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Sunn-hemp mosiac virus as a helper in the intercellular spread of southern bean mosaic in a resistant… Fuentes, Ana Lucía 1992

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SUNN-HEMP MOSAIC VIRUS AS A HELPER IN THE INTERCELLULARSPREAD OF SOUTHERN BEAN MOSAIC VIRUS IN A RESISTANT HOSTbyAna Lucia FuentesB.Sc., University of Costa Rica, 1981M.Sc., University of Costa Rica, 1986A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIESDEPARTMENT OF PLANT SCIENCEWe accept this thesis as conformingo the required standardTHE UNIVERSITY OF BRITISH COLUMBIASeptember 1992© Ana Lucia Fuentes, 1992Signature(s) removed to protect privacySignature(s) removed to protect privacyIn presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)Department of ‘ :\tThe University of British ColumbiaVancouver, CanadaDate Strt 2’DE-6 (2/88)Signature(s) removed to protect privacyAbstractFour bean cultivars (“Bountiful”, “Pinto”, “Top Crop”, and“Tendergreen”) allowed only subliminal replication of the cowpea strain ofsouthern bean mosaic virus (SBMV-C). Bean protoplasts, on the other hand,sustained replication of SBMV-C upon in vitro inoculation. Antigen accumulationof the bean strain of southern bean mosaic virus (SBMV-B) and SBMV-C in beanprotoplasts was similar, indicating that the replicating capacity of both virusesdoes not differ in bean cells.When the four bean cultivars were inoculated with a mixture of sunnhemp mosaic virus (SH1VIV), a tobamovirus, and SBMV-C, the latter was readilydetected in the inoculated primary leaves. The rate of spread of SBMV-C in thepresence of SHMV was compared to the rate of spread of SBMV-B in bean coinoculated with SHMV. Virus accumulation in leaf blades, lateral veins, mid-ribs,petioles, stems and roots was similar for both strains in the non-vascular tissue ofthe inoculated leaf; a sharp decline in SBMV-C accumulation was observedstarting from the lateral veins towards the mid and distal parts of the petiole,where virtually no virus could be found. These results contrasted with theuniform presence of SBMV-B throughout infected bean plants.Leaf strips blotted on nitrocellulose paper and developed as forWestern blotting confirmed these results, with SBMV-C antigen being detected inmesophyll tissue and in epidermal cells of the lateral veins of the inoculated11primary leaves. Electron micrographs of immunogold-labelled sections revealedthe absence of uniform SBMV-C particles in the mesophyll cells; instead, heavilylabelled, amorphous protein clumps in the vacuole were found. SBMV-C coatprotein from infected cowpea and bean plants showed no difference in its mobilityduring electrophoresis in denaturing polyacrylamide gels.These results indicate that SHMV facilitates cell to cell spread ofSBMV-C in inoculated bean leaves but does not allow for the movement of thelatter through the vascular system. Lack of efficient assembly of SBMV-C doesnot impede cell-to-cell movement of the virus in the doubly-infected leaves, yet itis probably an important factor involved in determining the inability of SBMV-Cto move into andlor through the vascular system.111Table of ContentsAbstract . . . . . . . . iiList of’l’ables . • • viiilast of Figures . . . . . . . . . . . . . ixList of Abbreviations . . . . . . . . . . . . .. . . . xiiA.ckiiocvledgenients. . . . . . . xvi1. Introduction 11.1 General Introduction 11.2 Objectives 152. Biological characterization of the interactionbetweenSBMV-CandSHMV 172.1 Introduction 172.2 Materials and methods 182.2.1 Virus propagation and purification 182.2.2 Virus inoculation 202.2.3 Collection and storage of samples 202.2.4 Bioassay 212.2.5 Production of antibodies and detectionof antigen by ELISA 212.2.6 Temperature study 232.2.7 Multiple passage study 23iv2.3 Results . 242.3.1 Symptoms in singly and doubly inoculatedplants 242.3.2 SBMV-C accumulation in singly anddoubly inoculated plants 302.3.3 Effect of sequential inoculation 352.3.4 Effect of temperature shifts 362.3.5 Effect of multiple passage 362.3.6 Effect of other viruses that replicate in bean on thereplication of SBMV-C 378. Replication of viruses in singly and doubly inoculated beanprotoplasts 893.1 Introduction 393.2 Materials and methods 393.2.1 Protoplast isolation 403.2.2 Protoplast inoculation and incubation 403.2.3 Assays for virus in protoplasts 413.2.4 Protoplast fixation andfluorescent antibody staining 413.3 Results 423.3.1 Proportion of SBMV-C-infected cells in singly and doublyinfected leaves 42v3.3.2 Replication of SBMV-C in singly inoculatedprotoplasts 433.3.3 Replication of SBMV-C in doubly inoculatedprotoplasts in comparison to SBMV-Binoculated protoplasts 454. Distribution of SBMV-C within the inoculated leaves and throughoutthe plant 474.1 Introduction 474.2 Materials and methods 474.2.1 Collection of samples 474.2.2 ELISA and bioassay 484.2.3 Dot-blot hybridization assayfor viral RNA 484.2.4 Tissue-print immunoblotting 514.2.5 Cryosectioning and immunofluorescentlabelling of thick sections 534.2.6 Immunosorbent electron microscopy 544.2.7 Tissue fixation and embedding forelectron microscopy 554.2.8 Inimunogold labelling of antigens 584.2.9 Electrophoresis and Western blotting of viral proteins 594.3 Results 60vi4.3.1 Distribution of SBMV-C capsid protein compared to thatof SBMV-B in double infections with SHMV 604.3.2 Distribution of SBMV-C RNA in doublyinoculated plants 644.3.3 Viral antigen distribution in thick sections ofleaf and petiole by tissue blot analysis 644.3.4 SBMV-C antigen distribution in thick sections bycryosectioning and inimunofluorescent labelling . . . . 724.3.5 Viral particle distribution in thin sections of leaf andpetiole by standard osmium tetroxide fixation 834.3.6 Viral particle distribution in thinsections of leaves by immunogold labelling 904.3.7 Electron microscopic observations after trapping ofSBMV-C from doubly-infected leaves 1104.3.7 Electrophoresis and Western blot of viralproteins 1105. Eiscussion 1156. Bibliography 127viiList of TablesTable I. Effect on the replication of SBMV-C by other virusesthat infect bean systemically 38Table II. Replication of SBMV-C and SBMV-B and replication of SBMV-Cin single and mixed infections with SHMV in “Bountiful’ beanprotoplasts detected by double antibody sandwichenzyme-linked immunosorbent assay (ELISA) 46viiiList of FiguresFig. 1 “Pinto” bean singly inoculated with SBMV-C 25Fig. 2 Symptoms, during the months of June, July and August,in “Bountiful” primary leaves 27Fig. 3 “Bountiful” plant, doubly-inoculated with SHMV and SBMV-C 29Fig. 4 Symptoms in “Pinto” primary leaf inoculated with A) SHMV andSBMV-C and B) SBMV-B 31Fig. 5 Detection of SBMV-C by bioassay on cowpea “Georgia 21” inprimary leaves of different bean cultivars doubly-inoculatedwith SHMV and SBMV-C 33Fig. 6 Detection of SBMV-C antigen, by ELISA, in primary leavesof four different bean cultivars doubly-inoculated withSBMV-C and SHMV 34Fig. 7 Time course of the replication of SBMV-C in “Bountiful”protoplasts detected by ELISA and bioassay on Vignaunguiculata L. var 399414 44Fig. 8 Accumulation of SBMV-C and SBMV-B capsid proteins, detectedby ELISA in bean, in mixed infections with SHMV 62Fig. 9 Detection of the SBMV-C RNA in different parts of bean plantsby dot-blot hybridization using a nick-translated 32P labelledc-DNA probe of SBMV-C 65ixFig. 