoglobulin superfamily, decay accelerating factor (CD55) and sialic acid as their receptors (Table 1.4 and 1.5). Sometimes a virus like HIV may use different receptor types to enter different cell lines or even to infect the same cell type (210). Several viruses belonging to different families may use the same molecule as their cellular receptor. For example integrin avp3 serves as receptor for adenovirus and coxsackie virus A9, whereas poliovirus receptor (PVR) is used by poliovirus and two herpesviruses (Table 1.4). Some of the factors that influence the choice of the receptor are: 1) close proximity of the receptor molecule to the viral attachment site (210); 2) nature, availability and abundance of the receptor molecule on cell surface (102), and 3) ability of the receptor molecule to trigger further events that prime the virus for the subsequent steps in infection process (14, 210). In several viruses, it was reported that a single amino acid substitution in the VAP or the capsid often results in the change of receptor recognition and cellular tropism (14). For example, a substitution of Leu226 to Gin in the receptor binding pocket of haemagglutinin (HA) glycoprotein of influenza virus changes its receptor specificity from NeuAc o2-6 Gal to NeuAc oQ-3 Gal (205). Based on the above observations it has been suggested that the structure and function of the virus  37  Table 1.4.  Viral protein receptors and co-receptors'  Family  Virus''  Retroviridae  HIV  Receptor  GALV/FeLV/SSAV MLV-E MLV-A ALV-A BLV  CD4, C X C R 4 , CCR5, CCR3, CCR2B, CCR8, STRL-33, T Y M S T R , BOB/GPR15 CCR5, STRL-33, T Y M S T R , GPR1,CD4 PiT-1 MCAT-1 PiT-2 TVA BLVR  Poliovirus Coxasckie-B Coxasckie-A9 Major rhinoviruses Minor rhinoviruses Echoviruses  PVR CAR avp3 ICAM-1 L D L R , a2MR, LRP a2pi , CD55  Herpesviridae  BHV-1 HSV-1, HSV-2 HHV-7 HSV-1 EBV  PVR, P r r l . H v e C Prr2, HveB, HveC, Prrl CD4 HveA CR2  Reoviridae  Reovirus  JAM  Adenoviridae  Ad-2, Ad-5 Adenovirus  CAR  Coronaviridae  MHV-A59 Coronavirus-229E  MHVR Aminopeptidase-N  Togaviridae  Sindbis  Laminin receptor  SIV  Picomaviridae  4  4  avP3, avP5 4  4  This table was adapted and modified from reference # 210.  Adenovirus; A L V , Avian leukemia virus; B H V , Bovine herpes virus; Bovine leukemia virus; E B V , Epstein Barr virus; F e L V , Feline leukemia virus; G A L V , Gibbon ape leukemia virus; HIV, Human immunideficienvy virus; HHV, Human herpes virus; HSV, Human herpes virus; H S V , Herpes simplex virus; M L V - A , Amphotropic murine leukemia virus; M L V - E , Ectopic murine leukemia virus; M H V , Mouse hepatitis virus; SIV, Simian immunodeficiency virus; S S A V , Simian sarcoma associated virus; Abbreviations of virus names: A d , BLV,  Abbreviations of receptors and coreceptors: B L V R , bovine leukemia virus receptor; C A R , coxasckie adenovirus receptor; C C R , CC-chemokine receptor; CR, complement receptor; C X C R , CXC-chemokine receptor; GPR, G-protein coupled receptor; Hve, herpesvirus entry protein; I C A M , intracellular adhesion molecule; J A M , junction adhesion molecule; L D L R , low density lipoprotein receptor; LRP, lipoprotein receptor related protein; a2MR, a2-macroglobulin receptor; M C A T , murine cationic amino acid transporter; M H V R , mouse herpesvirus receptor; PiT, inorganic phosphate transporter; Prr, poliovirus receptor related; PVR, poliovirus receptor; T Y M S T R , T-Lymphocyte-expressed seven-transmembrane domain receptor; 4  lntegrins  38  Table 1.5. Viral carbohydrate receptors  1  Receptor  Family  Virus  Coronaviridae  Human coronavirus Bovine coronavirus  Sialic acid-containing oligosaccharides Sialic acid-containing oligosaccharides  Herpesviridae  HSV, Human CMV  Heparan sulfate  Orthomyxoviridae  Influenza A Influenza C  Sialic acid-containing oligosaccharides 9-O-acetylsialic acid  Paramyxoviridae  Sendai  Sialic acid-containing oligosaccharides  Parvoviridae  Canine parvovirus  Sialic acid-containing oligosaccharides  Papovaviridae  Murine polyomavirus  Sialic acid-containing oligosaccharides  Reoviridae  Reovirus-3  Sialic acid-containing oligosaccharides  Retroviridae  HIV  Heparan sulphate  'This table  2  was adapted and modified from reference  Abbreviations of virus names: HIV, Human virus; CMV,  #210  immunodeficiency virus;  Cytomegalovirus.  39  HSV, Herpes simplex  receptor is not a major determinant in the evolution of virus structure within a family. Instead, viruses have evolved under different selection pressures to recognize different receptors, while maintaining the similarities in their genomes and keeping the key threedimensional structural motifs in their capsids and receptor attachment proteins (47, 48).  1.3.2 Secondary receptors In addition to primary receptors, several viruses use other cell surface molecules as secondary receptors. Secondary receptors can be divided into two types: 1) Initial attachment receptors and 2) co-receptors. Usually virus affinity to the secondary receptor(s) is low compared to the primary receptor (69, 79). Initial attachment receptors for several animal viruses have been reported. In many cases, heparan sulphate, an extracellular matrix associated proteoglycan, was identified as initial attachment receptor. It has been suggested that heparan sulphate recruits large amounts of virus to the cell surface, allowing virus to find adhesion strengthening, high affinity receptors that facilitate virus entry into the cell (79, 170). Co-receptors are often involved in post primary receptor binding. Co-receptors for several viruses have been identified. These include integrins for adenoviruses, chemokine receptors (CC and CXC variety) for HIV, SIV and poxviruses and fibroblast growth factor receptor for adeno-associated virus-2 (Table 1.4; 170). In HIV, co-receptor binding is tightly coupled with primary receptor binding. The primary receptor CD4 induces conformational changes in the virion glycoprotein gpl20, so that the chemokine receptor binding site is exposed. Binding to this chemokine receptor induces further  40  conformational changes which facilitates virus-cell membrane fusion by exposing the fusion peptide (51).  1.3.3 Conformational changes involved in virus particles during cell entry After binding to receptors, viruses adopt at least four major mechanisms to enter cells: 1) pH independent fusion at the cell surface, 2) pH dependent fusion in acidic endosomes, 3) receptor-mediated endocytosis and 4) receptor-mediated conformational changes in the virion at the cell surface and subsequent genome delivery (170). In all these entry mechanisms, the virus capsid or the VAP needs to undergo conformational change that allows the exposure of internally located or hidden regions ofthe VAP or the capsid. Conformationally restructured virions are always hydrophobic and proteolytically sensitive. In enveloped viruses, the hidden hydrophobic fusion peptide becomes exposed and lodged into the cellular membrane causing viral and cellular membrane fusion. In non-enveloped viruses, internally located N-terminal hydrophobic sequence becomes externalized leading to pore formation or destabilization of the cellular membrane (51, 48, 82, 107). Due to their hydrophobic nature, the fusion peptides of enveloped viruses and the N-terminal CP segments of nonenveloped viruses are always protected and hidden in unbound virus, so a signal is needed to expose these hydrophobic regions at an appropriate time. The trigger that causes conformational changes is either a receptor or changes in pH or ionic environment (140). For receptor induced conformational changes, a higher affinity interaction is needed between the virus and the receptor (48).  41  1.3.3.1 Enveloped viruses Enveloped viruses bind to cellular receptors with their surface glycoproteins, which subsequently results in viral and cellular membrane fusion. The membrane fusion could either occur with plasma membrane at neutral pH (HIV, measles and herpes simplex viruses), or with endosomal membrane in clathrin coated pits at low pH (influenza, dengue and vesicular stomatitis viruses) (51, 130). Viral membrane fusion mechanisms are well studied in influenza and HIV. Although, these two viruses belong to two entirely different families with different physical morphologies, genome organizations and replication strategies, they share a lot of common features with regard to membrane fusion and cell entry mechanisms (51). Fusion proteins of both these viruses are trimers and synthesized as single chain precursors. Later, these are cleaved by host proteases making them metastable. These metastable fusion proteins are separated from the stable form by an energy barrier. Upon receiving the activation signal, pre-fusogenic fusion proteins undergo extensive conformational changes releasing free energy, which is used for the fusion of viral and cellular membranes (40).  The influenza virus haemagglutinin: The influenza virus envelope glycoprotein, haemagglutinin (HA), is synthesized as HAO in infected cells and it is proteolytically cleaved to HA1 and HA2 by host proteases. HA1 and HA2 are connected by disulfide bonds. Native HA is a 135A long trimer consisting of a globular head, made of HA1 and a stem region made of a part of HA1 and all of HA2. The globular head of HA is a distorted jellyroll structure containing a receptor (sialic acid) binding pocket at the tip (205). The stalk contains a trimeric coiled coil structure in which the amino terminal  42  fusion peptide is buried inside. The approximate distance between the fusion peptide in native HA and the host cell membrane is 100A. Large conformational changes are needed in order for the fusion peptide to reach the host cell membrane. As discussed earlier, low pH is the trigger needed to convert HAfroma native state to a fusogenic state. Low pH activated HA is proteolytically sensitive and hydrophobic in nature. Residues 55 to 75, which are maintained as an extended loop in native HA, are converted in a helical structure. Due to this conformational rearrangement, the central trimeric-coiled coil becomes extended, allowing the buried fusion peptide to reach the host cell membrane (51, 175)..  HIV envelope glycoprotein: The envelope glycoprotein of HIV (gpl60) is synthesized in the ER and cleaved by host proteases in the Golgi apparatus to yield gpl20 (surface glycoprotein, SU) and gp41 (transmembrane glycoprotein, TM). The gpl20 and gp41 are connected by noncovalent interactions. The gpl20 consists of an inner domain that interacts with gp41 and an outer domain, which is exposed on the surface of the trimer. These two domains are connected by a /3-sheet called the bridging sheet. The gp41 consists of an ectodomain and a membrane spanning domain. The ectodomain consists of a trimeric coiled coil structure in which the N-terminus fusion peptide is hidden (209). Upon binding to CD4, gpl20 undergoes major conformational changes involving the inner and outer domain shift. Due to this change the CD4/gpl20 unit bends in such a way that it exposes the chemokine (coreceptor) receptor-binding site. Several lines of evidence show the refolding of up to 100 aa residues in gpl20 upon CD4 binding (209, 40). The chemokine receptor binding induces further conformational changes in the  43  gp41ectodomain, resulting in the insertion of the fusion peptide into the host cell membrane. The trimeric coiled coil structure is a common motif found in several other viral fusion proteins and cellular vesicle fusion proteins (51).  1.3.3.2 Nonenveloped viruses The host cell entry of nonenveloped viruses does not involve fusion, since they lack a lipid bilayer. Instead, multiple copies of one or more capsid proteins are involved in causing a pore or disruption through which viral nucleic acid or nucleoprotein complex enters the host cell cytoplasm (41, 140). Most of the nonenveloped viruses enter the host cell via receptor-mediated endocytosis (130). The activation signals needed for virus uncoating at the endosomal membrane are not well characterized in nonenveloped viruses except in picornaviruses, where the virion disassembly is triggered upon receptor interaction (82). Virus entry mechanisms of nonenveloped viruses are well-studied in picorna (polio- and rhinoviruses) and reoviruses. Although these viruses are distinct from each other in terms of virus architecture, genome organizations and replication strategies, they share many common features with regard to entry mechanisms (41, 107).  Picornavirus entry: Picornaviruses are icosahedral particles 30 nm in diameter, composed of 60 protomers. Each protomer is made of four polypeptides, VP1, VP2, VP3 and VP4. The particle surface is made up of VP1, VP2 and VP3, all of which contain approximately 240 to 290 aa residues. VP4 is a shorter polypeptide (~70 aa) lying across  44  the inner surface of the capsid. VP2 and VP4 are cleaved products of VPO. The VP4 functions as a separated N-terminal extension of VP2, and contains a covalently linked N-terminal myristic acid group, which is predicted to play an important role in the penetration of the cellular membrane (82, 144). It is known that receptor binding sites of several picornaviruses are present in a depression at the base ofthe particle icosahedral 5fold axis known as the canyon (166). The poliovirus particle (160S), upon binding to its cellular receptor (PVR) undergoes conformational changes to convert it into a 135S "A" particle. This 135S particle is hydrophobic and proteolytically sensitive compared to the 160S native virion (59). It has been suggested that, upon receptor binding, the virus particle undergoes expansion in which the VP1, VP2 and VP3 at the particle 3-fold axis move outwardly, creating significant gaps between subunits at the base ofthe canyon. This movement of subunits is similar to movement of subunits during swelling of several isometric plant viruses (76, 82). During this structural transition, the N-terminal ends of VP1 and VP4 come out through the openings at the base of icosahedral five-fold axis to form an amphipathic helical bundle. Insertion of this helical bundle causes a pore, or disruption in the host cell membrane, through which viral RNA enters the cell (82, 178).  Reovirus entry: Reovirus virions are large nonenveloped icosahedral particles 85 nm in diameter. The virion architecture is complex, containing a T=l viral core harbouring a 10 segment dsRNA genome. The viral core surface is formed by a major core protein XI, a second core protein o"2 and a third core protein X2. The X2 projects from the surface at each five fold axis in the form a turret through which viral mRNA  45  passes into the host cell. A trimeric viral attachment protein c l is associated with each turret (107, 41, 150). The viral attachment protein consists of an elongated tail domain and a globular head domain. The tail domain binds to a carbohydrate receptor, sialic acid, and the globular head binds to a protein receptor, junction adhesion molecule (JAM) (57) The viral core is layered by a membrane penetration protein /xl and its protector protein o"3. There are 200 heterohexameric complexes of these two proteins in the form of T=13 cover over the viral core (41, 107). Reovirus undergoes a series of structural transitions during the host cell invasion: 1) shedding of o~3 due to proteolytic cleavage, converting the native virion into Inter Sub Virion Particles (ISVPs); 2) an autolytic cleavage at the N-terminal myristoylated ends of penetration protein ill trimer. These two events convert /il trimer into a proteolytically sensitive, hydrophobic, metastable structure, which is ready to penetrate the host cell membrane. Upon receiving the activation signal in the endosome, helical domains of til trimer separate from each other at the particle three fold axis, inserting N-terminal myristilated ends of ill. The insertion of til in to the endosomal membrane changes X2 turret conformation, releasing a 1 attachment protein from the particle and viral RNA is released into the cell cytoplasm (41, 107). The activation signal that is triggering these conformational changes has not yet been determined. It has been suggested that an interaction with the lipid head group or the conditions in the lumen of the endosome could act as possible triggers (107)  1.4 Cucumber necrosis virus Cucumber necrosis virus was first identified in 1952 on greenhouse cucumber plants in southern Ontario, Canada (122). Infected plants showed a marked malformation  46  of leaves and systemically infected leaves exhibited chlorosis and necrosis. The number of fruits producedfroman infected plant is greatly reduced andfruitsshowed an occasional conspicuous green mottling. Under experimental conditions, only 10 to 20% of CNV infected plants showed systemic infection (122). Although the natural host range of CNV is very narrow, its experimental host range includes several dicotyledonous plants. Initially, CNV was thought to be a member ofthe Necrosis virus genus, but later Rochon and Tremaine (156) reported that it belongs to the tombusvirus group. CNV is a spherical virus 30 nm in diameter that contains a monopartite single-stranded, positive sense RNA genome comprised of approximately 4,700 nt. CNV is serologically distinct from other tombusviruses probably due to the variable nature of the protruding domains of tombusvirus coat proteins (156). CNV is the only definitive member of the tombusvirus genus that is transmitted by a soil-borne fungus, but several other viruses that belong to the family Tombusviridae are transmitted by fungi (Table 1.2)  1.4.1 Genomic organization The entire genome of CNV has been cloned and sequenced (157). The genome contains five long open readingframes(ORFs) encoding 33, 92, 41, 21 and 20 kDa proteins (Figure 1.3). The 92 kDa protein is produced via translational readthrough of the UAG stop codon of ORF1, which produces a 33 kDa protein. Two 3' co-terminal sub genomic RNAs (sgRNAl and sgRNA2) of sizes 2.1 and 0.9 kb are producedfromCNV RNA during infection. SgRNAl serves as a template for translation ofthe 41 kDa protein from ORF3. sgRNA2 is a bifunctional mRNA encoding 21 and 20 kDa proteins from  47  ORF 5  UAG  H ORF1  ORF2  41 k D a  ORF  3  120 k D a l \-m k DJ — ORF 4 sgRNA 1 sgRNA 2  Figure 1.3. Genomic organization of CNV. The five open reading frames (ORFs) present on the CNV genome and the sizes of their corresponding proteins are indicated. The 33 and 92 kDa proteins are produced from genomic length RNA. Subgenomic RNAs 1 and 2 serve as templates for the 41 and 21/20 kDa proteins, respectively. SubgenomicRNA 2 is bicistronic wherein expression of the 20 kDa protein occurs via a leaky scanning mechanism (91).  48  distinct overlapping open reading frames (ORFs 4 and 5 respectively) (159). Expression of the ORF for the 20 kDa protein occurs via a leaky scanning mechanism (91).  1.4.2 Functions of CNV encoded proteins. p33. p33 is proposed to be involved in viral replication based upon the data available on ORF1 products of tombusviruses (162). In several other tombusviruses, p33 along with p92 have been proposed to be a part of the replication complex and are known to be associated with membranes (172). The level of accumulation of p33 is 20 fold more than that of p92 in TBSV-infected plants and protoplasts (172). The p33 protein encoded by Cymbidium ring spot virus (CymRSV) has been shown to localize to peroxisomal membranes and a 7 kDa segment of p33 has been suggested to contain the peroxisome membrane targeting signal (126). p92. p92 is produced as a result of translational readthrough of the p33 stop codon. Initial amino acid sequence comparisons by Rochon and Tremaine (157), showed that CNV p92 contains the glycine-aspartate-aspartate (GDD) motif, which is characteristic of the RNA dependent RNA polymerases (RdRp) of several positivestranded RNA viruses (94, 157). Recently Nagy and Pogany (124) have isolated and characterized the CNV RdRp and showed that CNV p92 does indeed contribute to viral replicase activity. In addition, an arginine/ proline rich sequence termed the RPR motif was found to be important for viral RNA replication (129). p41. Amino acid sequence comparisons and immunoprecipitation studies have shown that p41 is the coat protein (CP) (89, 157). A coat protein mutant (PD-) which lacks the sequence coding for the CP protruding domain (PD) was found to replicate well  49  and move systemically on mechanically inoculated N. benthamiana plants. Neither CNV virions nor CP subunits were found in PD (-) infected plant tissues due to further deletion of the CNV CP ORF (120). Frameshift and deletion mutations in the protruding domain were also proven to be deleterious to particle formation (173). These studies demonstrate that the CNV P domain is important for virus assembly probably by promoting CP dimer formation, which is an important initial step in the assembly of several T=3 icosahedral viruses (77, 142, 177, 179). CNV CP was also shown to be important for fungus transmission (for details see Section 1.2.4.3.1). For details on the CNV CP subunit and particle structure, see Section 1.5.4. p20 andp21. A CNV mutant, which does not express the p20 gene showed highly attenuated symptoms on mechanically inoculated plants even though levels of RNA replication in N. clevelandii indicating that p20 has a major influence on symptom induction (159). Recently it has been shown that the equivalent protein of other tombusviruses (pi 9) acts as a suppressor of gene silencing (173). More recently, CNV p20 has also been shown to act as a suppressor of gene silencing (Rochon and Yu Xiang, personal communication). CNV p21 is a product of ORF4 and is essential in cell-to-cell movement (90). The manner in which p21 facilitates viral movement has not yet been elucidated.  1.4.2 Defective interfering RNAs (Dl RNAs) in CNV Defective interfering (Dl) RNAs are frequently found in tombusvirus-infected plants (146, 162). Dl RNAs are naturally occurring deletion mutants of the viral genome which are unable to replicate in the absence of viral genome. It is believed that Dl RNAs  50  are produced as a result of errors made during replication of the viral genome. Dl RNAs obtained during CNV infections have been isolated and characterized. CNV Dl RNAs retained 5'untranslated and 3' terminal regions along with small portion of the ORF for the 92 kDa replicase protein (55). A CNV mutant that does not express p20 has been shown to generate Dl RNAs rapidly (158). Moreover, CNV coat protein is not required for the efficient generation of Dl RNAs (161).  1.4.4 Structural aspects of CNV The CNV coat protein subunit consists of 380 aa. Putative structures of the CNV coat protein subunit, asymmetric unit and particle have been obtained by homology modelling (see Chapter 2; 93a) based on the crystal structure of the closely related Tomato bushy stunt virus, TBSV (74). The CNV subunit folds into three distinct domains termed the RNA binding (R; 58 aa), shell (S; 167 aa) and protruding (P; 116 aa) domains. The R and S domains are connected by a 34 aa arm, and the S and P domains are connected by a 5 aa hinge (Figure 1.4). The R and arm domains are located internal to the particle; the S domain forms the shell of the particle and the P domain projects outward from the shell. The P domain interacts in pairs across the two-fold axis to form protrusions on the surface (Figure 1.7). The CNV particle, by analogy to TBSV and several other plant viral capsids is arranged in the form of T=3 icosahedral symmetry. The 180 CP subunits that comprise the T=3 structure are chemically identical, but in order to satisfy quasi-equivalence requirements, subunits adopt slightly different three-dimensional structures. These conformationally  51  Figure 1.4. Linear and three dimensional structure o f the C N V C P subunit (C-type). (A) The different C P structural domains are depicted in different colors and are designated as follows: R N A binding (R, white); arm (a, yellow); shell (S, blue); hinge (h, red); and protruding (P, gray). The number o f amino acids in each domain are indicated below individual domains. A l s o shown are flanking sequences in the C N V genome. (B) Surface representation of the homology modelled C N V subunit (C-type). The different structural domains are represented in the same colors as in (A). The disordered R domain is not shown.  