10 Transfer of proteins from leaf sections and tissue pressedonto nitrocellulose membranes 67Fig. 11 SBMV-C antigen detected in SBMV-C-infected cowpea primaryleaf blade and vein tissue by tissue print immunoblotting 68Fig. 12 SBMV-C antigen detected in SBMV-C-infected cowpea primaryleaf blade and vein tissue by tissue print immunoblotting 70Fig. 13 SBMV-C antigen detected in “Bountiful” bean primary leaftissue, doubly-inoculated with SB1VW-C and SHMV, bytissue print immunoblotting 73Fig. 14 Tissue prints of healthy “Bountiful primary leaf bean tissue 75Fig. 15 Inimunofluorescent labelling of a thick section of a primary“Bountiful” leaf doubly-inoculated with SBMV-B and SHMV 77Fig. 16 Immunofluorescent labelling of a thick section of “Bountiful”bean primary leaf mock-inoculated with phosphate buffer 79Fig. 17 Immunofluorescent labelling of a thick section of “Bountiful”bean primary leaf mock-inoculated with phosphate buffer 81Fig. 18 Thin section of Epon-embedded “Bountiful” primary leaf tissuedoubly-inoculated with SHMV and SBMV-B 84Fig. 19 Thin section of Epon-embedded “Bountiful” primary leaf tissuedoubly-inoculated with SHMV and SBMV-C 86Fig. 20 Thin section of Epon-embedded “Bountiful” primary leaf tissue 88Fig. 21 Thin sections of RNase-treated “Bountiful” primary leaf tissuexdoubly-inoculated with SHMV and SBMV-B.91Fig. 22 Thin section of RNase-treated “Bountiful” primary leaf tissuedoubly-inoculated with SBMV-C and SHMV 93Fig. 23 RNase-treated “Bountiful” primary leaf tissue mock-inoculatedwith phosphate buffer 95Fig. 24 Specific gold-labelling of SBMV in leaf-dips of homogenatesof infected tissue 97Fig. 25 Gold-labelling of SBMV-B antigen in thin sections of “Bountiful”primary leaf tissue doubly-inoculated with SBMV-B and SHMV ... 100Fig. 26 Gold-labelling of SBMV-C antigen in thin sections of cowpeaprimary leaf tissue doubly-inoculated with SBMV-C and SHMV 102Fig. 27 Thin section of “Bountiful” primary leaf doubly-inoculatedwith SBMV-C and SHMV 104Fig. 28 Gold-labelling of thin section of “Bountiful” primaryleaf doubly-inoculated with SBMV-C and SHMV 106Fig. 29 Gold-labelling of thin sections of “Bountiful” primaryleaves doubly-inoculated with SBMV-C and SHMV 108Fig. 30 SBMV-C-specific trapping of antigen from homogenatesof infected tissue 111Fig. 31 Western blot analysis of SBMV-C coat protein migrationfrom cowpea and bean primary inoculated leaves 113xiList of AbbreviationsA2 - absorbance at 260 nmA280 - absorbance at 280 nmA405 - absorbance at 405 nmA1MV - alfalfa mosaic virusATP - adenosine triphosphateBCIP - 5-bromo-4-chloroindoxyl phosphateBGMV - bean golden mosaic virusBRMV - bean rugose mosaic virusBSA - bovine serum albuminBSMV - barley stripe mosaic virusBYMV - bean yellow mosaic virusC - CelsiusCaC12 - calcium chloridecAMP - cyclic adenosine mono phosphatecDNA - complementary DNAcm (mm, nm, urn)- centimeter (millimeter, nanometer, micrometer)CPMV- cowpea mosaic virusDAS-ELISA - double antibody sandwich enzyme-linked immunosorbent assayDDSA - dodecenyl succinic anhydrideDIECA - sodium diethyldithio-carbamatexiiDMP - 2,4,6-tn dimethyl-aurinomethyl phenolDNA - deoxyribonucleic acidds - double-strandedEDTA - ethylenediaminetetraacetic acidELISA - enzyme-linked immunosorbent assayEM - electron microscopyFITC - fluorescein iso thio cyanateg (mg, ug) - gram (milligram, microgram)g - gravityh-hourHAuCL4 - hydrochloroauric acidHC1 - hydrochloric acidH20 - waterIgG - immunoglobulin GISEM - immunosorbent electron microscopyK - potassiumkDa (Da) - kilodalton (dalton)1 (ml, ul) - liter (milliliter, microliter)LiCl - lithium chlorideM (mM, uM) - molar (millimolar, micromolar)MgCl2 - magnesium chloridemm - minutexliiN - nitrogenNaAc - sodium acetateNaC1 - sodium chlorideNaOH - sodium hydroxideNBT - nitroblue tetrazoliumNH4C1 - ammonium chlorideNMA - nadic methyl anhydrideORF - open reading frame0804 - osmium tetroxideP - phosphorusPBS - phosphate-buffered saline (127 mM NaC1, 2.6 mM KC1, 8.5 mM Na2HPO4,1.1 mM KH2PO4)PEG - polyethylene glycolPLRV - potato leaf roll virusPVP - polyvinylpyrrolidonePVX- potato virus XPVY- potato virus YRCMV - red clover mottle viruss - secondRNA - ribonucleic acidSBMV - southern bean mosaic virusSBMV-B - bean strain of southern bean mosaic virusxivSBMV-C - cowpea strain of southern bean mosaic virusSDS - sodium dodecyl sulfateSHMV - sunn-hemp mosaic virusss - single-strandedSSC- a buffer containing 0.15 M sodium chloride and 0.015 M trisodiumcitrateTE- a buffer containing 10 mM Tris-HC1 pH 7.5 and 1 mM EDTATMV- tobacco mosaic virusTMV-L - TMV strain which induces hypersensitive reactionsTNE- a buffer containing 10 mM Tris-HCL pH 7.5, 10 mM NaCT and 1 mM EDTATomRsV - tomato ringspot virusTRV - tobacco rattle virusUA - uranyl acetateUV - ultravioletv - volumevRNP - virus-specific ribonucleoproteinxvAcknowledgementsI wish to thank my supervisor, Dr. R.I. Hamilton for his guidance andpatience during the course of this work. I also wish to express my appreciationand admiration for him as a scientist who not only cares about, but also workstowards, solving plant pathology problems in the developing countries.I thank the members of my supervisory committee, Dr. R.J. Copeman, Dr.C.J. Douglas, Dr. D.M. Rochon, Dr. V.C. Runeckles and Dr. M. Weintraub, fortheir help and valuable advice throughout my work on this project.I am indebted also to Dr. M. Weintraub, former Director, VancouverResearch Station, Dr. D.L. Struble, Director, and the Research Station staff forproviding the facilities for this work. Special thanks to Dr. Fran Leggett for herhelp, excellent advice and patience in the electron microscopy lab.I thank M. Weis and Dr. L. Samuels for their assistance in the electronmicroscopy lab at the University of British Columbia.I thank my husband John; my children Irene, Lorena and Alexander and mymother for their understanding and support during the writing of this thesis.xviChapter 1Introduction1.1. General IntroductionWhen a plant is inoculated with a virus, its ability to establish aninfection depends upon two processes: 1) the capacity of its genome to replicate inthe cells of the inoculated plant and 2) the movement of the virus from cell to celland to other parts of the plant. The capacity of a virus to perform these functionsin a particular plant species depends on the genome of both plant and virus(Atabekov and Dorokhov, 1984; Zaitlin and Hull, 1987). Thus, both virus andhost have an active role in determining the course of the infection process.Responses by plants to inoculation with a virus have been categorizedby Matthews (1991) as immune and infectible. In an immune plant, the viruscannot replicate in protoplasts or in cells of the intact plant and no virus progenyare produced. An infectible plant is one in which the virus can infect andreplicate in protoplasts. Several kinds of infectible plants or hosts have beendefined: 1) resistant: virus multiplication is limited to initially infected cells,resulting in a subliminal infection; 2) hypersensitive: infection is limited to anarea of cells around the initially infected cells, usually with the formation of1visible necrotic local lesions; and 3) susceptible: virus replicates and movessystemically. The host can be sensitive, i.e., exhibit more or less severe symptomsof disease, or tolerant, when there is little or no apparent effect on the plant(Matthews, 1991).The basis for the different responses of plants to viruses is not yetfully understood. There are few examples in which plants are a source