52  Figure 1.5. Ribbon diagrams of homology modelled C N V CP subunits and their location on the particle icosahedral axes. I) A subunit (red), II) B Subunit (blue) and III) the ordered C subunit (green). IV) Subunits A , B and C constitute the icosahedral asymmetric unit (Q3). Pentamers of A subunit are located on the particle icosahedral five-fold axis (5) and B and C subunits constitute the hexamers at the icosahedral threefold axis (3). The C/C subunits interact at the icosahedral two fold axis (2). The oligomer was generated using the oligomer generator at: http://mmtsb.scripps.edu/viper/util.php (149). Images of the C N V CP subunit and oligomer were manipulated using WebLab Viewer Lite software (Molecular Simulations, Inc.).  5 3  Figure 1.6. Surface representation of the CNV P-annulus at the particle 3-fold axis viewed from the interior of the capsid. The B and C-type subunits are coloured in gray and white respectively. The extended arms of C-type subunits are depicted in red, blue and yellow for the purpose of clarity.  54  Figure 1.7. Structure of the CNV particle. A, B and C-type subunits are represented in red, blue and green respectively. The different icosahedral axes are labelled as 5 (five-fold), 3 (three-fold), 2 (two-fold) and Q3 (quasi three-fold). (A) Diagrammatic representation ofthe CNV particle. The cutaway section shows the region that the disordered R domain may occupy in the particle interior. This diagram was adapted from reference # 74. (B) Surface representation of homology modeled CNV particle. The oligomer was generated using the oligomer generator at:http://mmtsb.scripps.edu/viper/util.php (149). The image of the CNV particle was manipulated using Web Lab Viewer Lite software (Molecular Simulations, Inc.)  55  by Harrison et al (74). The A and B subunits are identical but the C subunit acquires a slightly different hinge conformation so that the S and P domains are quite differently oriented. In addition, the arm region is ordered in the C subunit but is disordered in the A and B subunits. The ordered 34 aa C subunit arm is divided into an 18 aa '/3' and 16 aa 'e' regions. The ordered arms make a U-turn, fold along the inner edge of the shell domain, and extend towards the particle 3-fold axis. The B regions of the three-fold symmetry related C subunits inter-digitate to form a 54 aa annular structure called the 8annulus. The /3-annular network of all C subunits forms a scaffold inside the particle, which is believed to give stability to the particle (Figure 1.6). In the CNV particle there are 12 icosahedral five-fold axes (a total of 60 copies of the CP subunit) consisting of pentamers formed from A subunits. The B and C subunits interact at the icosahedral three-fold axis to form 20 hexamers (120 copies). As in other T=3 viruses, the icosahedral asymmetric unit consists of A, B and C subunits, which interact at the particle quasi three-fold axis (Figure 1.5 and 1.7).  1.5 Role of N-terminal arm in T=3 icosahedral plant viruses In T=3 icosahedral viruses, the arrangement of CP subunits follows quasiequivalent interactions as suggested by Casper and Klug (36). To achieve this quasiequivalence, subunits in virus shells adopt slightly different conformations. These different conformations are often obtained with the help of segments of the CP polypeptide acting as molecular switches (78, 85). In several small plant and insect RNA viruses these molecular switches are N-terminal segments that are ordered in only one or  56  two ofthe three independent subunits. In some animal viruses, like polyomavirus, the Cterminal segment acts as a molecular switch (180). In the case of Flock house virus (FHV), an insect virus, the N-terminal 20 to 30 aa of the C subunit along with the RNA acts as molecular switch (56, 85). In several plant viruses the N-terminal segments contain basic aa residues which are shown to be important in RNA binding and assembly (44, 109, 167, 143). Deletion of this N-terminal ordered arm in many T=3 plant viruses has resulted in T=l particles devoid of RNA (52, 109, 169). Crystal structures of several T=3 plant viruses have been determined (1, 35, 74, 97, 98, 110, 127, 177, 179). In several of these viruses, it is evident that a part of the internally located N-terminal region of the coat protein subunit is ordered only in C, or B and C subunits. A part of this ordered N-terminal segment is involved in an annular network of/3-strands at the particle three-fold axis. The interaction of ordered N-terminal segments at the particle three-fold axis is one of the common structural themes found in T=3 plant viruses (86). In the case of many tombus-, sobemo- and necroviruses only the C subunit arms are ordered and they interact at the particle three-fold axis to form the /3annulus. In tymo- and bromo viruses, N-terminal segments of both the B and C subunits are ordered and interact with each other at the particle three-fold axis to form an annular structure called the j3-hexamer. It is believed that assembly of several T=3 icosahedral viruses starts from dimers of the coat protein subunit (77, 142, 177, 179). An individual dimer can exist in two conformational states: one where the arms are ordered and dimer structure is flat and the other where the arms are disordered and the dimer structure is curved. In the particle the former structure is designated as the C/C dimer and the latter is called the A/B dimer.  57  Harrison and his co-workers (77, 177) have proposed that assembly of TBSV and TCV initiates by the formation of the /3-annulusfroma trimer of the C/C subunit dimers. Further assembly occurs by the addition of the dimers (either C/C or A/B) with no further distinct intermediate structures. As dimers are being added onto the initiation structure, the N-terminal arm of the C subunit will predetermine the structural state of the incoming dimer (77). Thus the local rules theory (17) applies here wherein the conformation of the incoming protein unit is determined by the structure of the protein to which it attaches. Similarly, in the case of bromo-and tymoviruses, it has been proposed that the /3-hexamer may be the initiation structure during assembly (35, 87, 179). Recent work on Flock house virus (FHV), Cowpea chlorotic mottle virus (CCMV) and Physalis mottle virus  (PhMV) has shown that the ordered N-terminal segment or /3-annular structure is not necessary for virus assembly (49, 168, 206). These studies suggest that virus assembly could be initiated from the 5-fold axis by a pentamer of the A/B dimers (211). Although it seems that the /3-annular structure and order/disorder mechanism is not required for correct assembly, the evolutionary conservation of these structures in several T=3 plant viruses suggests that they might still be important in making the assembly process more efficient and stabilizing the assembled virion (106).  1.6 Virus particle dynamics in T=3 icosahedral plant viruses Both animal and plant viruses are highly flexible in solution and have a propensity to undergo conformational changes under physiological conditions (88, 207). In the case of several animal viruses it has been shown that this ability to undergo  58  conformational changes plays an important role in the process of infection (82, 107, 175). Recently we have shown that conformational changes that are occurring in CNV upon binding to Olpidium zoospores play a key role in its vector-mediated infection (93). In many viruses it has been demonstrated that internally located CP polypeptide segments are occasionally exposed to the outer surface, indicating the dynamic nature of viruses in solution (23, 88, 104). Several plant virus capsids contain divalent cations, which play an important role in maintaining the structural integrity of the virus. X-ray crystal structures of several plant viruses have revealed the presence of calcium or magnesium ions between the subunits (105, 110, 127). In TBSV, there are two calcium ions per subunit which link neighbouring subunits in the CP asymmetric unit (74). These calcium ions interact with carboxylate groups from the side chains of glutamate and aspartate residues. It has been suggested that an important function of these ions is to provide a means of disassembly during the initial stages ofthe infection process (105). Many small spherical plant viruses undergo a reversible change in their structural conformation in the presence of metal chelating agents at elevated pH (207). This process is often referred to as swelling. During this process, the metal ions are chelated and due to the repulsion between negatively charged aspartate residues, the particle undergoes expansion. Incardona and Kaesberg (84) first observed this dynamic behaviour in BMV. Later Bancroft and coworkers studied this phenomenon extensively in CCMV (11) The crystal structure ofthe swollen form of TBSV (152) has revealed that during expansion the particle opens at the quasi three-fold axis between A, B and C subunits (where calcium ions are located). There are 60 openings appearing in the particle and each opening is about 18 to 20 A in  59  diameter. The hexamers along with the /3-annuli and the pentamers essentially remain structurally unchanged due to their extensive interactions with other CP subunits thus preventing complete dissociation of the swollen particle (88). In addition, contacts between A/B subunits at the particle quasi two-fold axis are strengthened due to the formation of a new B- strand resulting from a six residue ordering in A and B subunits (152). As a result, in the expanded structure, the icosahedral two-fold axis and quasi twofold axis become more similar. Moreover, the protruding domains of C/C subunits are rearranged due to structural changes occurring elsewhere in the virus. It has also been shown that a part of the internally located R and arm domains of the A and B subunits are extruded through the large openings becoming exposed on the outer surface of the particle and making swollen virus proteolytically sensitive (63, 75, 93). The atomic structure of the expanded form of CCMV has also been determined (179) at 28 A by Xray crystallography and cryo electron microscopy. Although the compact form of TBSV and CCMV differ significantly, the expanded structures share a relatively higher degree of similarity. This indicates common structural transitions occurring during expansion of T=3 plant viruses (88). It is possible that in solution, the unfolded A and B arms may "breath out" of the virion only occasionally (207) but, the addition of EDTA at higher pH favours the equilibrium towards the conformation in which arms are exposed more frequently. This also means that most of the T-3 plant viruses where expansion has been reported are probably programmed to undergo this conformational transition in nature. As mentioned earlier, the swollen conformation of plant viruses has been implicated in virion disassembly during the initial stages of the infection process (5, 27, 28, 152). But, Albert et al. (5) reported that a CCMV mutant, which is deficient in  60  swelling in vitro, is as infectious as the wild type virus indicating that, in this case swelling may not be required for disassembly. We have recently demonstrated that the swollen-like conformation of CNV is important in the vector transmission by zoospores of the fungus O. bornovanus (93).  1.7 Summary of Thesis Objectives The overall goal of this thesis is to characterize the molecular determinants (viral and vector) required for CNV transmission by zoospores of its vector O. bornovanus. The thesis objectives are divided into three parts.  1. To identify specific sequences and regions on the CNV coat protein and capsid that are important for fungus transmission. A. Analysis of naturally occurring CNV transmission-deficient mutants. Serial mechanical passage of plant viruses in the absence of vector often results in mutants which have lost their ability to be transmitted by their vector. This approach was successfully employed by Robbins et al (153) to isolate naturally occurring transmissiondeficient CNV mutants. In one such mutant LL5, it was found that a single aa substitution in the shell domain was responsible for inefficient zoospore binding and transmission. The same approach and methodology was followed to further characterize regions on the CNV capsid that are important for fungus transmission. B. Location of mutated amino acids on homology-modelled CNV subunit and capsid. CNV is structurally closely related to Tomato bushy stunt virus. Three-  61  dimensional structures of the CNV CP subunit, asymmetric unit and particle were obtained by homology-modelling in order to locate the position of mutated amino acids of CNV transmission mutants on the CNV subunit and capsid. Results of these studies are described in Chapter 2. C. Site-directed mutational analysis. Site-specific in vitro mutagenesis of the CNV CP subunit was conducted to identify CP regions that are important for fungus transmission focussing on regions that were previously identified as being important for transmission (see Chapter 2; 92a). In addition, four prolines in the arm domain and two prolines in the hinge were also mutated to determine the importance of the arm and the hinge in CNV transmission. Results of these experiments are described in Chapters 4 and 5.  2. Biochemical characterization of putative receptor(s) for C N V on O. bornovanus zoospores. A. Pre-treatment of zoospores with periodate, trypsin and phospholipase C. Previous studies in our laboratory (154) suggested the presence of specific recognition molecules (receptors) on the surface of O. bornovanus zoospores for CNV. In order to determine the basic biochemical nature of putative receptor(s), zoospores were treated with trypsin, sodium periodate and phospholipase C and used in in vitro binding assays using CNV. B. Virus overlay assays. CNV overlay assays using total zoospore proteins were conducted in order to determine whether any zoospore protein(s) or glycoprotein(s) are involved in CNV attachment.  62  C. Sugar inhibition studies. Previous work in our lab (154) showed that the surface of O. bornovanus zoospores contains mannose-/glucose- and fucose- containing sugars. Sugar inhibition studies were conducted using virus overlay assays and enzymelinked zoospore binding assays to determine the role of different sugars in CNV binding to zoospores. The results of the above experiments were described in Chapter 2.  3. To determine if zoospore-bound CNV is conformation ally different from unbound virus and whether confomational changes are important for vector transmission of CNV. Several animal virus particles undergo conformational changes as apart of their entry into the host cell. Several plant viruses undergo swelling in the presence of EDTA at alkaline pH. Experiments therefore were conducted to assess the possible significance of swelling in CNV attachment to zoospores. Protease digestion experiments using zoospore-bound, swollen and unbound (native) CNV were conducted to determine if CNV undergoes conformational changes upon binding to zoospores and whether conformationally altered zoospore-bound CNV is structurally similar to swollen CNV. Two highly conserved prolines in the arm region of CNV CP that are predicted to be important for particle conformation were mutated to determine if conformational changes in CNV particles are important for fungus transmission. The results of these experiments are described in Chapter 4.  63  1.8 References 1. Abad-Zapatero, C , S. S. Abdel-Meguid, J. E. Johnson, A. G. W. Leslie, I. Rayment, M . G. Rossman, D . 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Tobravirus 2B proteinacts in trans to facilitate transmission by nematodes. Virology 279:478-487. 199. Vellios, E., G. Duncan, D. Brown, and S. Macfarlane. 2002. Immuno-gold localization of tobravirus 2b nematode transmission helper protein associated with virus particles. Virology 300:118-124. 200. Visser, P. B., and J . F. Bol. 1999. Nonsrtuctural proteins of tobacco rattle virus which have a role in nematode-transmission: Expression pattern and interaction with viral coat protein. J. Gen. Virol. 80:3272-3280. 201. Walkey, D. G. A. 1991. Applied Plant Virology, 2  nd  ed. Chapman and Hall, U K .  202. Wang, J. Y., C. Chay, F. E . Gildow, and S. M . Gray. 1995. Readthrogh protein associated with virions of barley yellow dwarf luteovirus and its potential role in regulating the efficiency of aphid transmission. Virology 206:954-962. 203. Ward, E., and M . J. Adams. 1998. Analysis of ribosomal DNS sequences of Polymyxa species and related fungi and the development of genus and species specificprimers. Plant Pathol. 43:872-877.  79  204. Watson, M . A., and F. M . Roberts. 1939. A comparative study of of the transmission ofHyocymus virus 3, potato virus Y and cucumber virus 1 by the vector Myzus persicae (Sulz), M. circumfexus (Buckton), and Macrosiphum gu (Koch). Proc. R. Soc. Lond. Biol. Sci. 127:543-576. 205. Wiley, D. C , and J. J. Skehel. 1987. The structure and function of the haemagglutinin membrane glycoprotein of influenza virus. Annu. Rev. Biochem. 56:365394. 206. Willits, D., H . Zhao, N. Olson, T. S. Baker, A. Zlotnick, J . E . Johnson, T, Douglas, M . J. Young. 2003. Effects of the cowpea chlorotic mottle bromovirus Bhexamer structure on virion assembly. Virology 306:280-288. 207. Witz, J., and F. Brown. 2001 Structural dynamics, an intrinsic property of viral capsids. Arch. Virol. 146:2263-2274. 208. Woolston, C. J., S. N. Covey, J. R. Penswick, and J. W. Davies. 1983. Aphid transmission and a polypeptide are specified by a defined region of cauliflower mosaic virus genome. Gene 23:15-23. 209. Wyatt, R., and J. Sodroski. 1998. The H I V - l envelope glycoproteins: Fusogens, antigens, and immunigens. Science 280:1884-1888. 210. Young, J . A. T. 2001. Virus entry and uncoating. pp. 87-103. In: D . M . Knipe and P.M. Howley, Editors, Fields Virology, Lippincott Williams and Wilkins. 211. Zlotnick, A., R. Aldrich, J . M . Johnson, P. Ceres, and M . J . Young. 2000. Mechanism of capsid assembly for an icosahedral plant virus. Virology. 277:450-456.  80  2 CHAPTER TWO Identification of Specific Cucumber Necrosis Virus Coat Protein Amino Acids Affecting Fungus Transmission and Zoospore Attachment* 2.1 Introduction Efficient transmission of the majority of plant viruses requires distinct invertebrate or fungal vectors. In most cases, transmission has been shown to be a highly specific process in which only certain vectors can transmit certain viruses (for reviews, see references 4, 6, 13, 14, 23, 35). These observations suggest that virus particles as well as vectors contain specific sites that mediate their recognition. The coat protein (CP) of a plant virus has been shown to play an important role in transmission, and particular amino acids within the CP have been shown to be essential for this process (for reviews, see references 4,6, 13, 14, 23, 35). However, for the most part, the exact role of these amino acids in transmission including their potential role in vector attachment, is not known. Recent work with cucumber necrosis virus (CNV) has suggested that attachment of virions to vector zoospores is an important aspect of the transmission process (24).  * A version of this chapter has been published in the Journal of Virology. Kakani, K., J. Y. Sgro, and D. Rochon. 2001. Identification of cucumber necrosis virus coat protein amino acids affecting fungus transmission and zoospore attachment. J. Virol. 75:5576-5583.  81  C N V , a member ofthe genus Tombusvirus, is a 30-nm spherical virus with a monopartite positive-sense R N A genome (25). Transmission of C N V in nature occurs via zoospores of the Chytridiomycete  fungus, Olpidium bornovanus (6, 9, 24). Zoospores and  virus particles are released independently into the soil from the roots of infected plants. Virus is adsorbed onto the plasma membrane of zoospores and then enters into roots upon zoospore encystment. Studies of C N V transmission by O. bornovanus, and Olpidium transmission of several other small spherical plant viruses, have shown that the transmission process is highly specific (1, 6). For example, O. bornovanus transmits C N V but not Tobacco necrosis necrovirus (TNV), and conversely, O. brassicae transmits T N V but not C N V (10, 34). Moreover, different isolates of O. bornovanus transmit different viruses with varying efficiency (7), and different strains of T N V are transmitted with varying efficiency by the same O. brassicae isolate (17, 33, 34). Electron microscopy studies have shown that adsorption of virus to the zoospore plasmalemma is specific and reflects the virus-vector associations observed in nature (34). Together, these studies indicate the existence of a specific recognition mechanism between virus and vector zoospores.  Previous work has shown that the C N V CP contains determinants that specify its interaction with zoospores of O. bornovanus (20, 24). Reciprocal exchanges between the CP gene of C N V and that ofthe nontransmissible cherry strain of Tomato bushy stunt virus (TBSV) in infectious full-length cDNA clones showed that particles obtained from the TBSV genome containing the C N V CP were transmissible but particles from the C N V genome containing the T B S V CP were not. Also, a single amino acid mutation  82  (Glu to Lys) in the C N V CP shell domain results in lowered transmission efficiency of C N V by O. bornovanus. In vitro binding studies demonstrated that this mutant bound zoospores less efficiently than C N V , indicating that specific regions of the C N V coat protein can mediate zoospore adsorption (24). In this study, we have isolated and characterized several distinct naturally occurring C N V transmission mutants. In each mutant, transmission deficiency was found to be due to a single amino acid substitution in the C N V CP. Moreover, each transmission mutant bound zoospores less efficiently than C N V , suggesting that the altered amino acids affect features of the C N V capsid involved in vector attachment.  2.2 Materials and Methods 2.2.1 Isolation of CNV transmission mutants C N V transmission mutants were obtained following serial passage of virus essentially as described previously (24) except that cucumber cotyledons were used as the local lesion host for isolation of individual mutants.  2.2.3 Virus purification A miniprep procedure (24) similar to the procedure described below was employed to partially purify C N V and C N V mutants for use in the initial screenings for transmission mutants. For all other experiments, virus was purified by differential centrifugation as follows. Infected leaves were ground in 2 volumes of 100 m M sodium acetate (pH 5.0) containing 5 mM /3-mercaptoethanol and allowed to stand on ice for 30 to 60 min. The slurry was then filtered with Miracloth (Calbiochem) and centrifuged at 8,000 g in a GSA rotor. The supernatant was adjusted to 8% polyethylene glycol  83  (molecular weight, 8,000; Sigma) and stirred at 4°C for 1 to 2 h. Virus was pelleted by centrifugation at 8,000 g in a GSA rotor, resuspended in 10 m M sodium acetate (pH 5.0), and subjected to high-speed centrifugation (145,000 g for 2.5 h in a Ti 50.2 rotor) at 4°C. Virus pellets were resuspended as before and centrifuged at 20,800 g in an Eppendorf microcentrifuge. The supernatant was collected and passed through a 0.2-pm-pore-size filter. Concentration of virus was determined spectrophotometrically using an extinction coefficient of 4.5 (absorption of 4.5 at 260 nm is equal to one milligram of virus in a millilitre). The concentration of virus purified by the miniprep procedure was determined by electrophoresis of several dilutions of virions through 1% agarose gels buffered in 45 mM Tris-45 m M borate, (pH 8.3) followed by ethidium bromide staining in buffer containing 1 mM E D T A . Dilutions of purified virus with known concentrations was used as standard for determining the mass of mini-prepped virus loaded on the gel  2.2.4 Fungus transmission assay Purified virions were tested for transmission by O. bornovanus zoospores as previously described (5, 7, 20). Virus (1 pg) was incubated with 10 ml of zoospores (10 /ml in 50 mM glycine, pH 7.6). After a 15-min acquisition period, the mixture was 4  poured onto pots containing 12- to 16-day-old cucumber seedlings. Five days later, roots of cucumber seedlings were tested for the presence of virus by double-antibody sandwich (DAS) enzyme-linked immunosorbent assay (ELISA) using polyclonal antisera raised to C N V particles (20). Absorbance readings greater than fivefold over background (i.e., 0.1 at ^ 4 0 5 ) were considered positive. Each transmission experiment included a wild-type (WT) C N V control, a test to determine any background level of C N V transmission in the absence of zoospores and a test for the presence of contaminating virus in zoospore  84  preparations. Transmission in the absence of zoospores was not detectable in any ofthe experiments.  