ofresistant protoplasts (immune plants). Beier et al. (1977, 1979) found that out of50 lines of cowpea that were operationally immune to cowpea mosaic virus(CPMV), “Arlington” was the only one that was a source of resistant protoplasts.The resistance of “Arlington” protoplasts is thought to be mediated by an inhibitorof the proteolytic processing of CPMV polyproteins (Ponz and Bruening, 1986).This kind of extreme resistance seems to be an exception, most plant cells beingcapable of replicating essentially any plant virus (van Loon, 1983).Three types of response have been defined above for infectible hosts.In the case of a subliminal infection, virus replication in the initially infected cellsis not blocked, yet the virus cannot move beyond these cells. This type of responsecould be due to the lack of a functional transport factor that would allow the virusto move away from the initially infected cells, andlor to an antiviral defencereaction operating in the plant tissue but not in protoplasts or individual cells(Atabekov and Dorokhov, 1984). Limitation of viral spread also occurs in the2hypersensitive host reaction, which is usually accompanied by the developmentof a necrotic lesion, localized at the site of virus infection.The hypersensitive host reaction is a generalized response to differentpathogens and is thought to depend on the interaction between a host resistancegene and a pathogen avirulence gene (Flor, 1971). Most of the genetic studies onviral-induced hypersensitivity have been done with tobacco mosaic virus (TMV)interacting with plants in the genus Nicotiana. The N’ gene in N. sylvestris hasbeen identified as the gene controlling the hypersensitive response against mosttobamoviruses, except the common strain (reviewed by van Loon, 1987; Culver etal., 1991). Coat protein substitutions and alterations in nucleotide sequence in thecoat-protein open reading frame of the hypersensitive reaction-inducing TMV-Lstrain, demonstrated that coat protein gene sequences are involved in theinduction of the N’ gene (Saito et al., 1987; Knorr and Dawson, 1988). However,the N gene of N. glutinosa, which controls the hypersensitive reaction againstalmost all strains of TMV, is not induced by the coat protein (Dawson et al., 1988),while the Tm-2 and TM-22hypersensitive resistance factors in tomato seem to betriggered by the TMV 30 kDa movement protein (Meshi et al., 1989).Recognition of infecting viruses by plants does not involve a singleviral component, as demonstrated by the results of studies on induction of thehypersensitive response. Thus, plants seem to have evolved to recognize any3viral-specific product presented to them (Culver et al., 1991). Viral genomes, onthe other hand, are completely dependent on the host’s replicating machinery forsuccessful replication and must specifically interact with host macromolecules tomove into other cells. A direct implication of this phenomenon is that evolutionof viral genomes is closely tied to the evolution of the genome of the host plant.Most of the recent molecular studies undertaken to understand theinteractions between plant viruses and their hosts have focused on the viruses,given the relative simplicity of their genomes. After infection and replication inthe initially infected cells, efficient establishment of a viral infection in a plantdepends on the virus spreading systemically. Viral factors responsible fortranslocation have been identified for several viruses and some models on theinteraction of these factors with host factors have arisen.Plant viruses move in their hosts by slow, cell-to-cell transport withinthe parenchyma cells (i.e., short distance movement) and by rapid spread throughthe conducting system (i.e., long distance movement) (reviewed by Atabekov andDorokhov, 1984; Zaitlin and Hull, 1987; Hull, 1989). Cell-to-cell spread is believedto occur through plasmodesmata, which are plasma membrane-lined channels thattraverse plant cell walls, thus connecting adjacent cells (reviewed by Robards andLucas, 1990). Long-distance movement, on the other hand, occurs in either thephloem or xylem (reviewed by Matthews, 1991).4In some cases, whole virions have been seen in the plasmodesmata(Esau et al., 1967; Allison and Shalla, 1974; Weintraub et aL, 1976; ), yet properencapsidation does not seem to be a requirement for cell-to-cell movement of allplant viruses. Examples of cell-to-cell movement ofviruses after deletions or pointmutations in the capsid protein are tobacco mosaic virus (TMV) (Dorokhov et al,1983, Takamatsu et aL, 1987, Dawson et al, 1988), tomato golden mosaic virus(Garcliner et al., 1988), turnip crinkle virus (Hacker et al., 1992) and cucumbernecrosis virus (M.A. McLean, personal communication).Thus, there is no uniform evidence to suggest that all viruses moveacross plasmodesmata as one entity, be it virus particles, virus nucleic acid orvirus-specific ribonucleoprotein (vRNP). The same is true for long distancemovement through the conducting elements. In fact, with some viruses it appearsthat local and long distance spread have different requirements for coat protein.Using mutants with modifications in the coat protein gene or in the origin ofassembly on the genomic RNA of TMV, Saito et al. (1990) showed that both coatprotein, with a capacity to assemble into virus particles, and the assembly originin the genomic RNA were involved in efficient long-distance movement. However,coat protein is entirely dispensable for systemic movement of some viruses, forexample, barley stripe mosaic virus (BSMV) (Petty and Jackson, 1990), andtomato golden mosaic virus (Gardiner et al, 1988).5These differences in the requirements for coat proteins could indicatedifferences in the mechanisms and principles controlling viral movement, not onlyin terms of cell-to-cell and long distance movement, but also among differentviruses. In spite of this, understanding of the mechanisms involved in viral cell-to-cell movement was greatly increased after it was established that a specificvirus-encoded protein, the 30-KDa product of TMV, was required for cell-to-celltransport of that virus (Nishiguchi et al., 1978; Leonard and Zaitlin, 1982; Meshiand Okada, 1987; Deom et al., 1987).Several studies, in which the migration of fluorescent molecules ofdefined sizes between cells was recorded, showed that plasmodesmata have amolecular size exclusion limit or “gating” capacity of approximately 800 Da(Madore et al., 1986, Wolf et at, 1989). Since mature virions and viral genomesare considerably greater in size than the molecular exclusion limit ofplasmodesmata (Gibbs, 1976), it is generally accepted that viruses must modiVyplasmodesmata to migrate into adjacent cells. The 30-KDa TMV movementprotein has been localized within plasmodesmatal connections by immunogoldcytochemical studies (Tomenius et al., 1987). It has also been shown to modify thesize-exclusion limits of plasmodesmata in leaf tissue of transgenic tobacco plantstransformed with the 30-KDa movement protein gene and expressing the geneproduct (Wolf et aT., 1989; Deom et al., 1990,1991). In such plants,plasmodesmata allowed the movement of fluorescein isothiocyanate-labelled6dextrans with average molecular masses of 9.4 KDa (approximately ten times aslarge as those which moved between control cells) (Wolf et al., 1989).Recently, the movement protein of TMV was identified as a singlestranded (ss) nucleic acid-binding protein (Citovsky