2.2.5 Cloning and sequence analysis of transmission mutants Double-stranded cDNA copies of the CP coding regions of transmission mutants were obtained by reverse transcription-PCR (RT-PCR) (30). The template was total R N A extracted from either infected leaves or purified virus particles. The plus-sense primer (CNV oligonucleotide 81, 5 A A G A G G T T G A A T T C T G T C A G G 3 ' ) corresponded to C N V nucleotides 2148 to 2168 upstream up the C N V CP open readingframe(ORF) and included a unique EcoRl site (underlined). The minus-sense primer (CNV oligonucleotide 7, 5 'TGTTCCCTAGCGTCGC3') corresponded to the complement of C N V nucleotides 3854 to 3869 and lies downstream ofthe CP ORF. Following amplification, the RT-PCR product was digested with EcoRI and Ncol (both enzymes cut at regions flanking the CP ORF) and ligated into similarly digested pK2/M5, a full-length cDNA clone of WT C N V (26). The sequence of the transferred region of each transmission mutant was determined by cycle sequencing using dye-labeled terminators and AmpliTaq D N A polymerase FS (Perkin-Elmer Applied Biosystems). Samples were sequenced using an ABI PRISM 310 Genetic Analyzer (Perkin-Elmer).  The double mutant LL5K8 was prepared by digestion of LLK8 with Bglll and Ncol (which cleave at unique sites surrounding the LLK8 mutation) followed by insertion ofthe gel-purified fragment into 5g/II/A coI-digested LL5 (24). The presence of both the /  LLK8 and LL5 mutations was verified by sequence analysis.  2.2.6 In vitro transcription and inoculation of plants 85  Preparation of T7 polymerase runoff transcripts and inoculation of plants were as described previously (26).  2.2.7 In vitro binding assay The assay used was a modification of the one described by Robbins et al. (24). One hundred micrograms of purified virus was incubated with 5 x 10 O. bornovanus 5  zoospores in 1 ml of 50 mM sodium phosphate buffer (pH 7.6) for 1 h. Following incubation, zoospores were pelleted by centrifugation at 5,000 rpm for 7 min in an Eppendorf microcentrifuge. The pellet was washed with 1.5 ml of binding buffer and then resuspended in sterile water. The zoospore pellet was assayed for the presence of virus by either Western blot or slot blot analysis using C N V monoclonal antibody 57-2 (24) and an enhanced chemiluminescence detection system (Amersham Pharmacia Biotech). The quantity of virus in the pellet was determined by densitometric analysis of exposed film using the ImageQuant program (Molecular Dynamics). The amount of C N V that pelleted in the absence of fungus was subtracted from the amount of C N V that pelleted in the presence of fungus. Antibody 57-2 was confirmed to react equally to WT virus and mutants in slot blot analysis using denatured virus.  2.2.8 Homology modelling The three-dimensional coordinates of the C N V proteins were modeled after the published coordinates of TBSV, a virus with a known similar structure (PDB entry 2TBV) (22). The virus has icosahedral symmetry with three, independent quasiequivalent structural positions, A , B, and C. Each protein was modeled after its cognate structural homolog with the program Modeler (29) on a Silicon Graphics computer  86  (Silicon Graphics Inc., Mountain View, Calif.). Images of the modeled C N V subunit and trimer were manipulated using WebLab ViewerLite software (Molecular Simulations, Inc.). Surface representations were obtained using the "solvent surface" option. The C N V - T B S V alignment was from a multiple alignment using the CPs of several members ofthe Tombusviridae, including artichoke mottled crinkle virus (PIR2:S24926), carnation Italianringspot virus (PIR2:S52718), cucumber leafspot virus (21), cymbidium ringspot virus (PIR1:VCVGCR), melon necrotic spot virus (PIR1:VCVEMN), pelargonium leaf curl virus (PIR1:A48355), pothos latent virus (SP_VI:Q84832), type T B S V PIR2:S07259, and the cherry strain of T B S V (PIR1:VCVGTB). The program Pileup (version 10.1; Genetics Computer Group) (8) was used to create the multiple alignment.  2.3 Results 2.3.1 Isolation of transmission mutants from mechanically passaged CNV C N V was mechanically passaged 12 times through Nicotiana clevelandii, and individual local lesions were isolated following inoculation of cucumber cotyledons. The CP ORFs and flanking regions of six putative transmission mutants (as determined by reduced transmissibility [data not shown]) were amplified by RT-PCR and cloned in place ofthe WT C N V CP ORF in an infectious C N V cDNA clone. The cloned region was then sequenced to determined the presence of mutations. Transcripts of each ofthe clones were inoculated onto plants, and purified virus from infected plants was tested for transmissibility. Of 87 local lesions analyzed, 7 were ultimately found to contain virus with reduced transmission. Results ofthe transmission tests (Table 2.1) show that cloned mutants designated L L K 8 , LLK10, LLK63, LLK82, LLK84, and LLK85 were less  87  Table 2.1.  Transmission and in vitro binding efficiencies of C N V mutants  Virus  Transmission  % Binding^  WT C N V LLKOO LLK8 LLK10 LLK26 LLK63 LLK82 LLK84 LLK85 LL5K8  49/51 (96) 10/10(100) 3/14(21) 4/15 (27) 1/10(10) 3/21 (14) 15/20 (75) 10/20 (50) 19/25 (76) 0/8 (0)  100.0 130.0 ± 8 . 5 68.3 ± 17.5 39.0 ± 2 2 . 3 ND 20.8 ± 17.2 64.3 ± 2 4 . 1 1 53.0 ± 7 . 5 53.0 ± 12.3 21.7 ± 6 . 6  1  Number of pots showing transmission/number of pots tested. Values in parentheses indicate percentages of pots showing transmission. The data represent a compilation of at least three separate experiments for each virus. 1  Percentage of virus bound in an in vitro binding assay relative to the amount of WT C N V binding (average ± standard deviation of three separate experiments for each mutant). ND, not determined.  transmissible than WT C N V (transmission efficiency, 96%). L L K 8 , LLK10, and LLK63 transmitted at lower efficiencies (i.e., 21, 27, and 14%, respectively), whereas LLK82, LLK84, andLLK85 transmitted at higher efficiencies (75, 50, and 76%). Anuncloned mutant (LLK26) also transmitted with reduced efficiency (10%). Sequence analysis of LLK26 showed that it is identical to LLK8 (see below).  88  We wished to examine the infectivity and level of accumulation of each mutant in order to determine whether the reduced transmission efficiency was due to reduced ability of virus to accumulate in plants following transmission. LLK8, LLK10, LLK63, LLK82, LLK84, and LLK85 virions were inoculated onto N. clevelandii, and plants were monitored for symptoms and for R N A and virion accumulation. All mutants produced symptoms typical of WT C N V on N. clevelandii, resulting in large necrotic lesions on inoculated leaves and subsequent systemic necrosis and death of the plants within 10 to 14 days postinoculation (dpi) (data not shown). Agarose gel electrophoresis of total R N A extracts of infected plants at 3 dpi indicated that each mutant accumulated to approximately the same level as WT C N V (Figure 2.1). In addition, in three separate experiments, DAS-ELISAof leaf extracts at 5 dpi indicated that, on average, virions of L L K 8 , LLK10, LLK63, LLK82, and LLK85 accumulated to approximately the same level as WT C N V (data not shown). LLK84 virions accumulated to approximately 50% of the WT C N V level. All mutants were also capable of infecting cucumber and produced equivalent-sized necrotic lesions on inoculated cotyledons (data not shown). In addition, virion accumulation in cucumber was monitored by agarose gel electrophoresis, and all mutants, including LLK84, accumulated to approximately the same level as WT C N V (data not shown). The integrity of virus particles used for transmission tests was also assessed. Virus particles of each of the transmission mutants were analyzed by agarose gel electrophoresis and found to migrate as discrete bands (Figure 2.2). L L K 8 and LLK10 particles comi grated with WT C N V , whereas particles of LLK63 and LLK84 migrated  89  viral genomic R N A  Figure 2.1. Agarose gel electrophoresis of total leaf R N A extracts from plants infected with C N V transmission mutants. N. clevelandii plants were inoculated with equal amounts of WT C N V or the indicated mutant virions, and total R N A was extracted from inoculated leaves 3 dpi. Equal amounts of total R N A were loaded onto a nondenaturing 1% agarose gel. The gel was stained with ethidium bromide.  90  Figure 2.2. Agarose gel electrophoresis of particles of C N V fungus transmission mutants. The indicated viruses (500 ng of each) were electrophoresed through a 1% agarose gel buffered in Tris-borate (pH 8.3). Virions were visualized by ethidium bromide staining in the presence of 1 mM E D T A .  91  slightly slower than WT C N V and those of LLK82 and LLK85 migrated faster. The greater mobility of LLK82 and LLK85 particles is likely due to the higher net negative charge of the mutated CP (Gly to Glu and Asn to Asp, respectively [see below]). The basis for the slightly slower mobility of LLK63 and LLK84 is not known, but possibly these particles have a slightly expanded conformation, as previously suggested for the C N V LL5 transmission mutant (24). The ability of L L K 8 , LLK10, LLK63, LLK82, and LLK85 to accumulate to approximate WT levels in infected plants suggest that factors other than transmissibility do not likely contribute substantially to their reduced transmission frequencies. DAS-ELISA values for LLK84-infected leaves were approximately twofold less than that of WT CNV. As discussed below, it is possible that the lower accumulation of LLK84 may contribute to the lower transmission frequency of this mutant.  2.3.2 Mutations in CNV transmission mutants map to either the CP shell or protruding domain Based on the structure ofthe related T B S V CP, the C N V CP contains three major structural domains: the R domain, which in the capsid faces the interior; the S domain, which forms the shell of the capsid; and the P domain, which projects outward from the capsid. The linear arrangement of these domains on the C N V CP as well as their predicted structures in the particle subunit and capsid are shown in Figure 2.3D. The CP ORFs as well as flanking sequences used in the construction of cloned transmission mutants described above were sequenced to determine the location and nature of the mutations responsible for the reduced fungus transmission (Figure 2.4). In addition to the unique mutations present in each clone, all transmission mutants also contained a T-to-G  92  Eco^l  Ncol  coat protein  p92  p21  R  nt 2824, T to G aa 66, Phe to Cys p92  R  nt 3674, G to T silent  t  a  WTCNV  p20-  _p2J  LLK00  nt 3507, C to T p20 p92  R  nt 2224, C to T silent  *  P  h  S  nt 2897, A to G silent R  p92  a  LLK8  nt 3511,TtoC aa 295, Val to Ala „  i,  p21  LLK10  p21  LLK26  nt 3507, C to T aa 294, Leu to Phe p92  —  >  R  a  S  h  —  P  nt 3639, A to G aa338, Serto Gly p92  1*  R  Ip20 r-Tp2i~  nt 3697, G to A nt 3781, T to C aa357, Gly to Glu non-coding  * '  R  nt 3289, G to T aa221, Gly to Val p92  p92  p92  R  R  a  *  a  * R  a  P nt 2952, A to G nt 3500, T to C aa 109, Asn to Asp silent S  h  S  h  nt 3156, G to A aa 177, Glu to Lys S  P nt 3507, C to T aa 294, Leu to Phe  p92  R  a  S  LLK82  p20 p21  LLK84  p20 p21  LLK85  "p2TT  h  P  p21  h  P  H I p21  nt 3156, G to A aa 177, Glu to Lys  LLK63  LL5K8 LL5  Figure 2.4. Locations of mutations in C N V fungus transmission mutants. The portion of the mutant genome analyzed for mutation is shown. EcoRl and Ncol restriction enzyme sites used for cloning the mutant CP gene and flanking sequences are indicated for W T C N V . R, a, S, h, and P correspond to the different structural domains of the C N V CP (Fig. 3D). p92, p20, and p21 indicate flanking C N V ORFs. The two mutations present in the transmissible LLK00 and in all C N V transmission mutants are shown by asterisks and are described in detail for LLK00. Mutations in LL5 are also shown. LL5 was made by in vitro mutagenesis of our WT C N V infectious clone and does not contain the two substitutions present in the other mutants. Details of mutations including nucleotide position in the C N V genome, nucleotide change, amino acid position in the C N V CP, and amino acid change are given for each mutant. 93  mutation at C N V nucleotide 2824, which results in a Phe-to-Cys change at amino acid position 66 in the CP arm domain, and a silent G-to-T mutation at nucleotide 3674 in the coding region ofthe CP protruding domain (LLKOO [Figure 2.4]). These same mutations were noted in the previously described C N V transmission mutant LL5 (24), and studies ruled out any effect ofthe amino acid substitution in the arm domain mutation in the low transmission efficiency of LL5. In addition, these studies showed that the LL5 shell domain mutation was sufficient to induce the loss of transmissibility. To determine if the arm and protruding domain mutations together affect C N V transmission, particles produced from transcripts of a cDNA clone containing only these two mutations (LLKOO) were tested for transmission. The results (Table 2.1) demonstrated that these mutations do not affect transmission efficiency. Subsequent sequence analysis of two other C N V clones from passaged virus showed that both mutations were present in both clones (data not shown). Therefore, it appears that these two mutations arose spontaneously following mechanical passage of the original full-length C N V cDNA clone and probably represent the predominant form of the WT transmissible virus from which subsequent transmission mutants arose. The following discussion of the transmission mutants is based on mutations unique to these viruses.  Figure 2.4 shows that each transmission mutant (LLK8, LLK10, LLK63, LLK82, LLK84, and LLK85) contains a single amino acid substitution in the CP and that these occur in either the C N V CP shell or protruding domains; no amino acid changes were found in the R and arm domains, which are located in the particle interior. Two of the transmission mutants, LLK85 and LLK84, contain single amino acid substitutions in the shell domain, whereas the remaining transmission mutants contain single changes in the  94  protruding domain (Figure 2.4). As described above, mutants LLK26 and LLK8 contained identical protruding domain mutations.  Additional nucleotide substitutions that do not affect the CP amino acid sequence were found in LLK10, LLK82, and LLK85. InLLKlO, two silent substitutions were found: one in the 3'-terminal region of C N V p92 ORF (the putative RNA-dependent R N A polymerase) (25) and the other in the arm region of the C N V CP ORF. LLK85 contained a silent substitution in the coding region of the C N V CP protruding domain. These mutations were not further investigated since they do not affect the protein sequence and are not present in areas ofthe genome which have known regulatory nucleotide sequences. LLK82 contains a T-to-C change in the core promoter for the subgenomic mRNA2 that encodes proteins involved in cell-to-cell movement (p21) and symptom induction (p20) (16, 26). However, as described above, several analyses of LLK82 accumulation levels failed to indicate that the T-to-C change affects virus accumulation (see above).  2.3.3 CNV transmission mutants show decreased binding to zoospores in vitro We have previously shown that C N V binds to zoospores in vitro and that the C N V , transmission mutant LL5 shows reduced in vitro zoospore binding (24). These data suggested that the LL5 CP lacks an important determinant for attachment to zoospores. We wished to assess the possibility that reduced transmission of LLK8, LLK10, LLK63, LLK82, LLK84, and LLK85 is due to inefficient ability of mutant particles to bind zoospores. One hundred micrograms of each transmission mutant was incubated with 5 x 10 zoospores for 1 h, followed by low-speed centrifugation to pellet zoospores and 5  95  washing to remove unbound or nonspecifically bound virus. The amount of bound virus in the pellet was determined by Western blot or slot blot analysis. Table 2.1 shows that each transmission mutant binds to zoospores less efficiently than WT CNV, with binding efficiencies ranging from approximately 21 to 68% of that of WT CNV. These results suggest that the reduced transmission of C N V mutants is at least partly due to their reduced abilities to attach to zoospores during the transmission process.  2.3.4 An artificial double mutant transmits and binds to zoospores at a lower efficiency than either of the individual mutants An artificial double mutant (LL5K8 [Figure 2.4]) containing the mutations present in both LLK8 and the previously described LL5 mutant (24) was constructed and assessed for transmission. Table 2.1 shows that this mutant is less transmissible (0%) than either LLK8 (21% transmission) or LL5 (20% transmission) (24). Corresponding results were obtained in in vitro binding studies, i.e., LL5K8 binds zoospores less efficiently (22%) than either L L K 8 (68%) (Table 2.1) or LL5 (50%) (24). When the double mutant was tested for its ability to infect and accumulate in N. clevelandii and cucumber, no substantial decrease in the level of R N A accumulation (Figure 2.1) or particle accumulation as determined by ELISA (data not shown) was observed. In addition, particles appeared intact, as determined by agarose gel electrophoresis (Figure 2.2). These results reinforce the role of both the LLK8 and LL5 mutations in the attachment and transmission processes.  2.4 Discussion We have isolated and characterized several naturally occurring C N V mutants deficient in transmission by O. bornovanus. Each mutant contains amino acid  96  substitutions in the CP, reinforcing previous studies on the role of this protein in fungus transmission (20, 24). All of the CP mutations occurred in either the shell or protruding domain. These portions of the CP, unlike the R and arm domains, form the surface ofthe particle, which raises the possibility that the affected amino acids may serve as attachment sites for interaction of C N V with a putative zoospore receptor (see below).  In vitro binding studies showed that each transmission mutant bound to zoospores less efficiently than WT C N V (Table 2.1). These data suggest that zoospore binding plays an important role in transmission of these mutants, although other unidentified viral or host factors likely contribute to the transmission process.  All transmission mutants accumulated in cucumber to approximately the same level as WT C N V , indicating that virus particles are stable and that defects in transmission cannot be attributed to an inability of particles to accumulate in cucumber following transmission. With the exception of LLK84, which accumulated to approximately 50% ofthe WT C N V level, all transmission mutants also accumulated to W T C N V levels in N. benthamiana (Figures 2.1 and 2.2). The basis for the slightly reduced accumulation of LLK84 in this host is not known, but considering the location of the LLK84 mutation in the trimer interface, it is possible that the particles are partly defective in assembly or disassembly. We note that accumulation data were taken from both inoculated and systemic tissue of infected N. benthamiana but only from inoculated leaves of cucumber. It is possible that the lower accumulation levels observed in N. benthamiana are due to decreased ability of LLK84 to move systemically.  97  LLK8 and LLK10 contain mutations corresponding to amino acids that are immediately next to each other in the linear structure of the CP P domain (amino acids 294 and 295, respectively [Figure 2.4]). Amino acids from other mutants did not cluster on the primary CP structure. However, it was of interest to assess whether the other mutations clustered in the secondary or tertiary structure of the subunit or capsid and whether these sites are potentially exposed on the surface. To do this, homology modelling ofthe C N V CP subunit was conducted using the known high-resolution X-ray crystal structure of the related T B S V CP subunit (15). Figures 2.3 A and B show ribbon and surface representations, respectively, of the modeled C N V subunit, and Figure 2.3C shows a surface representation of the modeled C N V CP trimer (the asymmetric unit). The surface representation models predict that with the exception of LLK10, all of the mutated sites (including the previously identified site in LL5) are exposed on the surface of the subunit or trimer. In addition, six of seven ofthe mutated sites (i.e., LLK82, LLK8, LLK84, LL5, and LLK85) are preferentially located on one side of the CP subunit (Figure 2.3B). Mutated amino acids in LLK8, LLK10, and LLK82 are all located on the outer wall of the protruding domain dimer, and those in LLK84 and LL5 are near each other in a region of subunit-subunit interaction in the trimer (Figures 2.3 A and B). The fact that the mutations map to distinct regions on the capsid is compatible with multiple mechanisms for transmission and binding defects. Nevertheless, the modeled C N V CP trimer predicts that most ofthe mutated sites (LL5, LLK8, LLK10, LLK82, and LLK84) are in or near a cavity formed by the trimer on the particle quasi-threefold axis. It is therefore possible that the trimer cavity represents an important site for recognition of a putative zoospore receptor. If these mutations disrupt binding to a receptor, it would  98  Figure 2. 3. Locations of mutated amino acids on the C N V CP subunit and trimer in C N V transmission mutants. (A) Ribbon diagram of the homology modeled C N V CP subunit (subunit C) showing locations of mutated sites (in white in ball-and-stick form) in each of the transmission mutants. The mutated site in L L K 1 0 is shown in red to distinguish it from the adjacent LLK.8 mutation. Locations of the P, S, and a domains are indicated (see panel D for details). The disordered R domain is not shown. (B) Surface representation of the C N V CP subunit (subunit C) showing locations of mutated sites in white. The position of the buried L L K 1 0 mutation is indicated by the white dotted lines. The LLK63 mutation is not visible in this orientation. (C) Surface representation of the C N V trimer (asymmetric unit) showing locations of mutated sites in each transmission mutant. The red, blue, and green areas correspond to the A , B , and C subunits. The asterisk shows the quasi-threefold axis of symmetry (D). (D) Diagrammatic representation of the structure of T B S V used for reference to the C N V structure, (a) Linear order of the different CP domains is shown along with the number of amino acids comprising each C N V domain (R, R N A binding domain; a, arm; S, shell domain; h, hinge; P, protruding domain), (b) Subunit structure with locations of indicated domains, (c) Particle structure with the A subunit in red, B in blue, and C in green. The cutaway section shows the region that the disordered R domain may occupy in the particle interior. (This diagram was adapted from reference 3).  99  suggest that the receptor has complementary symmetry. Alternatively, the affected amino acids in these mutants may affect subunit-subunit interactions and virion conformation, thereby indirectly affecting virion attachment and subsequent transmission. The slower electrophoretic mobility of mutants LL5 (24), LLK63, and LLK84 (and LL5K8) (Figure 2.2) is consistent with the notion that reduced binding and transmission efficiencies may be due to conformational changes in particle structure as a result of the amino acid substitution. As stated above and shown in Figure 2.3C, LLK84 and LL5 mutations lie in a region of subunit contact and could therefore affect subunit interactions. Similarly, the mutation in LLK63 lies in a region of protruding domain dimer interactions and could affect particle conformation by interfering with protruding domain contacts.  The mutation in LLK10 reduces transmission to about 27% of the WT C N V level and decreases binding to 39% as a result of a Val-to-Ala change at amino acid 295 in the CP protruding domain. This substitution lies immediately next to the mutated site in LLK8. The modeled C N V subunit does not predict that the affected LLK10 amino acid is exposed on the particle surface. It is possible that replacement of Val by Ala indirectly affects transmission and binding by changing the accessibility of other exposed amino acids in this region.  The structure of the shell domain of the tombusvirus CP subunit is similar to that of the picornavirus particle (27), which raises the question as to whether the putative zoospore attachment sites on C N V correspond to any of the known cellular receptor attachment sites on picornaviruses. In foot-and-mouth disease virus, an RGD motif in the G-H loop of VP1 has been implicated in receptor attachment (11, 19). Interestingly, the  100  Gly-to-Val mutation in the C N V mutant LLK84 lies within the structurally analogous GH loop and is located within an SGD triplet. Mutagenesis studies may help in the final identification of this region ofthe C N V capsid in zoospore attachment.  Virus attachment sites on animal viruses are for recognition of receptors that lie on host cells infected by the virus. Plant viruses do not recognize receptors for host cell attachment, but certain plant viruses are likely to possess attachment sites for recognition ofthe vector that transmits the virus to its host. In other cases, a virus-encoded helper factor is believed to mediate interaction between the vector and the virus particle (23). Specific virus attachment sites for cellular receptors have been identified for several animal viruses, including poliovirus, foot-and-mouth disease virus, and influenza virus (12, 28, 31). However, such sites have not yet been identified in plant viruses, despite their importance in the establishment and dissemination of plant virus disease. Specific regions ofthe capsid involved in transmission have been identified in several plant viruses (4, 6, 13,14, 23, 32, 35), but to our knowledge no experiments have been conducted to determine if these sites are involved in the vector attachment stage of transmission. In tomato spotted wilt virus, an R G D motif has been identified in one of the viral structural proteins (18) and has been implicated but not proven to be involved in vector attachment. Our studies represent the initial stages of work that aims to identify features of virion architecture required for attachment to a vector. It is hoped that further work will provide information on evolutionarily conserved features of virus particles that are involved in receptor attachment. In addition, virus attachment mutants should aid in the identification of virus vector receptors about which very little is known.  101  2.5 References 1. Adams, M . J . 1991. Transmission of plant viruses by fungi. Ann. Appl. Biol. 118:479492. 2. Blanc, S., E . D. Ammar, S. Garcia-Lampasona, V. V. Dolja, C. Llave, J. Baker, and T. P. Pirone. 1998. Mutations in the potyvirus helper component protein: effects on interactions with virions and aphid stylets. J. Gen. Virol. 79:3119-3122. 3. Branden, C , and J. Tooze. 1991. Introduction to protein structure. Garland Publishing, Inc., New York, N.Y. 4. Brown, D. J. F., W. M . Robertson, and D. L . Trudgill. 1995. Transmission of viruses by plant nematodes. Annu. Rev. Phytopathol. 33:223-249. 5. Campbell, R. N., H . Lecoq, C. Wipf-Scheibel, and S. T. Sim. 1991. Transmission of cucumber leaf spot virus by Olpidium radicale. J. Gen. Virol. 72:3115-3119. 6. Campbell, R. N. 1996. Fungal transmission of plant viruses. Annu. Rev. Phytopathol. 34:87-108. 7. Campbell, R. N., S. T. Sim, and H . Lecoq. 1995. Virus transmission by host-specific strains of Olpidium bornovanus and Olpdium brassicae. Eur. J. Plant Pathol 101:273-282. 8. Devereux, J., P. Haeberli, and O. Smithies. 1984. A comprehensive set of sequence analysis programs for the V A X . Nucleic Acids Res. 12:387-395. 9. Dias, H . F. 1970. Transmission of cucumber necrosis virus by Olpidium cucurbitacaerum Barr & Dias. Virology 40:828-839. 10. Dias, H . F. 1970. The relationship between cucumber necrosis virus and its vector, Olpidium cucurbitacaerum. Virology 42:204-211. 11. Fox, G., N. R. Parry, P. V. Barnett, B. McGinn, D. J. Rowlands, and F. Brown. 1989. The cell attachment site on foot-and-mouth-disease virus includes the amino acid sequence R G D (arginine-glycine-aspartic acid). J. Gen. Virol. 70:625-637.  12. Fry, E . E . , S. M . Lea, T. Jackson, J. W. I. Newman, R. M . Ellard, W. E . Blakemore, R. Abu-Ghazaleh, A. Samuel, A. M . Q. King, and D. I. Stuart. 1999. The structure of a foot-and-mouth disease virus-oligosaccharide receptor complex. E M B O J. 18:543-554. 13. Gray, S. M . 1996. Plant virus protein involved in natural vector transmission Trends Microbiol. 4:259-264.  102  14. Gray, S. M . , and D. M . Rochon. 1999. Vectors of plant viruses, p. 1899-1910. In A . Granoff, and R. Webster (ed.), Encyclopedia of virology, vol. 1. Academic Press, London, England. 15. Harrison, S. C., A . J. Olson, C. E . Schutt, F. K. Winkler, and G. Brigogne. 1978. Tomato bushy stunt virus at 2.9 A resolution. Nature (London) 276:368-373. 16. Johnston, J. C., and D. M . Rochon. 1995. Deletion analysis ofthe promoter for the cucumber necrosis virus 0.9 kb subgenomic RNA. Virology 214:100-109. 17. Kassanis, B., and I. MacFarlane. 1965. Interaction of virus strain, fungus isolate, and host species in the transmission of tobacco necrosis virus. Virology 26:603-612. 18. Kormelink, R., P. de Haan, C. Meurs, D. Peters, and R. Goldbach. 1992. The nucleotide sequence of the M R N A segment of tomato spotted wilt virus, a bunyavirus with two ambisense R N A segments. J. Gen. Virol. 73:2795-2804. 19. Mason, P. W., E . Reider, and B. Baxt. 1994. R G D sequence of foot-and-mouth disease virus is essential for infecting cells via the natural receptor but can be bypassed by an antibody-dependent enhancement pathway. Proc. Natl. Acad. Sci. U S A 91:19321936. 20. McLean, M . A., R. N. Campbell, R. I. Hamilton, and D. M . Rochon. 1994. Involvement ofthe cucumber necrosis virus coat protein in the specificity of fungus transmission by Olpidium bornovanus. Virology 204:840-842. 21. Miller, J. S., H . Damude, M . A . Robbins, R. D. Reade, and D. M . Rochon. 1997. Genome structure of cucumber leaf spot virus: sequence analysis suggests it belongs to a distinct species within the Tombusviridae. Virus Res. 52:51-60. 22. Olson, A . J., G. Bricogne, and S. C. Harrison. 1983. Structure of tomato bushy stunt virus. IV. The virus particle at 2.9 A resolution. J. Mol. Biol. 171:61-93. 23. Pirone, T. P., and S. Blanc. 1996. Helper dependent vector transmission of plant viruses. Annu. Rev. Phytopathol. 34:227-247. 24. Robbins, M . A . , R. D. Reade, and D. M . Rochon. 1997. A cucumber necrosis virus variant deficient in fungal transmissibility contains an altered coat protein shell domain. Virology 234:138-146. 25. Rochon, D. M . , and J. H . Tremaine. 1989. Complete nucleotide sequence of the cucumber necrosis virus genome. Virology 71:251-259.  103  26. Rochon, D. M . , and J. C. Johnston. 1991. Infectious transcripts from cloned cucumber necrosis virus cDNA: evidence for a bifunctional subgenomic mRNA. Virology 181:656-665. 27. Rossmann, M . G. 1987. The evolution of R N A viruses. Bioessays 7:99-103. 28. Rossman, M . G. 1994. Viral cell recognition and entry. Protein Sci. 3:1712-1725. 29. Sali, A., and T. L . Blundell. 1993. Comparative protein modeling by satisfaction of spatial restraints. J. Mol. Biol. 234:779-815. 30. Sambrook, J., E . F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N. Y . 31. Skehel, J. J., and D. C. Wiley. 2000. Receptor binding and membrane fusion in virus entry: the influenza virus hemagglutinin. Annu. Rev. Biochem. 69:531-569. 32. Smith, T. J., E . Chase, T. Schmidt, and K. L . Perry. 2000. The structure of cucumber mosaic virus and comparison to cowpea chlorotic mottle virus. J. Virol. 74:7578-7586. 33. Teakle, D. S., and C. Hiruki. 1964. Vector specificity in Olpidium. Virology 24:539-544. 34Temmink, J . H . M . , R. N. Campbell, and P. R. Smith. 1970. Specificity and site of in vitro acquisition of tobacco necrosiss virus by zoospores of Olpidium brassicae. J. Gen. Virol. 9:201-213. 35. Van den Heuvel, J . F. J. M . , S. A . Hogenhout, and F. van der Wilk. 1999. Recognition and receptors in virus transmission by arthropods. Trends Microbiol. 7:7176.  104  3 CHAPTER THREE Evidence that Binding of Cucumber Necrosis Virus to Vector Zoospores Involves Recognition of Oligosaccharides* 3.1 Introduction Animal viruses gain entry to host cells by using host cell surface molecules as receptors (11, 16, 23, 26). However, plant viruses generally gain entry into plants through specific invertebrate or fungal vectors which breach the otherwise impermeable cell wall. In most cases, transmission of plant viruses has been shown to be a highly specific process in which only certain vectors can transmit certain viruses (4, 9, 19, 31). These observations suggest that vectors contain specific sites that are recognized by virus particles. The coat proteins of several plant viruses have been shown to be important in the transmission process (4, 9, 19, 31). However, there are only few reports of the possible involvement of receptors in or on vectors that mediate transmission. Li et al. (15) have recently identified two proteins from head tissues of an aphid vector as potential receptor candidates for Barley yellow dwarf virus (family Luteoviridae). In addition, Bandla et al., (3) have reported that a 50-kDa midgut protein form Frankliniella occidentalis, the main thrip vector for Tomato spotted wilt virus (genus Tospovirus), is a potential receptor. Symbionin, a homologue of Escherichia coli GroEL chaperonin  * A version of this chapter has been published in the Journal of Virology. Kakani, K., Robbins, M . & Rochon, D. 2003. Evidence that binding of cucumber necrosis virus to vector zoospores involves recognition of oligosaccharides. J. Virol. 77, 3922-3928.  105  protein which is highly abundant in aphid hemolymph (31), has been shown to bind to luteoviruses and to play an important role in the transmission process. Certain other viruses, such as potyviruses, caulimoviruses, and tobraviruses, require additional virusencoded "helper factor" proteins for transmission. In the cases of potyviruses and caulimoviruses, the helper factor likely serves as a bridge between virus particles and attachment sites in the vector (9, 10, 31). Several small spherical viruses within the family Tombusviridae are transmitted by zoospores of the Chytrid fungus Olpidium (either Olpidium bornovanus or Olpidium brassicae) (4). It has been proposed that transmission occurs following the release of zoospores and virus from different plants into the soil and subsequent adsorption of virus particles onto the surfaces of zoospores. Bound virus then enters the cells of plants following the encystment of zoospores on roots (1, 4). Several studies have shown that the transmission process is highly specific (1, 4). For example, O. brassicae transmits the necrovirus Tobacco necrosis virus (TNV-A) but not the tombusvirus Cucumber necrosis virus (CNV), and conversely, O. bornovanus transmits C N V but not T N V - A (7, 30). Moreover, different O. bornovanus isolates transmit either CNV, Melon necrotic spot virus (MNSV), or Cucumber leaf spot virus (CLSV) with various efficiencies (5), and different necrovirus species are transmitted with different efficiencies by the same O. brassicae isolate (13, 29, 30). Electron microscopy studies have shown that adsorption of virus to the zoospore plasmalemma is specific and reflects the virus-vector associations observed in nature (30). Together, these studies indicate the existence of a specific recognition mechanism(s) between virus and vector zoospores.  Previous work has shown that the C N V coat protein contains determinants for the specificity of transmission by O. bornovanus (18, 22). More recently, it has been shown  106  that a cavity at the quasi-threefold axis is important in C N V attachment and transmission (12). In this study, we wished to determine whether the acquisition and subsequent transmission of C N V by O. bornovanus involves specific zoospore receptors.  3.2 Materials and Methods 3.2.1 Virus isolates and purification CNV, C L S V , and T N V - A were maintained by mechanical passage in Nicotiana benthamiana or Nicotiana clevelandii, and M N S V was similarly maintained in Cucumis sativis. The viruses were purified by differential centrifugation as previously described (12).  3.2.2 Maintenance of fungal cultures O. bornovanus (isolate SSI96) was maintained on cucumber roots (C. sativis cv. Poinsette 76), and O. brassicae (isolate SS58)was maintained on lettuce roots (Lactuca sativa cv. White Boston) as described by Campbell et al. (5).  3.2.3 In vitro zoospore binding assays In vitro zoospore binding assays were conducted as previously described (12) using Western blot analysis followed by densitometry for quantification of the amount of bound virus. Monoclonal antibody 57-2 was used for detection of C N V , and the respective polyclonal antibodies were used for all other viruses. The monoclonal antibody was prepared in mice using C N V particles as the immunogen (M. Robbins and D. Rochon, unpublished data). Saturation binding data was analyzed using nonlinear  107  regression analysis and the "one-site binding method" in the GraphPad Prism software package (http://www.graphpad.com).  3.2.4 Trypsin, periodate, and phospholipase C treatment of O.bornovanus zoospores One milliliter of O. bornovanus zoospores (5 X 10 /ml) was incubated in either 10 5  mM sodium periodate-0.1% trypsin (Sigma) or 5 mU of phospholipase C (Sigma) for 15 min. Periodate oxidation was done in deionized water, trypsin digestion was done in 100 m M sodium phosphate buffer (pH 7.6), and phospholipase C treatment was done in 10 m M Tris (pH 7.6)-5 m M CaCf;. Following treatment, the zoospores were pelleted at 2,000 x g for 7 min, resuspended in 1 ml of binding buffer (50 m M sodium phosphate buffer, pH 7.6), and then used in an in vitro binding assay as described above. C N V virions were assessed for resistance to residual trypsin digestion by incubating 100 pg of C N V in 0.002 to 0.2% trypsin for 40 min in binding buffer. The integrity of the virus was assessed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and agarose gel electrophoresis. Degradation of the C N V coat protein subunit was not observed following SDS-PAGE, and the virus particle appeared intact following agarose gel electrophoresis (data not shown).  3.2.5 Virus overlay assays Virus overlay assays were done essentially as described by Salas-Benito and del Angel (24) with some modifications. A total of 2 X 10 zoospores (20 pi) in denaturation 6  buffer (14) were loaded per lane on an SDS-12% P A G E gel (14). The proteins were blotted onto nitrocellulose membranes (0.45-um pore size; Bio-Rad) and then renatured overnight in phosphate-buffered saline (PBS)-4% bovine serum albumin at 4°C. The blots  108  were washed three times for 10 min each time in PBS and then blocked in PBS containing 5% dry milk powder for 1 to 2 h. After being washed as described above, the blots were incubated with 100 ug of virus in 10 ml of 50 mM sodium phosphate buffer (pH 7.6) for 3 h. The blots were washed and then incubated for 1 h with C N V monoclonal antibody 57-2. The antigen-antibody complexes were detected using peroxidase-labeled goat anti-mouse antibody (Jackson Immuno Research Laboratories) and the Enhanced Chemiluminescence Detection System (Amersham Pharmacia Biotech). Monoclonal antibody 57-2 was confirmed to react equally to wild type (WT) C N V and to the C N V transmission mutants in slot blot analysis using undenatured virions.  Sugar inhibition studies were conducted by preincubating C N V virions in 10 ml of binding buffer (pH 7.6) and either 0.1 M D-(+)-mannose, 0.1 M methyl a-D-mannoside, or 1 mg of mannan/ml for 45 min at room temperature prior to the addition of virions to the blots.  3.2.6 Microtiter plate binding assays A modification of a previously described microtiter plate-based binding assay utilizing lectins (17) was used. Polystyrene microtiter plates (96 well; Libro/Titerek) were coated with 100 ml of O. bornovanus zoospores (2.5 x 10 zoospores/ml) in 50 mM 6  potassium phosphate buffer, pH 7.0, and incubated overnight at 37°C. After incubation, the plates were blocked at 37°C for 2 h in PBS containing 5% milk powder and 5% bovine serum albumin. After being washed with distilled water, the plates were incubated for 1 h at 37°C with either 8 ug of C N V in 100 ul of 50 mM sodium phosphate (pH 7.6) binding buffer or 8 ug of C N V preincubated with sugar solution (see below) in binding  109  buffer. The plates were washed as described above, and the amounts of C N V bound were determined using C N V monoclonal antibody 57-2 followed by detection with goat antimouse antibody conjugated to alkaline phosphatase. The relative amounts of bound virus were determined 10 to 40 min following the addition of substrate by measuring the absorbance at 405 nm. Under these conditions, the relationship between bound virus and absorbance at 405 nm was linear, as determined by a dilution series of C N V virions.  The following sugars were tested for the ability to inhibit C N V binding to O. bornovanus zoospores: D-(+)-glucose, D-(+)-galactose, D-(+)-mannose, L-(-)-arabinose, L(-)-fucose, A'-acetyl-D-glucosamine, D-(+)-xylose, L-(-)-sorbose, D-(-)-fructose, methyl O-Dmannoside, yeast mannan, D-mannosamine, «3,a6-mannopentaose, mannotriose-bis[./Vacetyl-D-glucosaminyl], and iV-acetyl-D-mannosarnine. The sugars were initially tested for inhibitory activity using 14 10-fold serial dilutions beginning with 0.2 M sugar. Dilutions of a3,cr6-mannopentaose and mannotriose were as described above, beginning with 1,067 and 538 uM solutions, respectively. Sugars that showed inhibition were then further tested within the inhibitory range using several threefold dilutions. Inhibition in the case of yeast mannan was determined using a starting concentration of 15 mg/ml (equivalent to -0.1 M in terms of the mannose residue concentration) followed by a series of threefold serial dilutions. CNV-sugar solutions were incubated for 45 min at room temperature prior to being added to the zoospores.  3.2.7 Labelling of O. bornovanus zoospores with FITC-labeled lectins Sixteen micrograms of fluorescein isothiocyanate-labeled concanavalin-A (ConAFITC; Sigma) or 30 pg of FITC-labeled Tetragonolobuspupurea  110  agglutinin (TPA-FITC;  Sigma) were incubated for 10 min with washed O. bornovanus zoospores (2 x 10 to 2 x 5  10 ) in 1 ml of binding buffer. The zoospores were viewed with a Zeiss Axiophot 6  epifluorescence microscope using an excitation wavelength of 450 to 490 nm. The specificity ofthe labelling reaction was determined by preincubating the lectin with either 0.5 M D-(+)-mannose (in the case of ConA) or 0.15 M L-(-)-fucose (in the case of TP A) prior to adding it to the zoospores.  3.3 Results 3.3.1 Binding of CNV to zoospores is saturable and specific. Two major criteria for viral recognition sites as receptors are saturability and specificity (28). A n in vitro binding assay (12, 22) was used to determine whether C N V binding to O. bornovanus zoospores is saturable. Increasing amounts of C N V (2.2 to 55 pmol) were incubated with 4 X 10 O. bornovanus zoospores in binding buffer. Following 5  a 1-h incubation, the virus-zoospore suspensions were centrifuged at low speed, the pellets were washed, and the amounts of virus bound to pelleted zoospores were determined using Western blot analysis followed by densitometry. Figure 3.1 A shows that binding of C N V to zoospores is indeed saturable, becoming apparent at ~20 pmol of CNV.  To further assess the possibility that Olpidium zoospores contain specific virus attachment sites, C N V , M N S V , and CLSV—each known to be transmitted by O. bornovanus (but not by O. brassicae)—and TNV-A—known to be transmitted by O. brassicae (but not by O. bornovanus)—-were examined for the ability to bind to zoospores of either O. bornovanus or O. brassicae using the above-described binding assay. Figures  111  O. bornovanus zoospores  E  a.  0 0300 0120 0090 0060 0030 000-  0.120 0.090' 0.030 0.012 0.009 0.006 E 0_ 0.003 0.000  B  _r_  0.15CH  > o  >  > o  0. brassicae zoospores  > z o  >  >  Figure 3.1. Virus-zoospore binding assays. (A) Saturation binding of C N V to O. bornovanus zoospores. Increasing amounts of C N V were added to 4 x 10 zoospores, and the amounts of bound virus were determined using Western blot analysis followed by densitometry. The amount of virus bound to washes of uninfected roots was also determined for each concentration of virus, and this value was subtracted from the amount of virus bound to zoospores. All values represent the average of triplicate samples from one representative experiment. (B and C) Ten picomoles of either C N V , MNSV, CLSV, or T N V - A was incubated with 10 O. bornovanus (B) or O. brassicae (C) zoospores, and the amount of virus bound was determined as for panel A. The results are the averages of triplicate treatments from two separate experiments plus standard deviations. (This figure was derived from work done by former graduate student, Marjorie Robbins.) 5  6  112  3.1 B and C shows that C N V , MNSV, and C L S V each bind O. bornovanus zoospores more efficiently than T N V - A (2.6-, 2.8-, and 17-fold, respectively [Figure 3. IB]) and that T N V - A binds O. brassicae zoospores more efficiently than either C N V , MNSV, or C L S V (3.3- and 30-fold, with no detectable C L S V binding [Figure 3.1C]). These experiments show that specificity indeed occurs in the attachment of these viruses to Olpidium zoospores and, moreover, that the specificity observed in vitro reflects previously described biological specificities (4, 5). These results, in conjunction with the saturation binding experiments, support the possibility that specific zoospore receptors are involved in the acquisition and transmission of these viruses in nature.  3.3.2 Periodate, trypsin, and phospholipase C treatment of O. bornovanus zoospores. To initially characterize the biochemical nature of the molecule(s) on the surface of O. bornovanus zoospores involved in C N V attachment, zoospores were treated with either 10 mM sodium periodate, 0.1% trypsin, or 5 mU of phospholipase C. The treated zoospores were washed and subsequently used in an in vitro binding assay with C N V virions. Figure 3.2 shows that periodate treatment of zoospores decreased C N V binding by 72%, suggesting that carbohydrates are important for C N V binding. Trypsin digestion of zoospores reduced virus binding by 84%, indicating that proteins are also important for attachment. No decrease in C N V binding was observed using phospholipase C-treated zoospores. Together, these results suggest that proteins and/or glycoproteins on the zoospore surface play an important role in C N V binding.  113  140 .§  ]  120 "  No treatment  Trypsin  Periodate  Phospholipase C  Treatment Figure 3.2. Effects of trypsin, sodium periodate, and phospholipase C digestion on C N V binding to O. bornovanus zoospores. Zoospores were treated with 0.1% trypsin, 10 mM sodium periodate, or 5 mU of phospholipase C and used in an in vitro binding assay with 100 ug (10 pmol) of C N V particles. The amounts of virus bound were determined as for Fig. 1, using densitometry, and the results are expressed as the percentage of WT C N V binding. The results shown are the averages of triplicate samples (plus standard deviations) and are representative of three independent experiments for periodate and trypsin treatments and two independent experiments for phospholipase C.  114  3.3.3 CNV binds to specific-size proteins in O. bornovanus zoospore extracts. Virus overlay assays were conducted to further investigate the possibility that C N V recognizes specific proteins or glycoproteins present on zoospores. Total proteins from 2 X 10 zoospores (Figure 3.3, lane 9) were subjected to SDS-PAGE, transferred to 6  nitrocellulose membranes, and incubated with C N V virions. Bound virus was detected using a monoclonal antibody raised to C N V virions. Figure 3.3 (lane 2) shows that C N V bound predominantly to five low-molecular-mass proteins with estimated masses of 40, 39, 36, 34, 30, and 15 kDa and to several high-molecular-mass proteins of 119 and 63 kDa, along with a group of proteins ranging from 88 to 92 kDa. This binding pattern was observed repeatedly with different batches of zoospore preparations, but slight variations in the relative banding intensity were observed, as well as small variations in the number of high-molecular-mass proteins resolved. This was particularly true of the three proteins of -88 to 92 kDa shown in Figure 3.3 (lane 2), which sometimes resolved as only one or two species. The complexity of the banding pattern suggests that C N V virions may be recognizing a group of related proteins or a common residue on multiple proteins, such as a specific carbohydrate moiety.  