et al., 1990). Based on thisproperty, the TMV movement protein-RNA complex has been suggested as anintermediate in cell-to-cell movement (Citovsky and Zambryski, 1991). Accordingto this model, only RNA molecules bound to the movement protein can betransported through the plasmodesmata. The movement protein would have twofunctions; it would act on the plasmodesmata and increase their permeability, andit would “unfold” the RNA and shape it into a transferable form (Citovsky et al.,1990). The nature of the interaction ofthe movement protein-nucleic acid complexwith the plasmodesmatal subunits is unknown, but by analogy with gap junctionsin animals, could involve cAMP-dependent phosphorylation ofthe plasmodesmatalsubunits. This would be supported by the finding that cell-to-cell spread of TMVis dependent on intracellular levels of cAMP (Atabekov and Taliansky, 1990).After modification of the plasmodesmatal channels, the entire movement proteinnucleic acid complex or the nucleic acid alone would be translocated across theplasmodesmatal channel (Citovsky and Zambryski, 1991).Regardless of the mechanism underlying cell-to-cell movement, thereis evidence that in many cases the capacity of a given virus to be transported in7a plant determines whether that plant is a potential host for the virus (Talianskyet al., 1982a). Thus, although a virus may be able to replicate in the initiallyinfected cells of a plant, if it cannot be transported beyond these, the plant can beregarded as resistant to that virus (Matthews, 1991). In certain cases, resistanceof a plant to a virus can be overcome by co-infecting that plant with another virus.One virus, the “helper”, normally replicates and spreads in the host wheninoculated alone; the other, the “dependent” virus, may replicate in the inoculatedcells but does not spread unless the helper virus is present.Complementation ofvirus spread has been described for both relatedand unrelated viruses (for review, see Atabekov and Taliansky, 1990). Forexample, members of the tobamovirus group in different combinations and indifferent plants complemented one another’s systemic spread (Malyshenko et al.,1989). Spread of phloem-limited viruses, such as potato leaf roll virus (PLRV), aluteovirus, into mesophyll cells, was brought about by mixed infection with thepotato virus X potexvirus (PVX) (Atabekov et al., 1984) or the potato virus Ypotyvirus (PVY) (Barker, 1987). This phenomenon is not surprising if mostviruses code for movement proteins which operate in a similar fashion to alterplasmodesmata and facilitate viral movement. However, complementation doesnot occur with all viral combinations, nor does it occur for the same viruscombinations in all hosts. For example, complementation of PLRV movement byPVY is effective in N. clevelandii but not in potato; no complementation was8observed between tobacco rattle virus (TRV) and barley stripe mosaic virus(BSMV) in tobacco or BSMV and alfalfa mosaic virus (ALMV) in wheat(Malyshenko et al., 1989).Thus far there is no clear functional or structural classification ofmovement proteins that explains and predicts transport functioncomplementation. Several mechanisms to explain the occurrence ofcomplementation, which are not mutually exclusive, include: 1) non-homologousmovement proteins coded by unrelated viruses which function in a similar fashion,yet the degree to which the modifications of the plasmodesmata can be utilizeddepends upon the timing and location of the “dependent” virus in relation to thereplication and spread of the “helper” (Maule, 1991); 2) since complementationseems to be rather host-specific, it is possible that the host genotype can influenceand modify the phenomenon of complementation of transport function (Barker,1987; Atabekov and Taliansky, 1990); and 3) movement proteins of differentviruses (or of taxonomic groups of viruses), use different mechanisms, involvingother viral products and/or plant factors, for the expression of the transportfunction (Atabekov and Taliansky, 1990). In the third model, in order forcomplementation to occur, “dependent” virus-specific products should becompatible with the plant and/or to the “helper” movement protein.One mechanism for transport which is compatible with the first9possibility is that proposed by Citovsky and Zambrysky (1991). Movementproteins bind ss-nucleic acids in a non-specific manner (Citovsky et al., 1990,1991). The occurrence of complementation between two viruses would depend onwhether the RNA of the “dependent” virus is in sufficient quantity and in closeproximity to the movement protein of the “helper”. If these two conditions aremet, the movement protein could then bind the “dependent” RNA and effect itstransport, thus allowing it to move beyond the initially infected cells. If the“dependent” viral RNA does not meet these requirements, however, the movementprotein would not bind to the RNA and complementation would not occur.The second model is supported by the fact that complementation ofmovement between certain viruses occurs in some hosts and not others. Anexample, mentioned above, is the helper effect induced by PVY that effects PLRVmovement into non-phloem tissue in N. clevelandii but not in potato (Barker,1987).Finally, the third model is supported by recent findings on therequirements for complementation of red clover mottle virus (RCMV) by TMV(Taliansky et al, 1992). Previously it was shown that tobamoviruses enabled thetransport of RCMV in tobacco plants normally resistant to RCMV (Malyshenko etal., 1988). However, RCMV transport does not take place in transgenic tobaccoplants that produce the movement protein of TMV, whereas the transport of TMV10Lsl mutant (a temperature sensitive mutant in cell-to-cell movement) iscomplemented in these plants. Nevertheless, complementation ofRCMV transportdoes occur when the transgenic plants are infected with both RCMV and TMV Lslat the non-permissive temperature (33 C) (Taliansky et al., 1992). From theseresults, the authors suggest that the presence of the full-length TMV genome ora certain TMV-encoded product(s) in addition to the 30-KDa movement protein isnecessary for complementation of the RCMV transport function.Understanding how plant and viral genes control viral movement isnecessary if control of such movement could effect management of virus disease.One approach , taken in this study, is to examine the course of events which occurin a mixed infection in which one virus acts as a “helper” and another as a“dependent”.In previous studies, Molefe et al (1983), showed that bean (Phaseolusvulgaris L. “Pinto”), singly inoculated with SBMV-C sustained only subliminalviral replication in inoculated primary leaves; no symptoms developed and verylow amounts of virus were recovered. However, plants co-infected with SHMV,which infects bean systemically, showed local lesions in the inoculated primaryleaves due to the replication of SBMV-C. Furthermore, SBMV-C replicated tolevels detectable by ELISA and bioassay in the inoculated primary leaves, but notin the trifoliate leaves, of three other bean cultivars co-infected with SHMV11(Fuentes and Hamilton, 1988, 1991).The ‘helper’ virus, SHMV, is a member of the tobamovirus group andit occurs naturally in leguminous plants in several continents (Kassanis andVarma, 1975; Varma, 1985). The virus is rod-shaped; both full length (300 nmlong and 17 nm wide), and shorter particles are produced in