3.3.4 CNV transmission mutants bind with reduced efficiency in virus overlay assays. To determine the specificity of the interaction between C N V and zoospore proteins in virus overlay assays, we used three previously characterized C N V transmission mutants (LL5, LLK10, and LLK63 [12]). The mutants differ from WT C N V by a single amino acid substitution in the C N V coat protein subunit, have reduced  115  WT C N V  1  2  LL5  3  LLK10  4  5  6  LLK63  7  8  Coomassie  9  Figure 3.3. Binding o f C N V and C N V transmission mutants to total zoospore extracts using virus overlay assays. Total O. bornovanus zoospores (2 x 10 zoospores [zoos]/lane) were subjected to S D S - P A G E , transferred to nitrocellulose membranes, and incubated with 100 ug (10 pmol) o f either W T C N V , L L 5 , L L K 1 0 , or L L K 6 3 (lanes 2, 4, 6 and 8, respectively). B o u n d virus was detected using a monoclonal antibody specific to C N V . Lanes 1, 3, 5 and 7 each contain 200 ng of either C N V , L L 5 , L L K 1 0 , or L L K 6 3 virions used as an internal detection standard respectively. Lane 9, S D S - P A G E of O. bornovanus zoospores (2 x 10 zoospores) stained with Coomassie blue. 6  6  116  transmission efficiencies (30, 27, and 14% of that of WT C N V , respectively), and have reduced binding efficiencies in solution binding assays (40, 39, and 21% of WT C N V binding, respectively) (12, 22). Figure 3.3 (lane 4) shows that LL5 binds with reduced efficiency in virus overlay assays and that little or no binding is observed when LLK10 (lane 6) and LLK63 (lane 8) are used. The reduced abilities of virions of C N V transmission mutants to bind to proteins in zoospore extracts suggests that the multiple proteins detected by WT virions are due to a specific interaction important for zoospore recognition during transmission.  3.3.5 C N V binding to O. bornovanus zoospores is competitively inhibited by several mannose-containing sugars. The possibility that C N V may be recognizing a carbohydrate moiety present on multiple glycoproteins in zoospore extracts was examined by preincubating C N V virions with specific sugars prior to adding virus in a virus overlay assay. Figure 3.4 shows that preincubation of C N V with 0.1 M methyl w-D-mannoside or 1 mg of mannan/ml abolished detectable C N V binding to O. bornovanus proteins, whereas 0.3 M D-(+)mannose showed only a slight reduction in C N V binding. The specific inhibition of C N V binding by methyl a-D-mannoside and mannan suggests that these (or closely related) sugars may represent components of several glycoproteins that C N V binds to in overlay assays.  To facilitate the analysis of the inhibitory potentials of several other sugars, we modified a previously described microtiter plate-based lectin binding assay (17) in which C N V acts as the lectin. In the modified assay, zoospores are bound to microtiter plates, and virus, in the presence or absence of a specific sugar, is incubated with bound  117  No sugar C N V zoos  1  2  Mannose C N V zoos  3  4  Methyl mannoside C N V zoos  5  6  Mannan C N V zoos  7  8  Figure 3.4. Virus overlay assays using C N V incubated with mannose, methyl a-Dmannoside or yeast mannan. O. bornovanus zoospores (2 x 10 zoospores [zoos]/lane) were subjected to SDS-PAGE, blotted, and incubated with either 100 pg (10 pmol) of C N V without sugar (lane 2) or 100 pg of C N V in the presence of either 0.3 M D-(+)mannose (lane 4), 0.1 M methyl a-D-mannoside (lane 6), or 1 mg of yeast mannan/ml (lane 8). Bound virus was detected using a CNV monoclonal antibody. Lanes 1, 3, 5, and 7 each contain 200 ng of C N V used as an internal detection standard. 6  118  zoospores. The binding of virus to the zoospores is then quantified using a CNV-specific monoclonal antibody in an enzyme-linked immunosorbent assay-based assay.  Fifteen sugars were tested for inhibitory potential (Table 3.1) by preincubating C N V with several serial 10-fold dilutions of sugar prior to adding C N V to the zoospores. Sugars showing relatively significant inhibition (arbitrarily defined as those sugars with a 50% effective concentration [EC50] of <10 mM) were further analyzed for inhibitory potential by preincubation of virus with serial threefold dilutions ofthe sugar in the concentration range where inhibition was observed using the 10-fold dilutions. Table 3.1 shows that among 15 sugars tested, mannotriose, a3,cr6-mannopentaose, methyl E T - D mannopyranoside, mannan, L-(-)-fucose, and D-mannosamine showed EC50S of <10 mM. The EC50S for these sugars varied, with mannotriose showing the strongest relative inhibitory activity (EC50 = 1 2 8 uM) and D-mannosamine showing the weakest (2.7 mM) (Table 3.1). Inhibition curves for each of these sugars are presented in Figure 3.5. Among the sugars tested for inhibitory activities, no monosaccharides except methyl W-Dmannoside, D-mannosamine, and fucose were able to inhibit C N V binding to zoospores. Sugars containing three or more marmose residues showed strong inhibition at very low concentrations. These results, like those obtained using overlay assays, suggest that mannose-containing oligosaccharides may play an important role in C N V attachment to zoospores.  3.3.6 Surfaces of O. bornavanus zoospores contain fucose and mannose and/or glucose residues. Zoospores were assessed for the presence of specific sugar residues using a variety of FITC-labeled lectin probes. ConA-FITC, which recognizes mannose and/or  119  Figure 3.5. Inhibition of C N V binding to O. bornovanus zoospores by several sugars. Several threefold dilutions of the indicated sugars were incubated with 8 pig (0.8 pmol) of C N V , and the amounts of C N V bound to plated O. bornovanus zoospores were determined using a microtiter plate assay. The absorbance at 405 nm is plotted as a function of the concentration of sugar using nonlinear regression analysis. Each data point represents the average (± standard deviation) of triplicate treatments. The results shown were obtained in at least one other independent experiment.  120  Table 3.1. Sugars classified as inhibitors and noninhibitors in viruszoospore binding assays Sugar  EC (mM) 50  Inhibitors Mannotriose a3,a6-Mannopentaose Methyl a-D-mannopyranoside Mannan L-(-)-Fucose D-Mannosamine  0.128 0.157 .1.9 2.0 .2.3 .2.7  Noninhibitors D-(+)-Glucose L-(+)-Arabinose D-(+)-Galactose /V-Acetyl-D-glucosamine L-(-)-Sorbose A^-Acetyl-D-mannosamine D-(+)-Xylose D-(+)-Mannose D-(-)-Fructose 1  Inhibitors are arbitrarily classified as those sugars with  121  EC50S  of <10 mM.  glucose, and TPA-FITC, which recognizes fucose, were the only two lectins that gave detectable binding (Figure 3.6). ConA and T P A lectins bound densely and uniformly over the surface of the zoospore body and flagella. Labelling was eliminated or reduced when the lectins were preincubated with either mannose (in the case of ConA) or fucose (in the case of TPA) (data not shown), indicating that lectin binding is a result of a specific interaction. These results indicate the presence of D-(+)-mannose and/or D-(+)-glucose and L-(-)-fucose on the surfaces of O. bornovanus zoospores.  3.4 Discussion Despite the importance of vectors in the natural spread of plant viruses, little or nothing is known about the components of vectors that viruses recognize. In this study, we have examined the possibility that a fungal vector of a plant virus contains specific receptors for virus recognition and further examined the biochemical nature ofthe putative receptor. Toward this end, we have found that the binding of C N V to zoospores is both saturable and specific (Figure 3.1), two criteria used to demonstrate receptormediated attachment of animal viruses to their receptors (28). Extrapolation from the saturation binding curve indicates that -2.7 x 10 binding sites are present on zoospores 4  (i.e., 0.018 pmol per 4 x 10 zoospores). This number is within the range of virus receptor 5  sites normally found on eukaryotic cells (34). The saturation binding curve in Figure 3.1 was also used to determine the dissociation constant (KJ). A Kj of 5.7 x 10" M was 9  obtained, indicating a very tight association between virus and zoospores. Kj values of 10" 5  to 10~ are typically found for binding of animal viruses to their cellular receptors (32). 9  122  Figure 3.6. Labeling of zoospores with FITC-labeled ConA and TPA. FITC-labeled ConA (a and b) or TPA (c and d) were incubated with zoospores and photographed using either differential interference contrast microscopy (a and c) or fluorescence microscopy (b and d). Scale, 5 mm wlO pm. (This figure was derived from work done by former graduate student, Marjorie Robbins.)  123  Reduced CNV binding following trypsin and periodate treatment of O. bornovanus zoospores, along with the results of virus overlay assays, suggests the involvement of multiple proteins and/or glycoproteins in CNV attachment. Phospholipase C did not affect CNV binding, but further experiments are required to fully explore the possibility that membrane interactions are not involved in CNV attachment. The addition of protease inhibitors to zoospores during zoospore release from fungus-infected roots did not affect the number of bands observed (data not shown), suggesting that proteolytic degradation following zoospore release is likely not a factor in the generation of multiple bands. Incubation of virus with blots under high-stringency conditions (up to 0.55 M sodium salt) also did not affect the complexity or the intensity of the signal (data not shown), suggesting that nonspecific binding of CNV is not responsible for the multiple bands. Interestingly, incubation of CNV withmannan or methyl-a-D-marmopyranoside dramatically reduced CNV binding (Figure 3.4). These experiments therefore suggested that the multiple zoospore proteins that CNV recognizes in virus overlay assays may be due to the species possessing a common oligosaccharide component.  To assess the possibility that oligosaccharides play a role in virus attachment, we tested several sugars for the ability to inhibit CNV binding (Table 3.1). Interestingly, two mannose derivatives (methyl-o-D-mannopyranoside and D-mannosamine), as well as three mannose-containing oligosaccharides (mannotriose, a3,a6-mannopentaose, and yeast mannan), were found to be strong inhibitors, with EC50S ranging from 128 u M for mannotriose to 2.7 m M for D-mannosamine. L-(-)-Fucose was also found to be an efficient inhibitor, with an EC50 of 2.3 mM. D-Martnose did not show inhibition, suggesting that  124  specific features of mannose-containing sugars are required for efficient virus-sugar interaction. Taken together, our studies suggest that C N V may have lectin-like (lectins are carbohydrate binding proteins of plant, animal and viral origin) properties that contribute to its ability to bind oligosaccharides on its vector. This observation also raises the question of whether C N V may also require oligosaccharides in or on plant cells for successful infection or multiplication in plants.  The binding of individual lectins to monosaccharides (monovalent binding) is very weak, with affinities in the range of 0.1 to 10 mM (33). Analysis of saturation binding curves of C N V to zoospores suggests that the affinity of C N V for zoospores is 5.7 nM. The apparent high affinity of C N V for zoospores may indicate the involvement of more specific receptor or receptor-like interactions that do not exclusively involve carbohydrates. On the other hand, it is now recognized that highly avid (nanomolar range) lectin interactions can exist at cell surfaces due to multivalent binding (21, 33) between lectins and oligosaccharide receptors.  CNV, MNSV, C L S V , and T N V - A virions bind to vector zoospores more efficiently than to nonvector zoospores (Figure 3.1). T N V - A differs from the other three viruses in that it lacks the C-terminal P domain. This raises the possibility that the P domain of CNV, MNSV, and C L S V enhances binding to O. bornovanus zoospores and, further, that the presence of this domain may interfere with binding to O. brassicae zoospores.  The C N V mutants LL5, LLK10, and LLK63 contain single amino acid mutations in the coat protein which decrease transmissibility and zoospore binding (12, 22), as well  125  as binding to zoospore species in virus overlay assays. The mutation in LL5 is in the shell domain at the particle quasi-threefold axis, and the mutations in LLK10 and LLK63 are in the P domain, with the LLK10 mutation facing the quasi-threefold axis. The inability of these mutants to react in virus overlay assays may suggest the involvement of the specific mutations or their potential conformational effects in binding oligosaccharides and/or glycoproteins; however, further experiments will be required to assess this potential role.  Several animal viruses are known to bind host cells via oligosaccharide or proteoglycan receptors (11, 26). In the case of influenza virus, which binds sialic acidcontaining oligosaccharides, the interaction with the sugar is sufficient for cell attachment (27). In the cases of several other animal viruses that bind proteoglycans, it is thought that the proteoglycan is used as an initial attachment receptor before further higher-affinity receptors strengthen the attachment (25). Whether acquisition of C N V requires more than one type of receptor for stable attachment remains to be determined.  Cytochemical and structural analysis of the zoospores of certain Chytrid species have indicated that the zoospore is surrounded by a polysaccharide-containing cell coat and that a common component of the coats of some chytrids is mannose (8, 20). The lectin binding studies in Figure 3.6 show that O. bornovanus zoospores contain both L - ( - ) fucose and D-(+)-mannose and/or -glucose. Indeed, several other lectins with a variety of different sugar specificities did not react with zoospores (data not shown). Thus, C N V appears to utilize sugars that are prominent on the zoospore surface for attachment.  Structural studies of the coat protein subunit of tomato bushy stunt virus, a close relative of C N V , have shown that the shell domain folds into a jellyroll-type structure  126  typical of several plant and animal virus coat protein subunits (6). Interestingly, the lectin ConA also folds into a jellyroll-type structure (6). Indeed, it has been suggested (2) that the overall similarity in structural topology between the tombusvirus capsid and ConA may indicate that tombusviruses have evolved from lectins. The studies described here support the hypothesis that C N V has lectin-like properties that may play a key role in the recognition of its vector.  127  3.5 References 1. Adams, M . J. 1991. Transmission of plant viruses by fungi. Ann. Appl. Biol. 118:479492. 2. Argos, P., T. Tsukihara, and M . G. Rossmann. 1980. A structural comparison of concanavalin A and tomato bushy stunt virus protein. J. Mol. Evol. 15:169-179. 3. Bandla, D. M., L. R. Campbell, D. E . Ullman, and J. L . Sherwood. 1998. Interaction of tomato spotted wilt tospovirus (TSWV) glycoproteins with a thrips midgut protein, a potential cellular receptor for TSWV. Phytopathology 88:99-104. 4. Campbell, R. N. 1996. Fungal transmission of plant viruses. Annu. Rev. Phytopathol. 34:87-108. 5. Campbell, R. N., S. T. Sim, and H. Lecoq. 1995. Virus transmission by host specific strains of Olpidium bornovanus and Olpidium brassicae. Eur. J. Plant Pathol. 101:273282. 6. Chelvanayagam, G., J. Heringa, and P. Argos. 1992. Anatomy and evolution of proteins displaying the viral capsid jellyroll topology. J. Mol. Biol. 228:220-242. 7. Dias, H . F. 1970. Transmission of cucumber necrosis virus and its vector, Olpidium cucurbitacaerum Barr & Dias. Virology 40:828-839. 8. Dorward, D. W., and M . J. Powell. 1983. Cytochemical detection of polysaccharides and the ultrastructure of the cell coat of zoospores of Chytriomyces aureus and Chytriomyces hyalinus. Mycologia 75:209-220. 9. Gray, S. M., and D. M . Rochon. 1999. Vectors of plant viruses, p. 1899-1910. In A. Granoff and R. Webster (ed.), Encyclopedia of virology, vol. 1. Academic Press, London, England. 10. Haas, M., M . Bureau, A. Geldreich, P. Yot, and M . Keller. 2002. Cauliflower mosaic virus: still in the news. Mol. Plant Pathol. 3:419-429. 11. Haywood, A. M . 1994. Virus receptors, binding, adhesion strengthening, and changes in viral structure. J. Virol. 68:1-5. 12. Kakani, K., J . Y. Sgro, and D. Rochon. 2001. Identification of cucumber necrosis virus coat protein amino acids affecting fungus transmission and zoospore attachment. J. Virol. 75:5576-5583. 13. Kassanis, B., and I. MacFarlane. 1965. Interaction of virus strain, fungus isolate and host species in the transmission of tobacco necrosis virus. Virology 26:603-612.  128  14. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. 15. L i , C , D. Cox-Foster, S. M . Gray, and F. Gildow. 2001. Vector specificity of barley yellow dwarf virus (BYDV) transmission: identification of potential receptors binding B Y D V - M A V in the aphid, Sitobion avenae. Virology 286:125-133. 16. Marsh, M . , and A. Helenius. 1989. Virus entry into animal cells. Adv. Virus Res. 36:107-151. 17. McCoy, J. P., J . Varani, and I. J. Goldstein. 1983. Enzyme linked lectin assay: use of alkaline phosphatase conjugated Griffonia simplicifolia B 4 isolectin for the detection of a-D-galactopyranosyl end groups. Anal. Biochem. 130:437-444. 18. McLean, J . S., R. N . Campbell, R. I. Hamilton, and D. M . Rochon. 1994. Involvement of cucumber necrosis virus coat protein in the specificity of fungus transmission by Olpidium bornovanus. Virology 204:840-842. 19. Pirone, T. P., and S. Blanc. 1996. Helper dependent vector transmission of plant viruses. Annu. Rev. Phytopathol. 34:227-247. 20. Powell, M . J . 1994. Production and modifications of extracellular structures during development of Chytridiomycetes. Protoplasma 181:123-141. 21. Rini, J . M . 1995. Lectin structure. Annu. Rev. Biophys. Biomol. Struct. 24:551-577. 22. Robbins, M . A., R. D. Reade, and D. M . Rochon. 1997. A cucumber necrosis virus variant deficient in fungal transmissibility contains an altered coat protein shell domain. Virology 234:138-146. 23. Rossmann, M . G. 1994. Viral cell recognition and entry. Protein Sci. 3:1712-1725. 24. Salas-Benito, J. S., and R. M . Del Angel. 1997. Identification of two surface proteins from C6/36 cells that bind dengue type 4 virus. J. Virol. 71:7246-7252. 25. Schneider-Schaulies, J. 2000. Cellular receptors for viruses: links to tropism and pathogenesis. J. Gen. Virol. 81:1413-1429. 26. Sharon, N., and H . Lis. 1993. Carbohydrates in cell recognition. Sci. Am. 268:8289. 27. Skehel, J. J., and D. C. Wiley. 2000. Receptor binding and membrane fusion in virus entry: the influenza virus hemagglutinin. Annu. Rev. Biochem. 69:531-569. 28. Tardieu, J., J. Weis, and H. L . Weiner. 1982. Interaction of viruses with cell surface receptors. Int. Rev. Cytol. 80:27-61.  129  29. Teakle, D. S., and C. Hiruki. 1964. Vector specificity in Olpidium. Virology 24:539-544. 30. Temmink, J. H . M . , R. N. Campbell, and P. R. Smith. 1970. Specificity and site of in vitro acquisition of tobacco necrosis virus by zoospores of Olpidium brassicae. J. Gen. Virol. 9:201-213. 31. Van den Heuvel, J. F. J. M . , S. A. Hogenhout, and F. Van der Wilk. 1999. Recognition and receptors in virus transmission by arthropods. Trends Microbiol. 7:7176. 32. Wang, J . 2002. Protein recognition by cell surface receptors: physiological receptors versus virus interactions. Trends Biochem. Sci. 27:122-126. 33. Weis, W. I. 1997. Cell-surface carbohydrate recognition by animal and viral lectins. Curr. Opin. Struct. Biol. 7:624-630. 34. Wickham, T. J., R. R. Granados, H . A. Wood, D. A. Hammer, and M . L . Shuler. 1990. General analysis of receptor-mediated viral attachment to cell surfaces. Biophys. J. 58:1501-1515.  130  4 CHAPTER FOUR Evidence that Vector Transmission of a Plant Virus Requires Conformational Change in Virus Particles* 4.1 Introduction Transmission of plant viruses in nature often involves invertebrate or fungal vectors and can be a highly specific process in which only certain vectors transmit certain viruses.(8, 10, 13, 14, 34, 46). The specificity of the transmission process has suggested that virus particles as well as vectors contain specific sites that mediate their interaction, (25, 26) and, moreover, that there may be some similarity between the way plant viruses attach to vectors and that of animal virus/host cell interactions. The coat proteins (CPs) of several plant viruses, as well as specific amino acid residues within the CP, have been shown to play important roles in transmission (5, 8, 10, 13, 14, 25, 28, 34, 35, 46). However, for the most part, the exact role of these amino acid residues in transmission, including their potential role in specific stages of the transmission process, is not known.  Cucumber necrosis virus (CNV), a member of the Tombusviridae in the genus Tombusvirus, is naturally transmitted by zoospores of the fungus, Olpidium bornovanus.(10, 12) It is believed that C N V particles are adsorbed to the surface of  * A version of this chapter has been published in the Journal of Molecular  Biology  Kakani, K., R. Reade, and D. Rochon. 2004. Evidence that vector transmission of a plant virus requires conformational change in virus particles. J. Mol. Biol. 338:507-517.  131  motile zoospores following independent release of virus and zoospores from the roots of infected plants. The virus then enters root cells upon zoospore encystment. (1, 10). Studies of C N V transmission by O. bornovanus have suggested that specific amino acid residues in the CP are important for transmission efficiency, and that a cavity at the particle quasi-3-fold axis may be an important virus attachment site (25). More recently, we have provided evidence that recognition of zoospores by C N V may involve glycoprotein receptors on the zoospore surface (26).  C N V is a 30 nm T=3 icosahedron consisting of 180 copies of a single CP subunit. The structure of the C N V particle and CP subunit is similar to that of tomato bushy stunt virus (TBSV), the type member of Tombusvirus genus. Putative structures of the C N V CP subunit and trimer have been obtained using homology modelling (25) based on the X-ray crystal structure of T B S V (17). The C N V CP subunit folds into three distinct regions termed the R N A binding (R), shell (S) and protruding (P) domains. The R and S domains are connected by a 34 amino acid residue arm, and the S and P domains are connected by a small hinge (h) (Figure 4.1(A)). The R and arm domains are located internally in the particle, the S domain forms the shell of the virus and the P domain projects from the surface. Three ordered C subunit arms are interconnected at the particle 3-fold axis to form an internal network called the /3-annulus (Figure 4.1(C)). The (3annulus stabilizes the particle and is believed to determine particle curvature during assembly.  Treatment of C N V (as well as several other small spherical plant viruses) with metal-chelating agents at alkaline pH results in particles with a swollen conformation. (49,  132  (A)  EcoRl  Pro73 Pro85  ii  Lys53  Belli  Arg95  (B)  ( Q  Figure 4.1. Location of Pro73 and Pro85 on the CNV CP subunit and P-annulus. (A) Location of Pro73 and Pro85 in the arm of the linear CP subunit. The different CP structural domains are indicated in different colours and are designated as follows: RNA binding (R, white); arm (a, yellow); shell (S, light gray); hinge (h, blue); and protruding (P, gray). The location of Pro73 and Pro85 in the arm are indicated with red arrows. Also shown are the location of the Lys53 and Arg95 tryspin cleavage sites in swollen wild-type CNV. EcoRI, Bglll and Ncol sites used for cloning the mutant CP genes and flanking sequences in the CNV genome are indicated. (B) Surface representation of homology modeled CNV CP subunit (subunit C). The different structural domains are represented in the same colours as in (A). The locations of Pro73 and Pro85 are indicated in red. The disordered R domain is not shown. (C) Representation of the CNV p-annulus showing the location of Pro73 and Pro85 residues. B and C correspond to the locations of the B an C subunits relative to the P-annulus. 