infected plants(Whitfeld and Higgins, 1976). Full length particles contain infectious single-stranded RNA of molecular weight 2 X 106 and the shorter particles containsubgenomic RNAs which serve as templates for the viral coat and movementproteins. SHMV has also been reported to complement the transport function ofbrome mosaic virus (BMV) in “Pinto” beans, which BMV normally does not infect(Taliansky et al., 1982b) and of the RNA of the B component of RCMV, (which isnot transported unless it is co-inoculated with the M component) in V. unguiculata(Malyshenko et al, 1988).The “dependent” virus, SBMV-C, is one of four strains described forSBMV. The cowpea and bean strains of SBMV share similar chemical andphysical properties (Ghabrial et al., 1967), but they differ in their host ranges.SBMV-B infects most common bean cultivars but does not infect cowpea, whereasSBMV-C is restricted to Vigna cultivars but cannot infect bean (Tremaine andHamilton, 1983). SBMV is an icosahedral virus, 30 nm in diameter, its capsidconsisting of 180 polypeptides arranged with T=3 icosahedral quasi-symmetry12surrounding the single-stranded viral RNA (Abad-Zapatero et al., 1980). Sequencecomparison of 400 bases from the 3’ end of both SBMV-B and SBMV-C showedvery little homology in the non-coding region yet extensive homology in the codingregion (Mang et al., 1982).Both the genomes of SHMV and SBMV have been sequenced. Thecomplete nucleotide sequence of the transport gene of SHMV has been determinedand the amino acid sequence of its 30-kDa protein is known (Meshi et al., 1982).On the basis of its similarity with other putative viral movement proteins, SHMVmovement protein has been grouped into a “transport” (movement) group togetherwith other tobamoviruses and with tobra-, caulimo-, nepo-, como-, and potyviruses(Atabekov and Taliansky, 1990). For SBMV, however, no transport gene has beendefinitively described, even though the complete nucleotide sequence ofits genomehas been determined (Wu et al., 1987).In this study, host and viral responses were compared in single anddouble virus infections. Inoculation of isolated bean protoplasts with SBMV-C,SBMV-B and SBMV-C plus SHMV showed that SB1VIV-C is capable of replicatingin isolated bean cells to the same extent as SBMV-B, and that its replication isnot affected by the presence of SHMV. These results would indicate that SHMVfacilitates the spread of SBMV-C in bean, rather than its replication.13The “helper effect” of SHMV on SBMV-C movement in mixedinfections ofbean was shown to be limited to short distance (cell-to-cell) movementin the inoculated primary leaves, with no virus (SBMV-C) moving into thevascular system. Relatively few SBMV-C virions were found in doubly infectedparenchyma cells, yet amorphous masses of what appeared to be SBMV-C viralcoat protein were found in these cells, probably indicating a lack of properassembly of SBMV-C coat protein in bean, which might be associated with thelack of systemic movement of the virus.141.2 ObjectivesIn spite of many recent advances that have taken place in plant viralresearch, the mechanism by which the transport gene(s) facilitates cell-to-cell andlong distance movement in plants is still unknown. Mixed infections with a“helper” and a “dependent” virus, allow comparisons to be made between singleinfections of either virus, thus providing an insight into the host-virus interaction.This study was undertaken to determine the effects of a mixed infection in whichone virus, SHMV, acted as a “helper”, and another, SBMV-C, as a “dependent” inbean, a host of SHMV but not of SBMV-CThe objectives of this research were to:1. Compare the general pattern of symptom development and antigenaccumulation in single infections with SB1VIV-C and mixed infections with SBMVC and SHMV in four bean cultivars.2. Determine whether the increase in the accumulation of SBMV-C in bean coinoculated with SHMV is due to complementation of a replicative or a movementfunction.153. Establish the pattern of accumulation, throughout the plant, of SBMV-Cand SBMV-B in mixed infections with SHMV.4. Study the distribution of SBMV-C in cells of doubly infected bean tissue,and compare and determine differences with that of SBMV-B in bean and that ofSBMV-C in cowpea.16Chapter 2Biological characterization of the interaction between SBMV-C andSRMV2.1 IntroductionVisible or otherwise detectable abnormalities in plants, termedsymptoms of a disease, can be caused by viruses. When viruses cause symptomswhich result in a significant deviation from normal growth of a crop plant, theybecome economically important. In mixed infections, viruses may interact toproduce a disease which is not caused by either virus on its own, or may increasethe severity of the symptoms (Reviewed by Matthews, 1991).In this chapter, symptom development during single and mixedinfections with SHMV and SBMV is described for four bean cultivars. The effectof SHMV on the amount of SBMV produced in these cultivars was monitoredsince, in many instances, the helper effect of a virus will be reflected in anincrease in the amount of the “dependent” virus (for examples see: Dodds andHamilton, 1972; Hamilton and Nichols, 1977; Goodman and Ross, 1974; Ishimotoet aL, 1990).Other factors such as time of inoculation, increase in temperature of17incubation and the effect of multiple passage on the interaction between SBMV-Cand SHMV were also studied. Finally, the influence of other viruses on thereplication of SBMV-C in bean was determined.2.2 Materials and methods2.2.1 Virus propagation and purificationThe different virus isolates used were obtained from the dried,infected tissue collection of the Agriculture Canada Research Station, Vancouver.SBMV-B and SBMV-C were maintained in Phaseolus vulgaris “Bountiful” andVigna unguiculata L., respectively, grown under greenhouse conditions at 20-26C. Both virus strains were purified from infected leaf tissue 12-18 days afterinoculation according to established protocols (Tremaine et al., 1981). Tissue wassprinkled with sodium diethyldithiocarbamate (quantity corresponding to 0.02 Min final extraction volume) and then ground in 0.2 M sodium acetate buffer (pH5.0) with 0.1% mercaptoethanol. The pulp was sieved through cheese cloth, theextract was adjusted to pH 4.8 by addition of 10% acetic acid, left at 5° C for 4hours and then clarified by centrifugation at 12,000 x g for 15 mm. Polyethyleneglycol (PEG) and NaC1 were added to the supernatant to give 8% (w/v) and 0.01M respectively, and the mixture was stirred for 1 hr at 4° C. The precipitate wasrecovered by centrifugation at 12,000 x g for 15 mm. and the pellet was18resuspended in 0.1 M sodium acetate buffer pH 5. The precipitation procedurewas repeated, after which virus was further purified in 10-40% linear sucrosedensity gradients, prepared by the method of Davis and Pearson (1978), bycentrifugation in a Beckman SW 40 rotor at 26,000 x g for 3.5 hrs at 4 C. Thevirus, which appeared as an opalescent band near the centre of each gradient, wasrecovered with a needle and syringe and dialysed extensively against 0.1 Msodium phosphate buffer pH 7. Virus concentration was determinedspectrophotometrically (E= 5.85 [mg/m11’ cm’ at 260 nm).SHMV was propagated in P. vulgaris “Bountiful” and purifiedby the method of Kassanis and Varma (1975). Leaves were harvested 15-18 daysafter inoculation and ground in a blender with 2 volumes of 0.2 M K phosphatebuffer