133  44). The swollen conformation of TBSV is due to repulsion of subunits at the particle quasi-3-fold axis and is believed to be accompanied, in part, by movement of all or part of the R and arm domains of the A and B subunits to the outside of the particle (18). It has been suggested that the swollen conformation of plant viruses may be an important part of the uncoating process during the initial stages of virus infection (7, 36, 47).  It is known that several animal viruses undergo conformational change upon receptor attachment (11, 20, 23, 41, 51,). We wished to investigate if attachment of C N V to putative receptor(s) on vector zoospores involves conformational change. We provide evidence that zoospore-bound C N V is conformationally different from native CNV. In addition, we report that a poorly transmissible C N V mutant capable of binding zoospores in vitro fails to undergo conformational change. We discuss the possible role that conformational change may play in the C N V transmission process.  4.2 Materials and Methods 4.2.1 Virus purification A miniprep procedure was employed to partially purify particles of C N V and C N V mutants for use in initial transmission tests (35). For all other experiments, the virus was purified by differential centrifugation as described (25).  4.2.2 Maintenance of O. bornovanus cultures O. bornovanus isolate SSI96 was maintained on cucumber roots (Cucumis sativis cv. Poinsette 76) essentially as described (9).  134  4.2.3 Agarose gel electrophoresis of purified virus Virus particles were electrophoresed through 1% (w/v) agarose gels in TB buffer (40 m M Tris-borate, pH 8.3) as described (25). Virions were stained with ethidium bromide and photographed under ultraviolet illumination.  4.2.4 In vitro mutagenesis Oligonucleotide directed in vitro mutagenesis was used to produce C N V CP mutants with altered Pro73 and Pro85 residues (Table 4.1). To produce mutants Pro73Gly and Pro85Gly, an EcoRI/ Ncol fragment, encompassing the C N V CP and flanking regions (Figure 4.1(A)) in a full-length infectious cDNA clone of C N V (PK2/M5) (39) was subcloned into EcoRI/NcoI-digested pT7 Blue (Novagen) and used as a template for in vitro mutagenesis. Oligonucleotide primers used for mutagenesis are described in Table 4.1. Mutants were screened by sequencing. Selected plasmid D N A was then digested with EcoRI/NcoI and the fragment containing the mutation was cloned into similarly digested pK2/M5. pK2/M5 was used as template for the production of Pro73Ala, Pro73Cys and Pro73Leu mutants ( Table 4.1). Following mutagenesis, plasmid D N A was digested with EcoRI/Bglll and the mutated fragment was cloned back into pK2/M5 to obtain pPro73 Ala, pPro73Cys and pPro73Leu. The regions between the EcoRI/NcoI sites for pPro73Gly, pPro85Gly and the EcoRI/Bglll sites for pPro73Ala, pPro73Cys and pPro73Leu were sequenced to confirm that spurious mutations were not introduced.  135  Table 4.1. Oligonucleotides used for constructing Pro73 and Pro85 mutants Mutant  Sequence  Pro73 mutants Pro73Gly Pro73Ala Pro74Cys Pro73Leu 01igo# 117  3  Pro 8 5 mutant Pro85Gly  1  Position  2  A T T C C C G C G G C G A T A G C C C C A G (-) A T T G C C G C G G C G A T A G C C C C A G (-) ATGCACGCGGCGATAGCCCCAG(-) A T T A G C G C G G C G A T A G C C C C A G (-) C T C T T A T G C C T A T G C G G T T A A A G (+)  (2848-2827) (2848-2827) (2848-2827) (2848-2827) (2849-2871)  C T A C C T T T C C T T C C T T T A A C C G C A T A G (-) G T T T C A A A C A G C A A A A G G A T C T G T G (+)  (2883-2855) (2884-2909)  Sequences of primers are shown 5'-3'. Underlined nucleotides correspond to mutated positions and (-) and (+) antisense or sense relative to the C N V genome, respectively. Nucleotide positions are relative to the complete C N V genome (Rochon &Tremaine, 1989). 01igo#l 17 was used as (+) sense primer for all Pro73 mutants. 3  136  4.2.5 In vitro transcription and inoculation of plants Preparation of phage T7 polymerase run-off transcripts and inoculation of plants were done as described (39).  4.2.6 Fungus transmission assays Purified virions were tested for transmission by O. bornovanus zoospores essentially as described but with slight modification (25). A sample (1 /xg) of virus was incubated with 10 ml of l x l O zoospores/ml in 50 mM glycine (pH 7.6). After 15 5  minutes, the virus/ zoospore suspension was poured onto 12-16 day old cucumber seedlings. Five days later, roots of cucumber seedlings were tested for the presence of virus by DAS-ELISA, using polyclonal antisera raised against C N V particles(30). Absorbance readings five times higher than background level were considered as positive for transmission. Each transmission experiment included a wild-type C N V control, a test to determine any background level of C N V transmission in the absence of fungus, and a test for the presence of contaminating virus in zoospore preparations. Transmission in the absence of zoospores was not detectable in any of the experiments.  4.2.7 In vitro binding assays In vitro binding assays were conducted as described (25). Briefly, 100 jug of purified virus was incubated with 5><10 zoospores in 1 ml of 50 mM sodium phosphate 5  buffer (pH 7.6) for one hour. Following incubation, zoospores were centrifuged at 2700g for seven minutes in an Eppendorf microcentrifuge. Unbound virus was carefully aspirated and the zoospore pellet was washed in 1.5 ml of 50 m M sodium phosphate buffer, pH 7.6. The zoospore pellet was assayed for the presence of virus by Western blot  137  analysis using a C N V polyclonal antibody (RAD) raised against the R and arm domains of C N V CP (see below. Bound antibody was detected with goat anti-rabbit/peroxidase conjugate and an enhanced chemiluminescence detection system (Amersham Pharmacia Biotech). The quantity of virus in the pellet was determined by densitometric analysis of exposed film using the ImageQuant program (Molecular Dynamics). The amount of virus pelleted in the absence ofthe fungus was subtracted from the amount pelleted in the presence of the fungus. R A D polyclonal antibody was confirmed to react equally to wildtype C N V and Pro73Gly CP subunit in Western blot analysis.  To obtain zoospore-bound viruses for the trypsin digestion experiments, ten in vitro binding experiments were conducted separately. Following the first centrifugation, pellets were resuspended in buffer and then all ten pellets were pooled. The volume ofthe combined pellets was adjusted to 1.5 ml, and the mixture was centrifuged for seven minutes at 2700g as above. The final pellet was resuspended in sodium phosphate buffer, pH 7.6.  4.2.8 Production of polyclonal antiserum R A D antibody was produced from a synthetic C N V CPfragmentcorresponding to the R and arm domains. The R and arm domain coding region (corresponding to CP amino acid residues 1-106) was cloned into pET24d (Novagen) adjacent to a polyhistidine tag. Transformed BL21 cells were induced and fusion protein was purified by cobalt chelate chromatography. Approximately 500 f*g of protein was used for injection of rabbits. Serum from the fifth boost was used for purification of R A D IgG.  138  4.2.9 In vitro swelling of virions C N V or Pro73Gly virions (either 600 ng of virus/10 /xl of reaction volume or 10 /xg of virus/10 /xl of reaction volume) were swollen in 50 m M sodium phosphate buffer (pH 7.6), 25 mM E D T A . The swelling reaction was allowed to proceed at room temperature for 30—40 minutes.  4.2.10 Limited proteolysis of native, swollen and zoospore-bound virus Pilot experiments were conducted to determine the amount of trypsin required to distinguish native and swollen forms of C N V when digestions were conducted in a 10 /xl volume using 600 ng of virus. Approximately 600 ng of native, swollen and zoosporebound forms of C N V or Pro73Gly were digested in 50 mM sodium phosphate buffer (pH 7.6) at room temperature with 100 ng of trypsin (porcine pancreas; Sigma) in a final volume of 11 /xl. Aliquots of 1 /xl were removed at 2, 4, 8, 16, 32 and 64 minutes timepoints. Digestion was terminated by the addition of 1 /xl of 50 mM phenylmethylsulfonyl fluoride (PMSF; Sigma). Two separate 1 /xl aliquots were removed prior to the addition of trypsin and incubated for two and 64 minutes to serve as controls.  Approximately one-sixth of each sample (equivalent to 10 ng of starting material) was subjected to SDS-12% P A G E gels and subsequently transferred onto PVDF membranes. CP was detected by R A D polyclonal antibody as above, using an enhanced chemiluminescence detection system (Amersham/Pharmacia Biotech.).  Limited proteolysis was also conducted using a higher concentration of virus: 100 /xg of virus was swollen as above in a 100 /xl reaction volume and 10 /xl aliquots were incubated with 20 ng of trypsin for two to 64 minutes at room temperature. Digestions  139  were terminated as above, electrophoresed through an SDS/PAGE gel, and then stained with Coomassie brilliant blue.  4.2.11 Amino acid sequence analysis of trypsin-digested swollen virions C N V virions (10 ng) were swollen in 50 mM sodium phosphate buffer (pH 7.6), 25 mM E D T A in a 10 [il reaction volume for 30 minutes at room temperature. Swollen virions were then treated with 100 ng of trypsin for five minutes at room temperature in an 11 ill volume. The digestion products were then electrophoresed through 4-12% NuPAGE Bis-Tris gels (Invitrogen) and then transferred to a PVDF membrane (Sequiblot; BioRad). The membrane was stained with Coomassie brilliant blue, and the 33.5 kDa and 36 kDa digestion products were excised from the membrane. The aminoterminal sequence was determined using Edman degradation (Nucleic Acid and Protein Synthesis Proteomics Facility, University of British Columbia, Vancouver, B C , Canada).  4.3 Results 4.3.1 Proteolytic digestion patterns of zoospore-bound CNV differ from those of native CNV As several animal virus capsids undergo conformational change upon binding to their cellular receptors, we wished to determine whether C N V undergoes conformational change upon binding to its zoospore vector. C N V was incubated with zoospores in an in vitro binding assay as described (25). Limited proteolysis was then conducted over a 64 minute time-course. A similar time-course experiment was conducted using native C N V virions and results were analyzed by Western blot analysis. As can be seen in Figure  140  4.2(B), zoospore bound C N V is susceptible to digestion by trypsin as early as 2 minutes post-incubation, resulting in prominent 36 kDa and 33.5 kDa digestion products. In contrast, and as expected, native C N V is resistant to trypsin digestion up to 64 minutes post-incubation (Figure 4.2(A)). The differential sensitivity of zoospore-bound and native C N V to digestion by trypsin suggests strongly that C N V particles undergo conformational change upon zoospore binding.  Previous studies with TBSV and several other small spherical plant viruses have shown that the swollen forms of virus particles are proteolytically sensitive under conditions where native virions are not (15, 18). The proteolytic sensitivity of zoosporebound C N V alerted us to the possibility that the bound form of C N V may resemble the swollen state of CNV. To assess this possibility, trypsin digestion time-course experiments were conducted as described above, using C N V virions swollen in vitro. Figure 4.2 shows that the proteolytic digestion pattern observed for swollen C N V is similar to that observed using zoospore-bound C N V (compare Figure 4.2 (C) and (B)) in that 36 kDa and 33.5 kDa fragments are observed as the major proteolytic digestion products in both. This suggests that zoospore-bound C N V may resemble the swollen state of C N V . However, as can been seen in Figure 4.2, the 36 kDa and the 33.5 kDa products in zoospore-bound C N V appear to be less stable than in swollen C N V , i.e. the 36 kDa and the 33.5 kDa bands are only barely visible in zoospore-bound C N V following longer incubation times, whereas they remain visible in swollen C N V (compare Figure 4.2(B) and (C)). In addition, swollen C N V digestions frequently contain a minor 39 kDa product (see Figure 4.2(C)), whereas zoospore-bound C N V only occasionally shows a 39 kDa  141  Native  Zoospore-bound  Swollen  Swollen  2  4  8 +T  16  32  64  Figure 4.2. Time-course of trypsin digestion of native, swollen, and zoospore-bound CNV and Pro73Gly. Approximately 600 ng each of (A) native, (B) zoospore-bound and (C) swollen forms of CNV or Pro73Gly were digested with 100 ng of trypsin for two to 64 minutes. Equal volumes of digested material (equivalent to 10 ng of starting material) were loaded on to an SDS/polyacrylamide gel, blotted, and detected with a polyclonal antibody specific to the CNV R and arm domains (RAD). The digestion times (in minutes) are indicated below the lanes. Lanes 1-6 are trypsin-treated (+T) samples and lane 7 and 8 are mock treated (-T) samples for (2 and 64 minutes, respectively). (D). Trypsin digestion of CNV and Pro73Gly using 10 ug of virus and 20 ng of trypsin over a 64 minute time-course. The numbers at the left correspond to the relative molecular masses (in kDa) of the major trypsin digestion products. The asterisks (*) correspond to full-length CNV CP. 142  product (data not shown). The results of these experiments therefore suggest that zoospore-bound C N V is similar to swollen C N V , but not identical.  4.3.2 Effect of mutations of Pro73 and Pro85 on C N V particle formation To further assess the potential importance of the conformational state of C N V in zoospore binding and transmission, a panel of C N V mutants was constructed with alterations in either Pro73 or Pro85, which are highly conserved residues located within the CP arm. We chose to alter these residues because it had been suggested that these residues may be important for proper particle conformation (2). In addition, both residues are located in the interior of the particle and therefore would not likely directly affect attachment to zoospores via their specific binding to a zoospore receptor. Figure 4.3 shows an alignment of the arm region of several Tombusviridae members demonstrating conservation of both proline residues among several plant virus genera. Based on the structure of TBSV,(2) it has been suggested that the conserved proline residue corresponding to C N V Pro73 may be important for formation of the /3-annulus, since it forms a hook at the start ofthe /3-annulus ( Figure 4.1(B)) and therefore may be required for proper oligomerization of the three C-subunit arms'. Similarly, it was suggested that the Pro85 residue might be important for quasi-equivalence by serving as a molecular switch to regulate the position of the arm.  Four C N V Pro73 mutants were constructed (pPro73Gly, pPro73Ala, pPro73Cys and pPro73Leu) which contain either Gly, Ala, Cys or Leu, respectively, in place of Pro73. One C N V Pro85 mutant was constructed (pPro85Gly), which has Gly in place of Pro85 (Table 4.1). Transcripts of each ofthe mutants were inoculated onto plants to  143  Pro73 CNV (Tombusvirus) MNSV (Carmovirus) CLSV (Aureusvirus) RCNMV ( D i a n t h o v i r u s ) TBSV-Ch (Tombusvirusj TBSV-BS (Tombusvirus) TCV (Carmovirus) SBMV (Sobemovirus) SeMV (Sobemovirus) RYMV (Sobemovirus) TNV-A (Necrovirus)  Pro85  LIAHPQAFPGAIAAfjISYAYAVKGRKfgRF-QTAKG ISYTEGAKPGAISAPVAISRRVAGMKPRF-VRSEG VQNQIVGGIGAIAAP^VSITKRVRGMRPSFRQ-TKG RNRTPNTSVKTVAI PJFAKTQIIKTVN|PPKPA-RG MINHVGGTGGAIMAPVAVTRQLVGSKJJKFTGRTSG IITHVGGVGG SI M A G J V A V SRQLVGSKEKFTGRTSG ;  ™  fcx|  PVAQPVQKVTRLSAPVALAYREVSTQPJRVSTARDG  GVSMAPIAQGTMVKLRP|MLRSS GI SMAPJS AQGAMVRI RNpAV S S S QLQRAGVAQASRISGTVPGPLSS GYIVAITSGGVVTRPIVIKFSNRG  Figure 4. 3. Location of conserved proline residues in the CP arm of several small spherical plant viruses. The CP arm sequences of several small spherical plant viruses are aligned. The acronym for the virus species is indicated at the left along with the name of the genus to which it belongs. The location of the conserved Pro residues in the arm is indicated with an arrow; the numbering of the residue is based on its location in the CNV CP subunit Underlined amino acid residues are part of the arm involved in P-annulus formation in TBSV-BS and R Y M V . Abbreviations and references for sequences are as follows: CNV, cucumber necrosis virus; (38). MNSV, melon necrotic spot virus; (37). C L S V , cucumber leafspot virus; (32). R C N M V , red clover necrotic mosaic virus; (50). TBSV-Ch, cherry isolate of tomato bushy stunt virus; (16). TBSV-BS, tomato bushy stunt virus; (24). T C V , turnip crinkle virus; (22). SBMV, southern bean mosaic virus; (40) SeMV, sesbania mosaic virus; (3) R Y M V , rice yellow mottle virus; (33). T N V - A , tobacco necrosis virus-A; (31).  144  assess their ability to initiate infection and produce virions in plants (Table 4.2). A l l mutants established infections on Nicotiana benthamiana and virus particles could be purified from the leaves of infected plants (Table 4.2). The yield of particles was very low in the case of Pro73Ala (about 10% that of wild-type CNV) and even lower for Pro73Cys, Pro73Leu and Pro85Gly (each about 2%) compared to that obtained from leaves infected with wild-type C N V . However, particle accumulation in Pro73Glyinfected N. benthamiana appeared to be approximately equal to that of wild-type C N V (Table 4.2).  To further examine properties of Pro73Gly, we conducted infectivity tests and we measured accumulation in plants using double-antibody sandwich enzyme-linked immunosorbent assays (DAS-ELISA). The dilution end point of wild-type C N V on Nicotiana clevelandii was determined by inoculating plants with different concentrations of purified C N V virions (three leaves per plant; nine plants per treatment) and then observing plants for the development of systemic symptoms. Six, three and zero plants became infected when using 50, 16.6 and 5.5 pg of C N V inoculum, respectively, per leaf. Similar results were obtained with Pro73Gly virions, wherein seven, five and one plant became infected. These data therefore suggest that Pro73Gly can establish infections in N. clevelandii as well as C N V does. To test the level of accumulation in cucumber (the host used for transmission studies), Pro73Gly was inoculated onto cucumber cotyledons. Necrotic local lesions similar in size to those produced by C N V were observed; however, DAS-ELISA of leaf extracts indicated that Pro73Gly virions accumulated to approximately 40%) of wild-type C N V (data not shown). Together, these data suggest that both Pro73 and Pro85 have important roles in C N V virion accumulation. In addition, the  145  Table 4.2. Properties of C N V Pro73 and Pro85 mutants  Mutant Pro73 mutants Pro73Gly Pro73Ala Pro73Cys Pro73Leu Pro 8 5 mutant Pro85Gly  Infectivity Virus Particles 1  2  Virus Yield  3  + + + +  yes yes yes yes  Similar to WT C N V ~10%ofWT C N V ~2%ofWT CNV -2% of WT C N V  +  yes  ~2%ofWT C N V  Refers to the ability of infectious clones of the mutants to infect TV. benthamiana. Virus particle formation was determined by agarose gel electrophoresis following "mini-prep" purification from inoculated leaves. Yields were assessed by comparison of equal volumes of purified virus from an equivalent mass of infected tissue.  2  3  146  data suggest that Pro73 can be replaced by glycine but that virion accumulation in cucumber is reduced by approximately 60%.  4.3.3 Transmission efficiency of Pro73Gly virions by O. bornovanus is reduced dramatically Transmission assays were conducted using Pro73Gly virions to determine its transmission efficiency relative to wild-type C N V . Transmission efficiency was scored by determining the number of pots infected versus the number of pots inoculated. Figure 4.4 shows that 23 of 25 pots (92%> efficiency) inoculated with CNV/zoospore mixtures became infected, whereas only one of 25 pots (4% efficiency) became infected when Pro73Gly/zoospore mixtures were used. Thus, the Pro to Gly mutation at amino acid position 73 of the C N V arm appears to affect transmission efficiency much more dramatically than its effect on particle accumulation in mechanically inoculated plants.  4.3.4 Binding efficiency of Pro73Gly to zoospores is similar to that of wild-type C N V We have previously shown that the transmission deficiency of several naturally occurring C N V transmission mutants is, at least partially, due to a reduced ability to bind to vector zoospores (25, 35). We wished to assess the possibility that reduced transmission of Pro73Gly is due to its inability to bind to zoospores during the transmission process using an in vitro virus/zoospore binding assay. Figure 4.5 shows that Pro73Gly binds to zoospores with approximately the same (or increased) efficiency (90-130%)) as wild-type C N V . These results suggest that reduced transmissibility of Pro73Gly is not a result of its inefficient binding to zoospores during the transmission process.  147  100  23/25  90 80 70 60 -\ fi H NO  50 40 30 H 20 10  1/25  0 CNV  Pro73Gly Virus  Figure 4. 4. Summary of fungus transmission assays using Pro73Gly virions. Transmission assays were conducted using 1 ug of virus (either C N V or Pro73Gly) in 10 ml of zoospores (1*10 zoospores/ml). The percentage of pots showing transmission is indicated on the X-axis. The numbers on the bars indicate the number of pots showing transmission versus the number of pots tested. The data represent a compilation of five separate experiments. 5  148  140 120 to)  100 80  u  60 40 20 0 Pro73Gly  CNV Virus  Figure 4. 5. Summary of Pro73Gly in vitro virus/zoospore binding assays. A sample (100 pg) of either Pro73Gly or C N V virions were incubated with 5x 10 zoospores in an in vitro binding assay. The amount of bound virus was determined using Western blot analysis followed by densitometry. Binding efficiency was determined as the percentage of virus bound relative to the amount of WT C N V binding. The results are the average of two separate experiments (± the standard deviation) using triplicate samples of each virus per experiment. 5  149  4.3.5 The swollen conformation of Pro73Gly is different from that of CNV The location of Pro73 in the /3-annulus of virus particles suggested that the Gly mutation might influence transmissibility through its effect on the conformation of particles, since the /3-annulus is known to play an important role in virus stability. Pro73Gly particles were therefore observed by electron microscopy; however, no obvious difference in morphology in comparison to C N V particles was apparent (data not shown). In addition, agarose gel electrophoresis, which can be used to monitor conformational changes in virus particles, (21) did not reveal differences in electrophoretic mobility of Pro73Gly and C N V viruses ( Figure 4.6, compare lanes 1 and 2). However, as can be seen in Figure 4.6, swollen Pro73Gly migrates slightly slower than swollen C N V , and resolves as a more discrete band. These results suggested that the swollen conformation of Pro73Gly may be different from that of C N V , and that this may be the feature that results in loss of transmission.  Limited proteolysis of native and swollen virus was therefore conducted to further assess potential differences in the native and swollen forms of Pro73Gly and CNV. Figure 4.2 shows the results of a proteolysis experiment where it can be seen that native Pro73Gly virions remain intact throughout the incubation time, similar to what was observed with native C N V particles (Figure 4.2(A)). However, the proteolytic digestion profile of swollen Pro73Gly appears different from that of swollen C N V ( Figure 4.2(C)). In particular, and most evident, is the observation that trypsin digestion of swollen C N V results in prominent 36 kDa and 33.5 kDa products, whereas little or no 36 kDa or 33.5 kDa product is observed in trypsin-digested swollen Pro73Gly. In addition, a comparison  150  Native CNV  Swollen  Pro73Gly  CNV  Pro73Gly  Figure 4.6. Agarose gel electrophoresis of native and swollen C N V and Pro73Gly. Approximately 500 ng of native or swollen virions was electrophoresed through a 1% (w/v) agarose gel buffered in Tris-borate (pH 8.3). Virions were visualized by ethidium bromide staining in the presence of 1 m M E D T A .  