pH 5.2. Sap was squeezed through cheese cloth and concentrated by twoor three cycles of differential centrifugation (10 mm at 10,000 X g; 1.5 h at100,000 X g) and the pellets were resuspended in water or 0.01 phosphate bufferpH 7.0. Virus concentration was determined spectrophotometrically (E= 3.2[mg/mli’ cm’ at 260 nm).Other viruses that infect bean systemically were tested fortheir effect on the replication of SBMV-C in bean. SBMV-C was paired withalfalfa mosaic virus (A1MV), bean yellow mosaic potyvirus (BYMV), tomatoringspot nepovirus (ToRsV), bean golden mosaic geminivirus (BGMV) and bean19rugose mosaic comovirus (BR1VIV), and propagated in “Bountiful”. Infected tissuewas harvested and kept at 700 C until needed.2.2.2 Virus inoculationFully expanded, Carborundum-dusted primary leaves of fourcuitivars of P. vulgaris (“Bountiful”, “Pinto”, “Top Crop” and “Tendergreen”) weremechanically inoculated (foam pad) with purified SBMV-C or SHMV (0.1 mg/mi)in 0.01 phosphate buffer (pH 7.0), or with a 1:1 mixture ofboth virus preparations(SBMV-C/SHMV both at 0.1 mg/mi). To test the effect of other viruses on thereplication of SBMV-C, infected leaf tissue was homogenized in 0.01 M phosphatebuffer (pH 7.0) and primary leaves were inoculated with each virus alone ortogether with SBMV-C.2.2.3 Collection and storage of samplesInoculated primary leaves and trifoliate leaves from 4 to 6different plants were harvested at different times post-inoculation and washedunder running tap water for 2-4 h in order to remove any viral inoculum from theleaf surface. Leaves were dried, weighed and stored in plastic bags at -70° C untiltested.202.2.4 BioassayPresence of SBMV-C infectivity was determined by bioassayon a cowpea local lesion host (V. unguiculata 399419 or Georgia 21). Sampleswere ground in 0.01 M potassium phosphate buffer pH 7.0 and the homogenatewas rubbed onto primary leaves of the cowpea plants. Local lesions appearingon leaves were counted 5-7 days post-inoculation.A systemic host was used to assess the presence of the virusin SBMV-C singly inoculated bean leaves because of the minimal amount of locallesions produced in local lesion hosts. Primary bean leaves were harvested 7 and15 days after inoculation with SB1VW-C and tissue homogenates were rubbed ontoa cowpea cultivar which supported systemic infection of SBMV-C.2.2.5 Production of antibodies and detection of antigen by ELISAThe amount of SBMV-C antigen was determined by thedouble antibody sandwich enzyme-linked immunoassay (DAS-ELISA) (Clark andAdams, 1977). Antibodies used for ELISA were raised in rabbits following fiveweekly intramuscular injections, 1 mg each, of purified SBMV-C. Rabbit bloodwas collected when titres reached 1:512 (rabbit H2) and 1:1024 (rabbit Hi).Immunoglobulins were precipitated from antisera with ammoniuin sulphate and21subsequently purified by DEAE-cellulose column chromatography as described byClark, Lister and Bar-Joseph (1986). Concentration of immunoglobulins wasadjusted to 1 mg/mi (A280= 1.4) and stored at 200 C.Flat-bottomed, 96-well, Linbro microtiter plates (FlowLaboratories, Mississauga, Ontario) were used for ELISA tests. Wells were coatedovernight at 4 C with SBMV antibodies by the addition to each well of 200 iil ofpolyclonal anti-SBMV immunoglobulin diluted to 1 jig/mi in phosphate-bufferedsaline (PBS)(127 mM NaC1, 2.6 mM KC1, 8.5 mM Na2HPO4,1.1 mM KH2PO4).They were then blocked with 0.2% BLOTTO (Johnson et al., 1984) in PBS(BLOTTO is 10 g of nonfat, dried milk made up to 100 ml with distilled water and0.02% sodium azide as a preservative), for 30 mm at room temperature. Leaftissue from inoculated and healthy plants was ground with a mortar and pestlein 1 ml sample buffer (0.05% Tween 20, 2% polyvinylpyrrolidone [PVP] and 0.2%ovalbumin in PBS)/0.1 g of leaf tissue, and 200 jil samples were added to eachwell. The plates were incubated for 1 h at 37 C and then washed with tap waterthree times. SBMV-immunoglobulin-enzyme conjugate was diluted in freshlyprepared sample buffer (200 jil conjugate/20 ml buffer), 200 p1 was added to eachwell and plate was incubated for 45 mm at 37 C. Plates were washed as above,substrate (p-nitrophenyl phosphate, Sigma Chemical Co., St. Louis, MO) at 0.5mg/mi in 10% diethanolamine pH 9.8 was added to the wells and the plates wereincubated for 1 h at 37 C. The absorbance of each well was recorded at 405 nm22(A405) in a Titertek Multiscan MCC plate reader (Flow Laboratories).2.2.6 Temperature studyThe effect of temperature on symptom development and virusconcentration was determined by inoculating “Pinto” and “Bountiful” primaryleaves with a mixture of SBMV-C and SHMV and incubating the plants at 32° C.Symptom development was recorded at 7, 14 and 21 days post-inoculation.Samples were collected and assessed for virus as described above.2.2.7 Multiple passage studyTwelve plants each of “Bountiful” and “Pinto” were inoculatedwith a mixture of SBMV-C and SHMV as described in section 2.2.2. After 15-20days, primary leaves were homogenized and re-inoculated to another set ofplants.This procedure was repeated 15 X; the experiment lasted from April 28 1988 untilApril 7, 1989. SBMV-C accumulation and infectivity on cowpea was tested everythree passages. After the seventh passage, SBMV-C was purified on a sucrosegradient (section 2.2.1) and its infectivity was tested on “Pinto”.232.3 Results2.3.1 Symptoms in singly and doubly inoculated plantsSymptoms in primary and trifoliate leaves of four beancultivars (“Pinto”, “Bountiful”, “Top Crop” and “Tendergreen”) were recorded 10,15 and 20 days after inoculation with SB1VIV-C alone, SHMV alone and a mixtureof the two viruses.None of the cultivars, singly inoculated with SBMV-C,presented symptoms (Fig. 1A, B). Primary and trifoliate leaves had the sameappearance and developed at the same rate as those in mock-inoculated plants.Symptoms of SBMV-C/SHMV mixed infections varied between the seasons inthree of the four cultivars (“Bountiful”, “Tendergreen” and “Top Crop”), yet werequalitatively similar to those in plants inoculated with SHMV alone. During themonths of June, July and August, primary leaves infected with both virusesshowed severe symptoms (Fig. 2A) while primary leaves inoculated with SHMValone showed no symptoms or very slight chiorotic symptoms (Fig. 2B). Duringthe rest of the year, primary leaves of SBMV-C/SHMV- and SHMV-inoculatedplants presented chlorosis and ring-like lesions extending from the mid-riboutwards, while trifoliate leaves developed a rugose yellow and dark green mosaic(Fig. 3). The relative severity of symptoms in doubly inoculated plants was notcorrelated with variations in SBMV-C concentration.24Fig. 1. “Pinto” bean singly inoculated with SBMV-C. A) Inoculated primary leafshowing no visible symptoms 7 days post-inoculation; and B) Whole plantpresenting no symptoms 20 days post-inoculation. The same results were obtainedwith “Bountiful”, “Top Crop” and “Tendergreen”.25‘-Iri-IFig. 2. Symptoms, during the months of June, July and August, in “Bountiful”primary leaves inoculated with: A) Mixture of SBMV-C and SHMV; and B)SRMV alone. Doubly-infected leaves presented severe symtoms, while singly-infected leaves presented only slight chlorosis or no visible symptoms.27ccC.,’—I1:14Fig. 3. “Bountiful” plant, doubly-inoculated with SHMV and SBMV-C. Primaryleaves presented chlorosis and ring-like lesions while trifoliate leaves developedyellow and dark