151  of the relative levels of CP in swollen Pro73Gly virions treated with trypsin for 64 minutes (lane 6) versus mock-treated, swollen Pro73Gly (lane 8) with the corresponding treatments of swollen C N V particles revealed that relatively lower levels of Pro73Gly are digested. To further assess the possibility that swollen Pro73Gly particles are more resistant to trypsin digestion, a time-course experiment was conducted using a lower mass ratio of trypsin to virus (i.e. 20 ng of trypsin and 10 fig of virus). Figure 4.2(D) shows that, indeed, swollen Pro73Gly particles are more resistant to proteolysis than are swollen C N V particles. From these studies, we conclude that swollen Pro73Gly particles are different from swollen C N V particles.  4.3.6 The N-terminal region of swollen CNV particles is accessible to trypsin digestion In many small spherical viruses, conversion to the swollen state involves translocation of the N-terminal region of the CP to the particle exterior (36). In view of the observed differences in the swollen states of wild-type and Pro73Gly particles, we wished to determine the location ofthe 36 kDa and 33.5 kDa trypsin cleavage sites in swollen C N V particles. Edman degradation analysis was therefore conducted using purified 36 kDa and 33.5 kDa protein products. It was found that the 36 kDa product consisted of primarily one protein species that begins with Lys53, which is just upstream from the CP arm domain. The 33.5 kDa species appeared to be a mixture of similar-sized products, with the most prominent cleavage product corresponding to digestion at Arg95, which is located just after the arm domain (Figure 4.1(A)). Thus, as with other small spherical plant viruses, at least part of the C N V CP N-terminal domain is exposed on the particle surface. In addition, the near absence of these protein species in trypsin-treated  152  Pro73Gly particles indicates that a major structural difference between swollen wild-type and Pro73Gly particles is the near absence of the N-terminal coat protein region on the surface of particles.  4.3.7 Zoospore-bound Pro73Gly does not undergo conformational change Our in vitro binding experiments in Figure 4.5 indicate that the poor transmissibility of Pro73Gly cannot be attributed to inefficient binding to zoospores. However, the observation that the swollen conformation of Pro73Gly is different from that of C N V (Figure 4.2), along with our observation that C N V undergoes a conformational change similar to that of swollen C N V upon zoospore binding ( Figure 4.2), raised the possibility that Pro73Gly may be poorly transmitted due to an inability to form the proper conformation upon zoospore binding. To test this possibility, we conducted trypsin digestion experiments on zoospore-bound Pro73Gly, as was conducted on wild-type zoospore-bound C N V ( Figure 4.2). Figure 4.2(B) shows that, unlike zoospore-bound wild-type C N V , zoospore-bound Pro73Gly particles are resistant to trypsin digestion throughout the 64 minute digestion reaction. These results suggest that the inability of Pro73Gly to be transmitted may be a result of its inability to undergo conformational change upon zoospore binding.  4.4 Discussion Previous work in our laboratory has shown that specific amino acid residues in the C N V CP S and P domains are important for zoospore attachment and subsequent transmission by O. bornavanus (7.) Here, we have examined the possible involvement of the internally located CP arm in fungus transmission, focusing on two proline residues in  153  the arm region. Substitution of either residue results in decreased particle accumulation in most cases (Table 4.2), suggesting the importance of the proline residue in either particle assembly, disassembly or stability. However, substitution of proline 73 with glycine had a less deleterious effect, and had no observable effect in N. clevelandii. Pro73 forms part of the /3-annulus, which is an important intermediate in assembly of particles, possibly by facilitating formation of trimers of dimers at the particle 3-fold axis (42). The observation that Pro73Gly particles accumulate to wild-type levels in N. clevelandii and to 40% of wild-type levels in cucumber (Table 4.2) suggests that substitution with Gly does not have a strongly detrimental effect on /3-annulus formation, or that the /3-annulus may not be absolutely required for virus assembly (48).  It is believed that Pro85 may be important for virus assembly by controlling the arm position and acting as a molecular switch to regulate the ratio of the two types of conformations required ofthe C and A/B subunits to satisfy quasi-equivalence (2). It is possible that the Pro to Gly substitution in Pro85Gly reduces the efficiency of particle formation by engendering an unacceptably high number of conformational states in the arm and thereby restricting efficient oligomerization during particle assembly.  The data presented here suggest that successful transmission of C N V requires conformational change of virus particles during or following the attachment phase of transmission. We believe that this is the first demonstration that a plant virus undergoes conformational change during acquisition by its vector. It also is the first time that in vivo evidence has been obtained for the biological significance of the swollen state of a small spherical plant virus.  154  Our studies indicate that zoospore-bound C N V resembles C N V particles swollen in vitro. It is believed that many viruses have a highly dynamic character in vivo (27, 4) and conformational changes in capsid structures are purported to be associated with the initial stages of infection in many viruses (11, 23, 51, 41, 20). In the well-studied poliovirus system, receptor attachment initiates a series of conformational changes that allow the virus to attach to membranes, to form a pore in the membrane and to release viral R N A from the capsid (23). It has been noted that the structural states associated with the various transitions of poliovirus particles are highly analogous to the expanded states of structurally similar plant viruses. In particular, the rotation and outward movement of poliovirus VP2 and VP3 along the 3-fold axis is analogous to the expansion at the quasi3-fold axis in plant virus capsids. In addition, the outward movement of the aminoterminal region of VP1 at the 5-fold axis is analogous to the extrusion of the plant virus A and B subunit N termini (23, 19). Indeed, these similarities have previously prompted the suggestion that the swollen state of plant virus capsids may correspond to an important intermediate during the initial stages of infection. Our observation that zoospore-bound C N V virions are structurally different from native virus, and that they resemble virions swollen in vitro, suggests that the biological significance of the swollen state may lie, at least in part, in the vector attachment process. Although further experiments are required to determine the extent of similarity between the swollen and bound forms of C N V , our results clearly show that conformational change is associated with zoospore attachment. Zoospore-bound C N V is proteolytically more sensitive than native virions (Figure 4.2(A) and (B)). This might reflect the dynamics of externalization in the native virus and the possibility that the arms of bound virus are trapped in a more exposed state.  155  Previous studies with other spherical plant viruses have shown that virions swollen in vitro are more readily translated in cell-free translation systems (7, 47, 6), suggesting that swelling in vivo may be associated with viral uncoating and concomitant translation ofthe genome. It is possible that the virus uncoating mechanism in vivo occurs on membranes within cells and is mechanistically similar to the vector attachment process.  Interestingly, exposure of Pro73Gly particles to in vitro swelling conditions results in particles with reduced mobility on agarose gels similar to that observed with swollen wild-type C N V particles (Figure 4.6). The reduced mobility of swollen C N V particles is likely due to the increase in particle size following repulsion of the three subunits at the quasi-3-fold axis. The observation that Pro73Gly particles also migrate more slowly suggests that they too undergo expansion. However, as shown in Figure 4.2, trypsin digestion of swollen Pro73Gly particles does not result in the same high-level accumulation of the 36 kDa and 33.5 kDa products seen in swollen C N V digestions, although these products can be observed when large amounts of swollen mutant particles are digested (see Figure 4.2(D), lanes 5 and 6). Thus, although Pro73Gly particles are expanded at the quasi-3-fold axis, it appears as though the N-terminal region of a significant proportion ofthe CP subunits have not translocated to the particle exterior.  Pro73Gly particles bind zoospores efficiently (Figure 4.5) but are largely insensitive to trypsin digestion (Figure 4.2(B)). This suggests that reduced transmission of Pro73Gly virions is due to a defect in the ability of particles to undergo some aspect of conformational change during zoospore attachment. As described above, binding of  156  poliovirus to its cellular receptor has been shown to result in distinct conformational changes and it has been suggested that the receptor acts as a catalyst to promote the required changes (45). In addition, in the case of human rhinovirus, the ability of the virus to undergo conformational change has been related directly to its ability to infect cells (29). Previous work in our laboratory has suggested that specific oligosaccharides and/or glycoproteins present on the zoospore surface act as receptors for C N V attachment (26). In analogy to that observed in poliovirus, we hypothesize that binding of C N V to its zoospore receptor may facilitate conformational change in C N V , and that this conformational change is required to complete subsequent stages of the transmission process. Our experiments did not assess if bound Pro73Gly particles have undergone the expansion aspect of swelling, but the observation that they do expand under in vitro conditions supports the notion that binding to zoospores fails to induce particles to extrude the N-terminal domain. Thus, at a minimum, extrusion of the N-terminal aims' appears to be highly essential for transmission.  It is believed that the main function of the zoospore in C N V transmission is to provide a means for entry of C N V into otherwise impermeable root cells. In consideration of the proposed parallels between poliovirus entry into host cells and C N V attachment to zoospores, it is possible that zoospores may play a more significant role than has been recognized so far. For example, attachment may contribute to root cell infection by "priming" particles for translation in the root cytoplasm. Alternatively, C N V may enter zoospores prior to root cell infection. Indeed, electron microscopy of CNV/zoospore-infected plants has suggested that C N V uptake by O. bornovanus zoospores involves endocytosis (43). In addition, it is known that transmission of some  157  plant viruses by Olpidium brassicae is believed to involve uptake of virus particles in the co-infected plant (10).  Our finding that C N V undergoes conformational change upon zoospore attachment raises the possibility that a similar change might be important for other aspects of the virus infection cycle. For example, it is possible that the uncoating process may occur on membranes within an initially infected cell. Further experiments are required to investigate this possibility.  In summary, the results of our experiments suggest strongly that C N V undergoes conformational change as part of the transmission process. Moreover, our data suggest that the basis for the reduction in transmission of Pro73Gly particles is due to the inability of the A and B subunit arms to be translocated to the surface. Thus, as has been suggested in poliovirus, extrusion of the arms of C N V may be important for interaction with the zoospore membrane and possibly for entry of the virus or viral nucleic acid into zoospores.  158  4.5 References 1. Adams, M . J. 1991. Transmission of plant viruses by fungi. Ann. Appl. Biol. 118:479-492. 2. Bergdoll, M . , M.-H. Remy, C. Cagnon, J.-M. Masson and P. Dumas. 1997. Proline dependent oligomerization with arm exchange. Structure 5:391-401. 3. Bhuvaneshwari, M . , H. S. Subramanya, K. Gopinath, H . S. Savitri, M.V. Nayudu and M . R. N. Murthy. 1995. Structure of sesbania mosaic virus at 3 A resolution. Structure 3:1021-1030. 4. Bothner, B., S. F. Dong, L . Bibbs, J. E . Johnson and G. Siuzdak. 1998. Evidence of viral capsid dynamics using limited proteolysis and mass spectrometry. J. Biol. Chem. 273:673-676. 5. Brault, V., M . Bergdoll, J.Mutterer, V. Prasad, S. Pfeffer, M . Erdinger, K. E . Richards, and V. Ziegler-Graff. 2003. Effects of mutations in the major capsid protein of beet western yellows virus on capsid formation, virus accumulation, and aphid transmission. J. Virol. 77:3247-3256. 6. Brisco, M . , R. Hull and T.M.A Wilson. 1985. Southern Bean Mosaic virus-specific proteins are synthesized in an in vitro system supplemented with intact and treated virions. Virology 143:392-398. 7. Brisco, M . , R. Hull and T.M.A Wilson. 1986. Swelling of isometric and bacilliform plant virus nucleocapsids required for virus specific protein synthesis in vitro. Virology 148:210-217. 8. Brown, D. J. F., W. M . Robertson, and D. L . Trudgill. 1995. Transmission of viruses by plant nematodes. Annu. Rev. Phytopathol. 33:223-249. 9. Campbell, R. N., S. T. Sim, and H . Lecoq. 1995. Virus transmission by host-specific strains of Olpidium bornovanus and Olpdium brassicae. Eur. J. Plant Pathol. 101:273282. 10. Campbell, R. N. 1996. Fungal transmission of plant viruses. Annu. Rev. Phytopathol. 34:87-108. 11. Chandran, K., and M . L . Nibert. 2003. Animal cell invasion by a large nonenveloped virus: reovirus delivers the goods. Trends in Microbiol. 11:374-3821 12. Dias, H . F., 1970. Transmission of cucumber necrosis virus by Olpidium cucubitacaerum Barr & Dias. Virology 42:828-839.  159  13. Gray, S. M . 1996. Plant virus protein involved in natural vector transmission. Trends in Microbiol. 4:259-264. 14. Gray, S. M . , and D. M . Rochon. 1999. Vectors of plant viruses, p. 1899-1910. In A. Granoff and R. Webster (ed.), Encyclopedia of virology, vol. 1. Academic Press, London, England. 15. Golden, J. S, and S. C. Harrison. 1982. Proteolytic dissection of turnip crinkle virus in solution. Biochemistry 21:3862-3866. 16. Haerne, P. Q., D. A. Knorr, B. I. Hillman, and T. J. Morris. 1990. The complete genome structure and synthesis if infectious R N A from clones of tomato bushy stunt virus. Virology 177:214-218. 17. Harrison, S. C., A. J . Olson, C. E . Schutt, F. K. Winkler, and G. Brigogne. 1978. Tomato bushy stunt virus at 2.9 A ° resolution. Nature (London) 276:368-373. 18. Harrison, S. C., P. K. Sorger, P.G. Stockley, J . Hogle, R. Altman and R. K. Strong. 1987. Mechanism of R N A virus assembly and disassembly, in Positive Strand Viruses (Brinton M A , Rueckert RR, eds), Alan R. Lis Inc. pp. 379-395. 19. Harrison, S.C., 1989. "Common Features in the Design of Small R N A Viruses" in Concepts in Viral Pathogenesis III .p 3-19 Springer-Verlag New York, Inc. 20. Haywood, A. M . , 1994. Virus receptors: Binding, adhesion strengthening, and changes in viral structure. J. Virol. 68:1-5. 21. Heaton, L . A., 1992. Use of agarose gel electrophoresis to monitor conformational changes of small, spherical plant viruses. Phytopathology 82:803-807. 22. Hogle, J. M . , A. Maeda, and S. C. Harrison. 1986. Structure and assembly of turnip crinkle virus. I. X-ray crystallographic structure analysis at 3.2 A resolution. J. Mol. Biol. 191:625-638. 23. Hogle, J. 2002. Poliovirus Cell Entry: Common structural themes in viral entry pathways. Annu. Rev. Microbiol. 56:677-702. 24. Hopper, P., S. C. Harrison, and R. T. Sauer. 1986. Structure of tomato bushy stunt virus V. Coat protein sequence determination and its structural implications. J. Mol. Biol. 177:701-713. 25. Kakani, K., J. Y . Sgro, and D. Rochon. 2001. Identification of cucumber necrosis virus coat protein amino acids affecting fungus transmission and zoospore attachment. J. Virol. 75:5576-5583.  160  26. Kakani, K., M . Robbins, and D. Rochon. 2003. Evidence that binding of cucumber necrosis virus to vector zoospores involves recognition of oligosaccharides. J. Virol. 77:3922-3928. 27. L i , Q., A . G.Yafal, Y . M - H . Lee, J. Hogle, and M . Chow. 1994. Poliovirus neutralization by antibodies to internal epitopes of VP4 and VP1 results from reversible exposure of these sequences at physiological temperature. J. Virol. 68:3965-3970. 28. Liu, S., X. He, G. Park, C. Josefsson, and K. Perry. 2002. A conservedcapsid protein surface domain of cucumber mosaic virus is essential for the efficient aphid vector transmission. J. Virol. 76:9756-9762. 29. Lewis, J.K., B. Bothner, T. J. Smith, and Siuzdak, G. 1998. Antiviral agent block breathing of the common cold virus. Proc. NatT Acad. Sci. U S A 95:6774-6778. 30. McLean, M . A . , R. N. Campbell, R. I. Hamilton, and D. Rochon. 1994. Involvement of cucumber necrosis virus coat protein in the specificity of fungus transmission by Olpidium bornovanus. Virology 204:840-842. 31. Meulewaeter, F. J. Seurinck, and J. Van Emmelo. 1990. Genome structure of tobacco necrosis virus strain A. Virology. 177:699-709. 32. Miller, J.S., H. Damude, M . A . Robbins, R. D. Reade, and D. M . Rochon. 1997. Genome structure of cucumber leaf spot virus, sequence analysis suggests it belongs to a distinct species within the Tombusviridae. Virus Research 52:51-60. 33. Ngon a Yassin, M . , C. Ritzenthaler, C. Brugidou, C. M . Fauquet, and R. N. Beachy. 1994. Nucleotide sequence and genome organization of rice yellow mottle virus RNA. J. Gen. Virol. 75:249-257. 34. Pirone, T. P., and S. Blanc. 1996. Helper dependent vector transmission of plant viruses. Annu. Rev. Phytopathol. 34:227-247. 35. Robbins, M . A., R. D. Reade, and D. M . Rochon. 1997. A cucumber necrosis virus variant deficient in fungal transmissibility contains an altered coat protein shell domain. Virology 234:138-146. 36. Robinson, I.K., and S. C. Harrison. 1982. Structure of the expanded state of tomato bushy stunt virus. Nature 297:563-568. 37. Riviere, C. J., and D. M . Rochon. 1990. Nucleotide sequence and genome organization of melon necrotic spot virus. J. Gen. Virol. 71:1887-1896. 38. Rochon, D., and J. H . Tremaine. 1989. Complete nucleotide sequence of cucumber necrosis virus genome. Virology 169:251-259.  161  39. Rochon, D. M . , and J. C. Johnston. 1991. Infectious transcripts from cloned cucumber necrosis virus cDNA: evidence for a bifunctional subgenomic mRNA. Virology 181:656-665. 40. Silva, A . M . , and M . G. Rossman. 1987. Refined structure of southern bean mosaic virus at 2.9 A resolution. J. Mol. Biol. 197:69-87. 41. Skehel, J. J., and D. C. Wiley. 2000. Receptor binding and membrane fusion in virus entry: the influenza virus hemagglutinin. Annu. Rev. Biochem. 69:531-569. 42. Sorger, P. K., P. G. stockley, and S. C , Harrison. 1986. Structure and assembly of turnip crinkle virus II. Mechanism of reassembly in vitro. J. Mol. Biol. 191:639-658. 43. Stobbs, L . W., G. W. Cross, and M . S. Manocha. 1982. Transmission of cucumber necrosis virus by Olpidium radicale zoozspores. Can. J. Plant Pathol. 4:134-142. 44. Speir, J.A., S. Munshi, W. G. Wang, T. S. Baker and J . E . Johnson. 1995. Structures of the native and swollen forms of cowpea chlorotic mottle virus determined by X-ray crystallography and cryo-electron microscopy. Structure 3, 63-78. 45. Tsang, S.K., B. M . McDermott, V. R. Racaniello, and. J. M . Hogle. 2001. Kinetic analysis of the effect of Poliovirus receptor on viral uncoating: the receptor as a catalyst. J. Virol. 75: 4984-4989. 46. Van den Heuvel, J. F. J. M . , S. A . Hogenhout, and F. van der Wilk. 1999. Recognition and receptors in virus transmission by arthropods. Trends Mi-crobiol. 7:71— 76. 47. Verduin, B., 1990. Early interactions between viruses and plants. Semin. Virol. 3:423-431. 48. Willits, D., H. Zhao, N. Olson, T. S. Baker, A . Zlotnick, J. E . Johnson, T. Douglas and M . J . Young. 2003. Effects of the cowpea chlorotic mottle bromovirus 8hexamer structure on virion assembly. Virology 306:280-288. 49. Witz, J., and F. Brown. 2001. Structural dynamics, an intrinsic property of viral capsids. Arch. Virol. 146:2263-2274. 50. Xiong, Z., and S. A . Lommel. 1989. The complete nucleotide sequence and genome organization of red clover necrotic mosaic virus RNA-1. Virology 171:543-554. 51. Young, J. A . T. 2001 Virus entry and uncoating. In Fields virology Knipe, D . M and Howley, P. M . , eds pp 87-103, Lippincott Williams and Wilkins.  162  5 CHAPTER FIVE GENERAL DISCUSSION This thesis explored some of the molecular determinants (viral and vector) required for vector transmission of C N V by its fungal vector O. bornovanus. One goal was to determine if any specific sequences on the C N V coat protein are important in recognizing zoospores during the attachment phase of the transmission process. Previous work in this lab showed that the CP plays an important role in determining the specificity of C N V transmission by O. bornovanus. It has been shown that continuous passage of plant viruses in the absence of the vector would result in variants that are deficient in vector transmission. Robbins et al. (1997) isolated and characterized LL5, one such naturally occurring transmission deficient mutant. It was shown that specific regions on the shell domain of C N V CP play an important role in fungus transmission. To further identify the regions important for C N V transmission the methodology developed for the characterization of LL5 was followed. We isolated and characterized six distinct, naturally occurring C N V transmission mutants. Each mutant was found to contain an amino acid substitution either in CP shell or protruding domains which is responsible for transmission deficiency. Transmission efficiencies of these mutants were variable (Table 2.1) and in vitro virus-zoospore binding assays showed that each mutant binds zoospores less efficiently than WT CNV, suggesting that reduced transmissibility is at least partially due to inefficient zoospore binding. A n artificial double mutant containing mutations present in both protruding and shell domains was shown to bind and transmit less efficiently by zoospores, reinforcing the role of both mutations in  163  attachment and transmission processes. In order to determine whether these mutated amino acids are exposed and clustered on the coat protein tertiary and quaternary structures, homology modelling of the C N V CP subunit, asymmetric unit and particle was conducted based on the closely related X-ray crystal structure of T B S V (Figure 2.3). Mapping of these mutated amino acids indicated most of the mutants are exposed on the surface of the particle. Several of these mutated amino acids are located in or near a cavity formed by the trimer at the particle quasi three-fold axis, suggesting that this region may be important in recognizing a putative receptor on the zoospore surface. It is not known whether these mutations contribute to reduced transmissibility by affecting interaction with the putative receptor or indirectly by changing the virion conformation. Isolation and characterization of more naturally occurring transmission mutants and mutagenic analysis of conserved amino acids in the quasi three-fold axis cavity in C N V or other related T-3 fungally transmitted viruses (CLSV, M N S V and TNV) might be useful in determining the further importance of this region in fungus transmission. Previous work from our lab has provided the preliminary evidence for the presence of receptor or receptor-like molecules on the zoospore surface (8). It was shown that binding of C N V to zoospores is saturable, indicating a discrete number of recognition molecules on zoospores for CNV. In addition, in vitro binding studies have demonstrated efficient binding between viruses and their respective vector zoospores and that this reflects their association in nature (8,9). In an attempt to further characterize the biochemical nature of the molecule(s) on the surface of O. bornovanus zoospores involved in C N V attachment, zoospores were treated with periodate, trypsin and phospholipase-C. Reduced C N V binding to pre-treated  164  zoospores suggested that glycoproteins on the zoospore surface may mediate C N V attachment. C N V bound to several specific-sized proteins in virus overlay assays, whereas C N V transmission mutants failed to bind or bound at