green mosaic and blistering.29In “Pinto”, mixed infections resulted in pinpoint local lesionsin the inoculated primary leaves (Fig. 4A) which were not induced upon singleinfection with either virus (not shown) but were similar, although smaller, tothose produced by single infection with SBMV-B (Fig. 4B)2.3.2 SBMV-C accumulation in singly and doublyinoculated plantsUpon inoculation with SBMV-C, the four bean cultivarsproved to be non-permissive hosts, allowing only subliminal replication ininoculated primary leaves. Replication at this level was not detected on the locallesion host V. unguiculata Georgia 21, and A405 values in ELISA were in the rangeof values found for mock-inoculated tissue. Sap from leaves of the four cultivarsinoculated with SBMV-C was infective in a systemic host for the virus.When these same bean cultivars were inoculated with amixture of SHMV and SBMV-C, SBMV-C was readily detected in the inoculatedprimary leaves by bioassay (Fig. 5) and by ELISA (Fig. 6). Homogenates of“Bountiful” primary leaves showed the greatest infectivity followed by“Tendergreen” and “Top Crop”, while “Pinto” showed the lowest. These resultswere correlated with the amount of SBMV-C capsid protein detected by ELISA inthe four different cultivars. Low infectivity and ELISA values found for “Pinto”30Fig. 4. Symptoms in “Pinto” primary leaf inoculated with: A) SHMV and SBMVC; and B) SBMV-B. Small necrotic local lesions developed in the mixed infection(A), while similar, but larger, necrotic local lesions, which later turned lightbrown, developed in “Pinto” primary leaves inoculated with SBMV-B.31-ricz605040‘I.30U)____020-J100BCu ItivarFig. 5. Detection of SBMV-C by bioassay on cowpea “Georgia 21” in primary leavesof different bean cultivars doubly inoculated with SHMV and SBMV-C. (P, “Pinto”;TC, “Top Crop”; T, “Tendergreen”; B, “Bountiful”).P TC T330.40.3In0.20.10CultivarFig. 6. Detection of SBMV-C antigen, by ELISA, in primary leaves of four differentbean cultivars doubly-inoculated with SBMV-C and SHMV. (H, Healthy; P,“Pinto”; TC, “Top Crop”; T, “Tendergreen”; B, “Bountiful”.)H P TC T B34can be explained by the fact that the virus is localized in necrotic local lesions.Trifoliate leaves of the four doubly inoculated cultivars wereharvested at different intervals up to 60 days after inoculation and tested for thepresence of SBMV-C by bioassay (on both systemic and local lesion hosts) andELISA. No virus was detected in the trifoliate leaves of any of the cultivars atany time.The effect of SHMV on the accumulation of SBMV-B wastested in cowpea but there was no measurable increase in the accumulation of thelatter, therefore the helper effect was not reciprocal in cowpea.2.3.3 Effect of sequential inoculationReplication of SBMV-C in primary leaves of the four beancultivars was detected when SBMV-C was inoculated up to 72 h after SHMVinoculation. When SBMV-C was rubbed onto primary leaves prior to SHMV, itsreplication could be detected when the interval between inoculations was shorterthan 24 h.No SBMV-C replication could be detected after inoculation ofeach half of a primary leaf with the individual viruses, nor was SBMV-C detected35in any part of the plants after simultaneous or sequential inoculation of trifoliateleaves with SBMV-C and SHMV.2.3.4 Effect of temperature shiftsTwo of the four cultivars described above (“Bountiful” and“Pinto”) were inoculated and maintained at 32 C. Results of several experimentsdemonstrated that the amount of SBMV-C antigen, measured by ELISA, inprimary leaves was greater and symptoms were more severe althoughqualitatively similar. Despite these results, SBMV-C was not found in trifoliateleaves of these plants when tested by bioassay and ELISA.2.3.5 Effect of multiple passageAfter sequential passage of SBMV-C and SHMV mixedinfections through 15 cycles of “Bountiful” plants, SBMV-C was virtually lost. Thedecline in SBMV-C was gradual and was accompanied by an increase of theseverity in symptoms produced by SHMV on the trifoliates. After seven passages,the viruses were purified from the primary leaves of “Bountiful”, yet no SBMV-Cwas recovered from the sucrose gradient, although the initial homogenate wasinfectious on a local lesion host for SBMV-C. When fractions from the sucrosegradient were tested on Pinto, samples from one visible band, found in the36position expected for SHMV, induced symptoms typical of those of SHMV in singleinfection.2.3.6 Effect of other viruses that replicate in bean on the replicationof SBMV-CFive viruses from different taxononñc groups were used to testtheir effect on the replication of SBMV-C. The common characteristic shared byall five viruses was their ability to infect bean systemically. Results of thesemixed infections are presented in Table I. Bean yellow mosaic virus (a potyvirus)and bean rugose mosaic virus (a comovirus) both acted as helpers for SBMV-C.Trifoliates of SBMV-CIBYMV- and SBMV-C/BRMV-inoculated plantswere also tested by infectivity assay on cowpea. No local lesions were produced,indicating that the helper effect of these two viruses was similar to that of SHMVin that it was limited to the inoculated primary leaves.37Table I. Effect on the replication of SBMV-C by other viruses that infect beansystemicallySBMV + Symptoms SBMV-C infectiona HelperA1MVb same as singlec none noBYMV same as single 30 lesions yesTomRsV same as single none noBGMV same as single 1 +1- 0.5 lesions noBRMV same as single 50 lesions yesa Infectivity was assayed on half leaves of V.unguiculata 399419, a local lesionhost for SBMV-C.b A1MV, alfalfa mosaic virus; BYMV, bean yellow mosaic virus; ToRsV, tomatoringspot virus; BGMV, bean golden mosaic virus; BRMV, bean rugose mosaicvirus.Symptoms were indistinguishable from those of single infection by the testedvirus.38Chapter 3Replication of viruses in singly and doubly inoculated bean protoplasts3.1 IntroductionThe introduction of techniques for the inoculation of plant protoplastswith viruses (Cocking, 1966; Takebe and Otsuki, 1969) has allowed the study of someaspects of virus behaviour that are difficult to examine in whole plants. The abilityor inability of a virus to replicate in protoplasts can give some insight into the levelat which plant resistance is operating. It also allows the study of the interaction oftwo viruses in the same host plant at a cellular level (Barker, 1980).The aim of this work was to determine whether SBMV-C was able toreplicate in isolated bean protoplasts and, if so, to compare the amount of virusproduced by SBMV-C with that of SBMV-B. Replication of SBMV-C alone was alsocompared to its replication in mixed infections with SHMV, in order to establishwhether SHMV had any effect on the amount of SBMV-C produced in isolated beanprotoplasts.3.2 Materials and methods393.2.1 Protoplast isolationAll manipulations were performed under sterile conditions in alaminar flow hood. Protoplasts were isolated from fully expanded leaves of‘Bountiful” bean by the one-step method (Power and Cocking, 1969). The tissue wassurface-sterilized with 70% ethanol and rinsed three times in distilled water. Leavesthen were cut into 1 mm2 strips which were incubated overnight, at roomtemperature, in a solution of 0.6 M mannitol, pH 5.4, 0.2% cellulase Onozuka R-10and 0.025% Macerozyme (Yakult Honsha Co., Minato-ku, Tokyo, Japan). Thesuspension was filtered through Miracloth (Calbiochem Corp., La Jolla, CA),centrifuged at 1,000 X g