significantly reduced levels. The possible involvement of specific sugars in virus binding was investigated using sugar inhibition studies. It was found that mannose-containing oligosaccharides and fucose inhibited C N V binding at relatively low concentrations. These studies suggest that binding of C N V to zoospores is mediated by specific mannose-/or fucose-containing oligosaccharides or glycoproteins. Interestingly previous studies from our lab revealed the presence of mannose/glucose and fucose containing sugars on the surface of the zoospores, reinforcing the possible involvement of these sugars as receptors or a part of receptors for C N V attachment (8). In addition chytrid zoospores are known to contain a prominent cell coat predominantly made of mannose-containing sugars (6). Carbohydrates play an important role as biological recognition molecules in critical cellular functions such as signal transduction, immune modulation and enzyme catalysis (2). Carbohydrates are excellent carriers of information due to their complex nature and structural diversity (11). The ability of cell surface carbohydrates to act as receptors is usually dependent on: 1) the presence of a particular sugar; 2) accessibility; and; 3) mode of presentation (for example, linkages in oligosaccharides) (11). Oligosaccharides are used as primary, as well as secondary receptors by different animal viruses (Table 1.5). Several animal viruses that recognize oligosaccharides as their entry receptors are considered as viral lectins. Lectins are carbohydrate binding proteins of plant and animal origin that recognize oligosaccharides present on cell surfaces. The tail spike protein (TSP) of Salmonella phage 22, haemagglutinin (HA) of influenza viruses,  165  VP1 of polyomaviruses and the VP4 of rotaviruses, are all known to contain a lectin-like fold in their proteins which harbour an oligosaccharide binding site (13, 3). The ability of several mannose-containing oligosaccharides to inhibit C N V binding to zoospores suggests that C N V has lectin-like properties that may play an important role in recognition of its fungal vector. Interestingly, the lectin Con-A has ajellyroll topology, which is a common structural motif found in several viral capsids. It has been suggested that there is broad structural similarity between Con-A and the tombusvirus capsid (1) This may indicate that tombusviruses have evolved from lectins. Taken together, these studies support the hypothesis that C N V has lectin-like properties that may play an important role in the recognition of its vector. In the case of some animal viruses that bind to proteoglycans, it is thought that the proteoglycan is used as initial attachment receptor before a further high affinity receptor strengthens the attachment (4, 10). Whether mannose-containing oligosaccharides are acting as primary or secondary receptors for C N V attachment remains to be determined. Future work may involve the testing of these sugars as inhibitors of transmission of C N V and other related viruses by Olpidium. Also, structural determination of C N V complexed with a sugar (mannotriose or a3-, a6-mannopentaose) either by X-ray crystallography or by cryo-electron microscopy image reconstruction techniques will be useful in determining the sugar binding region on the C N V capsid. Further identification of zoospore proteins involved in C N V recognition could involve 2-D gel electrophoresis of zoospore proteins (either total or membrane-associated) followed by overlay assays using C N V virions as a probe. Proteins that specifically bind to C N V can then be digested with trypsin and identified using mass spectrometry. It may be possible to obtain  166  c D N A clones corresponding to the proteins and to use those clones or expressed proteins to further assess their role as receptor(s) for CNV. Limited trypsin digestion of zoospore-bound, swollen and native (unbound) C N V revealed that zoospore-bound C N V is conformationally different and resembles the swollen conformation. In addition we have shown that reduced transmission of Pro73Gly, a C N V transmission-deficient mutant was due to its inability to undergo this conformational change. Together, these results suggest that conformational changes in virus particles are important for fungus transmission of CNV. During poliovirus entry into the host cell, the virus particle undergoes expansion upon binding to the receptor in which CP subunits VP1, VP2 and VP3 move outwardly creating significant gaps at the base of five-fold axis. The internally located N-terminal regions of VP1 and VP4 come out through these openings to form an amphipathic helical bundle, which not only promotes virion binding to the cellular membrane, but also causes a pore or disruption through which viral R N A enters the cell (5). Similar to the poliovirus system, our data shows that C N V undergoes a swollen-like conformation in which the N terminal arms of the A and B subunits are exposed. This structural similarity could indicate functional homology between CNV/zoospore interaction and poliovirus/host cell interaction. In the case of zoospore-bound C N V , the externalized arms could interact with the zoospore membrane resulting in stabilization of the virus/zoospore interaction and/or release of viral R N A into the zoospore during transmission. Interestingly, parallel studies in our lab have suggested the presence of a putative transmembrane domain in the arm region of the C N V CP. Green fluorescent protein (GFP) tagging experiments revealed that the arm is specifically targeted to chloroplast membranes (D. Rochon, Y . Xiang, &  167  R. Reade, unpublished observations). These studies suggest that externalization ofthe arm may promote CNV/zoospore membrane interactions. It would be interesting to investigate whether the conformational changes induced by zoospore binding are also important for critical steps of virus infection inside the plant cell, such as disassembly. It is also possible that the requirement for conformational change in virus particles could be a common phenomenon in plant virus/vector interactions. In the case of potyviruses these conformational changes could be induced by virus interaction with helper factor, since it has been suggested that helper factor play a critical role in determining the specificity of aphid transmission. It is generally well-accepted that virions bind to the outside of zoospores as a means for zoospore assisted entry into root cells. However, exactly how bound virus enters the root cytoplasm is not clear. Temmink and Campbell (12) suggested that virus attached to the flagellar sheath is transferred to the zoospore cytoplasm upon flagellar retraction and that C N V enters root cytoplasm via injection of the zoospore cytoplasm. However, this would require that C N V crosses the zoospore membrane to enter the host cell. It seems more likely that virus bound to the zoospore plasmalemma is injected into the root cytoplasm during injection ofthe zoospore protoplast. In this case, virus would be outside ofthe protoplast and within the root cytoplasm. The final stages of viral uncoating would then take place in the root cytoplasm. A major significance of the work described in this thesis is the emerging similarities between plant virus/vector interactions and animal virus/host cell receptor interactions. In addition, the work in this thesis is the first to demonstrate: 1) that specific regions ofthe virus capsid are important for vector attachment and transmission; 2) the  168  role of specific sugars in vector attachment and; 3) the importance of conformational changes in the virus particle for vector transmission. Based on the work described in this thesis we propose the following model (9, Figure 5.1) for C N V binding to O. bornovanus zoospores during transmission: 1) C N V recognizes a glycoprotein or oligosaccharide receptor on the surface of the zoospore wherein the cavity at the C N V particle quasi three-fold axis contains important sugar binding elements; 2) upon binding to the putative receptor, the C N V particle acquires a swollen or swollen-like conformation in which the A, B, and C subunits move away from each other to form openings through which disordered arms of the A and B subunits are translocated to the exterior of the particle; and 3) the hydrophobic portions of the A and B arms interact with the zoospore membrane resulting in stabilization of virus/zoospore interaction (9; Fig. 5.1).  169  Quasi three-fold axis  glycoprotein  Figure 5.1 Model for C N V binding to O. bornovanus zoospores. (A) A portion of the C N V particle showing the icosahedral five-fold and quasi three-fold axis. The A , B and C subunits are represented in red, blue and green colors respectively. Yellow circles represent the predicted positions where the A and B subunit arms come out during expansion. (B) Possible steps involved in the interaction between C N V and zoospore during the attachment phase of the transmission process. Onlythe A and B subunits are shown. This figure was adapted and modified from reference # 9 170  5.1 References 1. Argos, P., T. Tsukihara, and M . G. Rossmann. 1980. A structural comparison of concanavalin A and tomato bushy stunt virus protein. J. Mol. Evol. 15:169-179. 2. Baranowski, E . , C. M . Ruiz-Jarabo, N. Pariente, N. Verdaguer, and E. Domingo. 2003. Evolution of cell recognition by viruses: A source of biological novelty with medical implications. Adv. Virus Res. 62:19-110. 3. Dormitzer, P. R., Z. Y. J. Sun, G. Wagner, and S. C. Harrison. 2002. The rhesus rotavirus VP4 sialic acid binding domain has a galectin fold with a novel carbohydrate binding site. E M B O . J. 21:885-897. 4. Haywood, A. M . 1994. Virus receptors, binding, adhesion strengthening, and changes in viral structure. J. Virol. 68:1-5. 5. Hogle, J. M., A. Maeda, and S. C. Harrison. 1986. Structure and assembly of turnip crinkle virus. I. X-ray crystallographic structure analysis at 3.2 A resolution. J. Mol. Biol. 191:625-638. 6. Powell, M . J. 1994. Production and modifications of extracellular structures during development of Chytridiomycetes. Protoplasma 181:123-141 7. Robbins, M . A., R. D. Reade and D. M . Rochon. 1997. A cucumber necrosis virus variant deficient in fungal transmissibility contains an altered coat protein shell domain. Virology 234:138-146. 8. Robbins, M . A. 2000. Molecularcharacterization of the interactions between cucumber necrosis virus and zoospores of its fungal vector, Olpidium bornovanus. Ph.D. Thesis, Department of Plant Science, University of British Columbia, Vancouver, British Columbia, Canada. 9. Rochon, D., K. Kakani, M . Robbins, and R. Reade. 2004. Molecular aspects of plant virus transmission olpidium and plasmodiophorids vectors. Annu. Rev, Phytopathol. 42:211-241. 9. Schaulies, J. S. 2000. Cellular receptors for viruses: Links to tropism and pathogenesis. J. Gen. Virol. 81:1413-1429. 11. Sharon, N., and H . Lis. 1993. Carbohydrates in cell recognition. Sci. Am. 268:8289. 12. Temmink, J. H. M . , and R. N. Campbell. 1969. The ultrastructure of Olpidium brasscae. II. Zoospores. Can. J. Bot. 47:227-231.  171  13. Weis, W. I. 1997. Cell-surface carbohydrate recognition by animal and viral lectins. Curr. Opin. Struct. Biol. 7:624-630.  172  6 CHAPTER SIX APPENDIX 6.1 Introduction Isolation and characterization of several naturally occurring C N V transmission mutants showed that specific regions of the C N V capsid are important for transmission by O. bornovanus. Two regions, one on the protruding domain facing the cavity at the particle quasi three-fold axis and the other region on the particle shell at the quasi threefold axis could play an important role in recognizing zoospores during the attachment phase of transmission (2; see Chapter 2). We have also showed that C N V acquires a swollen-like conformation upon binding to the zoospores, and the extrusion of the internally located arms during this conformational change is important for fungus transmission (3). In the present preliminary study, we utilized in vitro mutagenesis to further assess the importance of the region on the CP protruding domain that is predicted to be important for fungus transmission. Two non-conserved proline residues in the arm and two additional proline residues,(one, a conserved proline at the base of the hinge, and another in the middle of the hinge) were also mutated to assess their effect on fungus transmission (Figure 6.1).  173  Figure 6.1. Surface representation of the C N V CP subunit (subunit C) in different orientations showing location of mutated amino acids. Different structural domains are indicated in different colors as follows: arm (yellow); shell (light gray); hinge (blue) (not visible in B); and protruding (P, gray). The disordered R domain is not shown. (A) Location of Pro63, Pro67, Pro85, Pro259, Pro262, Ser338 and Ser339 (B) Location of Thr292, Leu353, Trp296, and Gln297 on the protruding domain. All mutated amino acids are colored in red except Ser338 and Trp296, which are represented in pink to distinguish from adjacent Ser339 and Gln297 respectively.  174  6.2 Materials and Methods 6.2.1 Virus purification Partially purified virus particles of wild type C N V and C N V mutants were obtained by a miniprep procedure (4).  6.2.2 Maintenance of O. bornovanus cultures O. bornovanus isolate SSI96 was maintained on cucumber roots (Cucumis sativus cv. Poinsette 76) essentially as described (1).  6.2.3 In vitro mutagenesis Oligonucleotide-directed in vitro mutagenesis was used to produce C N V CP mutants (Table 6.2). To produce mutants Thr292Ala, Asp293Ala, Trp296Ala, Gln297Ala, Ser338Ala, Ser338Phe, Ser338Thr, Ser339Gly, Leu353Ala, Pro63Gly, Pro63Ala, and Pro85Ala, an EcoRll Ncol fragment encompassing the C N V CP and flanking regions (Figure 4.1(A)) in a full-length infectious cDNA clone of C N V (pK2/M5) (5) was subcloned into iscoRI/TVcoI-digested pT7 Blue (Novagen) and used as a template for in vitro mutagenesis. Oligonucleotide primers used for mutagenesis are described in (Table 6.1). Mutants were screened by sequencing. Selected plasmid D N A was then digested with EcoRl/Ncol  and the fragment containing the mutation was cloned  into similarly digested pK2/M5. pK2/M5 was used as the template for the production of Pro67Gly, Pro67Ala, Pro259Gly, Pro259Ala, Pro262Gly, Pro262Ala (Table 6.2). Following mutagenesis, plasmid D N A was digested with either EcoRI/Bglll  175  (in the case  TABLE 6.1. Oligonucleotide primers used for constructing various C N V Mutant  CP mutants.  Sequence  Position  1  2  Pro63Gly  T G T C C G T G G G C T A T G A G A G C (-)  (2818-2719)  Pro63Ala  T G T G C G T G G G C T A T G A G A G C (-)  (2818-2719)  G G C T T T T C C T G G G G C T A T C G C C C G (+)  (2819-2842)  C C A C C A A A A G C C T G T G G G T G G G C (-) C C A G C A A A A G C C T G T G G G T G G G C (-) G G C T A T C G C C G C G C C A A T C (+)  (2830-2808)  Pro85Ala  C T A G C T T T C C T T C C T T T A A C C G C A T A G (-) G T T T C A A A C A G C A A A A G G A T C T G T G (+)  (2883-2855) (2884-2909)  Pro259Gly  G T T C C T T G T G C C T C A A A C A G A T C (-)  Pro259Ala  G T T G C T T G T G C C T C A A A C A G A T C (-)  Oligo#190  G T C G C C T C T T C T G G A A T C G (+)  (3406-3383) (3406-3383) (3407-3425)  Pro262Gly Pro262Ala 01igo#193  A G A C C C G A C G T T G G T T G T G C (-) A G A G C C G A C G T T G G T T G T G C (-) T C T G G A A T C G T T G T T C C G G G A G A G C (+)  (3415-3395) (3415-3395) (3416-3437)  Thr292Ala  A A G G T C G G C A G C C G A A G C C A C C T C (-) G T A T G G C A G G C A C G T G T G C (+)  (3509-3486) (3510-3528)  Trp296Ala  G T A G C G C A G G C A C G T G T G C C C G (+)  (3510-3531)  Gln297Ala  G T A T G G G C G G C A C G T G T G C C C G G C (+)  (3510-3533)  A A G G T C G G T A G C C G A A G C C A C (-)  (3509-3489)  A G A G G C T C C G G C G G T G C T C (-) A G A G A A T C C G G C G G T G C T C A C A C (-) A G A G G T T C C G G C G G T G C T C (-) A C C G C T T C C G G C G G T G C T C A C A C (-) G C A T A T G T T G C A A A C A T C A C T A T A C (+)  (3643-3618) (3643-3622 (3643-3618) (3643-3622)  A C T C G C G T T G G C A T T C A C A C G T A T A G (-) C T T T C T G G G C T C A C G G G A G C G (+)  (3689-3664) (3690-3710)  01igo#178 Pro67Gly Pro67Ala 01igo#121  3  3  3  3  01igo#128 Ser338Ala Ser338Phe Ser338Thr Ser339Gly 01igo#112  4  5  Leu353Ala  (2830-2808) (2831-2849)  (3644-3668)  Sequences of primers are shown 5' to 3'. Underlined nucleotides correspond to mutated positions, and the antisense (-) or sense (+) orientation relative to the C N V genome. 2 Nucleotide positions are relative to the complete C N V genome (5). 3  Oligos 178, 121, 190 and 193 were used as (+) sense primers for Pro63, Pro67, Pro259 and Pro262 mutants, respectively. 4  01igo#128 was used as (-) sense primer for Trp296 and Gln297 mutants. 5  01igo#l 12 was used as (+) sense primer for Ser338 and Ser339 mutants.  176  of Pro67Gly and Pro67Ala) or EcoKMNcol (in the case of Pro259Gly, Pro259Ala, Pro262Gly and Pro262Ala) and the mutated fragment was cloned back into pK2/M5 to obtain a full-length clone containing the mutation.  6.2.4 In vitro transcription and inoculation of plants Preparation of T7 polymerase run-off transcripts and inoculation of plants were done as described (6).  6.2.5 Fungus transmission assay Partially purified virions were tested for their ability to be transmitted by O. bornovanus zoospores as described previously (3; Chapter 4).  6.3 Results and conclusions Mutations introduced into the C N V CP are listed in Table 6.1. The location of mutations in the three-dimensional structure of the CP subunit is shown in Figure 6.1. A l l the mutated amino acids in the P domain and hinge are predicted to be exposed on the surface of the particle except ser338 which is buried in the P/P domain contact region. Pro63, Pro76, Pro85 are present on the internally located arm domain. Transcripts corresponding to each of the mutants (Table 6.2) were inoculated onto N. benthamiana to assess their ability to produce virions in plants. All the mutants were found to be capable of producing symptoms in plants. Agarose gel electrophoresis of partially purified virus preparations (data not shown) of all mutants showed that virus particles could be purified from all mutants, except Asp293Ala, Trp296Ala, Ser338Phe, Ser339Gly and Leu353Ala  177  120  100-  o • ,-H CO CO  5/5  5/5  4/4  4/4  5/5  5/5  o  <  5/5  5/5  5/5  5/5  <  oo  O  14/15  34/39  80-  60-  CO  40-  202/20 J/25_  0-  >  o o  < o  O ro OH  ro 1-1  OH  o 00  o  <!  ON  ON  m  in (N O  O  PH  S-l  PH  ON CM O  O  H  a  O  CO  J-l  00  C/3  Mutant Figure 6.2. Summary of transmission efficiencies of C N V mutants. Transmission assays were conducted using 1 pg of virus in 10 ml of zoospores (lxlO zoospores/ml) as described in reference #3 and Chapter 4. The percentage of pots showing transmission is indicated on the X-axis. The numbers on the bars indicate the number of pots showing transmission versus the number of pots tested. The data represented for C N V , Pro63Gly, Pro259Gly and Pro259Ala is a compilation of at least three experiments. For the rest of the mutants, the data presented are from one experiment. 5  178  Table 6.2. Properties of C N V mutants  Mutation  Location  Particle formation  1  Fungus transmission  Pro63Gly Pro63Ala Pro67Gly Pro67Ala Pro85Ala  arm arm arm arm arm  + + + + +  + + + +  Pro259Gly Pro259Ala Pro262Gly Pro262Ala  S S hinge hinge  + + + +  + + +  Thr292Ala Trp296Ala Gln297Ala Ser338Ala Ser338Phe Ser338Thr Ser339Gly Leu353Ala  +  P P P P P P P P  -  + + -  + -  _  + Not tested  + + Not tested Not tested  + Not tested  Abbreviations used: S, shell domain; P, protruding domain 'Virus particle formation was determined by agarose gel electrophoresis following "mini-prep" purification of virus from inoculated leaves  179  (Table 6.2). The yield of particles appeared to be approximately equal to that of WT CNV. Transmission assays were conducted for all the mutants that produced virus particles to assess their transmission efficiency relative to WT CNV. Transmission efficiency was measured by determining the number of pots infected versus the number of pots inoculated. Figure 6.2 shows that Pro63Gly transmitted with very low efficiency (10%), whereas Pro259Gly was not transmitted by zoospores. WT C N V and all other mutants were transmitted with higher efficiency (92% to 100%). These results show that the majority of the mutants (Table 6.2) are capable of being transmitted by O. bornovanus. In the case of Ser338 it was previously shown that a mutation to glycine (LLK63) allowed for particle formation but interfered with transmission (2; Chapter 2). Here it is shown that changing Ser338 to Ala did not significantly affected particle formation or transmissibility. However, mutation to Thr or Phe prevented particle formation. Ser338 is located in a region wherein P/P domain contact occurs and it is therefore likely that the substitution with the bulky aromatic Phe or the larger Thr prevents particle formation by interfering with P/P domain interaction. Mutation of Pro259 and Pro63 to Ala or Gly did not interfere significantly with particle formation (Table 6.2). However, transmission was affected when Pro259 was mutated to Gly. It is possible that the greater conformational freedom engendered by Gly substitution does not support the proper conformational changes required for successful transmission as previously suggested for Pro73Gly (3; Chapter 4). Further infectivity studies are required to assess the fitness of the Pro259 and Pro63 mutants and experiments need to be designed to assess potential changes in conformation induced by these substitutions. Future work could involve testing whether these mutants : 1) bind to  180  zoospores efficiently using the in vitro binding assay; and 2) whether the mutants fail to undergo conformational changes upon zoospore binding as the possible reason(s) for reduced transmission. As far the other remaining transmissible mutants, further mutagenesis experiments are required to assess the importance of these particular amino acids in fungus transmission.  181  6.4 References 1. Campbell, R. N., S. T. Sim, and H. Lecoq. 1995. Virus transmission by host-specific strains of Olpidium bornovanus and Olpidium brassicae. Eur. J. Plant Pathol. 101:273282. 2. Kakani, K., J. Y. Sgro, and D. Rochon. 2001. Identification of cucumber necrosis virus coat protein amino acids affecting fungus transmission and zoospore attachment. J. Virol. 75:5576-5583. 3. Kakani, K., R. Reade, and D. Rochon. 2004. Evidence that vector transmission of a plant virus requires conformational change in virus particles. J. Mol. Biol. 338:507-517. 4. Robbins, M . A., R. D. Reade, and D. M . Rochon. 1997. A cucumber necrosis virus variant deficient in fungal transmissibility contains an altered coat protein shell domain. Virology 234:138-146. 5. Rochon, D., and J. H. Tremaine. 1989. Complete nucleotide sequence of the cucumber necrosis virus genome. Virology 169:251-259. 6. Rochon, D. M., and J. C. Johnston. 1991. Infectious transcripts from cloned cucumber necrosis virus cDNA: Evidence for a bifunctional subgenomic mRNA. Virology 181:656-665.  182  Co-authorship Statement Mr. Kakani played a major role in the design and execution of experiments in Chapter 2, 3, 4 and 6 which form the bulk of the research described in this thesis. He also contributed substantially to data analyses and manuscript preparation for each Chapter. The contributions of co-authors are listed below:  1. Dr. Jean-Yves Sgro (Chapter 2) conducted the homology modelling and subsequent three-dimensional structural representation ofthe C N V particle described in Section 2.2.8.  2. Dr. Marjorie Robbins designed, conducted and analyzed the experiments described in Fig. 3.1 and Fig. 3.6 in Chapter 3.  3. Ron Reade assisted in the development of procedures to analyze structural changes in C N V virions as a result of fungal zoospore attachment thereby contributing, in part, to Figure 4.2, excluding the parts of figures corresponding to Pro73Gly. Ron Reade also assisted in the determination of the location of trypsin cleavage sites in swollen C N V particles (Section 4.3.6).  4. D'Ann Rochon was the supervising scientist. She developed the broad goals and assisted in the design and interpretation of experiments as well as manuscript preparation.  D'Ann Rochon :  .-.^.. . r  (Supervisor and senior author)  Naga Kishore Kakani: ...  

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