for 3 mm, and the pelleted protoplasts were washed twicewith 0.6 M mannitol. Viability was assessed in a 10% solution ofEvans blue (Larkin,1976). Preparations with 80% or higher viability were used for viral inoculation.3.2.2 Protoplast inoculation and incubationFreshly prepared protoplasts were inoculated with SBMV-C,SBMV-B, or SHMV in a 30% PEG solution (Maule, 1983). Briefly, 10 or 20 .tg ofvirus was added to approximately 2 X 106 protoplasts. The cells were resuspended ina minimal volume of 30% PEG in 0.6 M mannitol and 3 mM CaC12,added within 30s while gently swirling to avoid protoplast disruption. The suspension wasimmediately diluted 10-fold with a solution of 0.6 M mannitol and 1 mM CaC12 and40was allowed to stand at room temperature for 5 mm. The inoculated protoplasts werewashed three times with 0.6 M mannitol and 1 mM CaCl2 at pH 5.6 and resuspendedto a final concentration of 5-10 x io cells/ml in a modified minimal salt medium(Takebe,1977) (0.2 mM KH2PO4,1.0 mM KNO3, 10 mM MgSO4,1 mM CaCl2, 1 j.tMKI, 0.01 .tM CuSO4and 50 p.g/ml gentamycin) at pH 5.4 (adjusted with HC1 or KOH).The protoplast suspension was incubated at 25° C under constant illumination ofabout 10,000 lux. At selected intervals, 1 ml samples were collected, centrifuged andstored at -70 C until further use.3.2.3 Assays for virus in protoplastsPresence of SBMV-C, SB1VIV-B and SHMV in protoplasts wasassayed by ELISA. Frozen, inoculated protoplasts were thawed, resuspended in 1 mlof 1 X PBS (20 mM phosphate buffer, pH 7.4, and 150 mM NaCl), sonicated andplaced in pre-coated wells as described in section 2.2.5. SBMV-C infectivity wastested on a cowpea local lesion host (V. unguiculata 399419). Inoculated protoplastswere resuspended in 1 ml of 0.01 M potassium phosphate buffer, pH 7.0, sonicatedand the extract was rubbed on eight half leaves of 1 week old cowpea plants.Numbers of lesions were recorded 7 days after inoculation.3.2.4 Protoplast fixation and fluorescent antibody staining41Protoplasts from inoculated “Bountiful” and “Pinto” leaves wereisolated as described in section 3.2.1. After enzyme digestion, cells were washed with0.6 M mannitol and a drop of the protoplast suspension was smeared onto amicroscope slide, previously coated with a layer of Mayer’s albumin (Cassels andGatebny, 1975), and dried in a stream of warm air.Fixation and staining were done essentially as described by Wood(1985). The protoplasts in the smear were fixed in acetone for 30-45 mm, gentlyrinsed with distilled water to remove mannitol, then equilibrated in PBS. SBMVantiserum was diluted 1/100 in PBS, a drop was placed on top of the smear and theslide was incubated at 37 C for 1 h in a moist chamber. After washing for 30 mmwith PBS, fluorescein-labelled, sheep anti-rabbit iminunoglobulin (Sigma ChemicalCo., St Louis, MO), diluted 1/20 with PBS, was added and incubated for 30 mm. Theslide was washed thoroughly in PBS, mounted in 10 mM sodium carbonate buffer in90% glycerol, pH8-9, and examined under a fluorescence microscope fitted with anexcitation filter (KP500) and barrier filters (LP520 and LP540).3.3 Results3.3.1 Proportion of SBMV-C infected cells in singly and doubly infectedleaves42Fluorescent antibody staining ofprotoplasts isolated from primary leavesof “Pinto” and “Bountiful” doubly inoculated with SBMV-C and SHMV resulted in thedetection of some stained protoplasts. In “Pinto” only 0.1% of the cells fluorescedwhile in Bountiful 5% of the cells fluoresced. Protoplasts from plants that weresingly inoculated with SBMV-C failed to fluoresce, probably due to the low amountof antigen present in subliminal infections.3.3.2 Replication of SBMV-C in singly inoculated protoplastsTo confirm that bean cells can support viral replication, Bountifulprotoplasts were inoculated with SBMV-C. Viral antigen in infected protoplasts couldbe detected using ELISA, in most experiments, 10 h after inoculation. It increasedsteadily until 42-58 h and remained constant thereafter. Bioassays of virusinfectivity were positive about 30 h after inoculation, and lesion numbers increasedsteadily until 68 h (Fig.7). Mock-inoculated protoplasts showed no increase inabsorbance values which were, on average, 0.015 at A405.Protoplast viability was assessed every time a sample was taken.The greatest decline was observed immediately after inoculation (viability decreasedfrom 80-90% to 60-70%) and at 48-53 h after placing in minimal salt medium(viability varied with different protoplast preparations).43U)Cl)00f IFig.7. Time course of the replication of SBMV-C in “Bountiful” protoplastsdetected by ELISA (.) and bioassay (i.) on Vina unguiculata L. var 399414.44Hours post- inoculation3.3.3 Replication of SBMV-C in doubly inoculated protoplasts incomparison to SBMV-B-inoculated protoplastsA single batch ofprotoplasts was divided into two equal parts; onewas inoculated with SBMV-C alone and the other with a mixture of SBMV-C andSHMV. This was done to determine whether the presence of SHMV would affectSBMV-C replication in bean protoplasts. Accumulation of SBMV-C antigen afterdouble infection of protoplasts with SHMV and SBMV-C was similar to that attainedwhen protoplasts were singly inoculated with SBMV-C (Table II).To determine the replicating efficiency ofthe two strains ofSBMV,two equal parts of a batch of protoplasts were infected with either SBMV-C orSBMV-B. The infection courses and antigen accumulation for both strains weresimilar (Table II). Thus, individual bean protoplasts sustained replication ofSBMV-Cand SBMV-B with the same efficiency, and the level of replication of SBMV-C in theisolated protoplasts was unaffected by co-inoculation with SHMV.45TABLE II. Replication of SBMV-C and SBMV-B and replication of SBMV-C insingle and mixed infections with SlIMY in Bountiful bean protoplasts detected bydouble antibody sandwich enzyme-linked immunosorbent assay (]JAS-ELISA).Absorbance (A/104protoplasts)Time post inocu- SBMVCa SBMV-B SBMV-C SBMV-Clation (hours)— +SHMV0 0.089 0.031 0.314 0.4385 0.044 0.008 0.272 0.37112 0.022 0.003 n.d n.d.24 0.377 0.414 0.275 0.34546 fl.d.” n.j 0.749 0.867a Experiment on first two columns was done simultaneously with the samepreparation of Bountiful bean protoplasts that was divided into two equal parts,one for each treatment and the last two columns with another lot of bountifulbean protoplasts also divided into two equal parts, one for each treatment.b n.d. indicates not determined.46Chapter 4Distribution of SBMV-C within inoculated leaves and throughout theplant4.1 IntroductionOne means of controlling the pathological effects of viruses isby blocking their systemic spread throughout the plant. In order to achieve thisgoal, it is necessary to understand how viruses move through plants and the wayvirus distribution varies within the tissues of the infected plants.In this study, the spread of SBMV-C in bean, in mixedinfections with SHMV, was followed using a combination of ELISA, dot-blothybridization, and light and electron microscopy. A parallel study, using anotherstrain of the virus which infects bean systemically (SBMV-B), allowed comparisonof the distribution of a virus in its host (SBMV-B in bean) with a “dependent”virus in a non-host (SBMV-C in bean with SHMV acting as the “helper”).4.2 Materials and methods4.2.1 Collection of samples47Various tissues from “Bountiful” inoculated with SHMV andSBMV-C or SHMV and SBMV-B were harvested from 6 different plants 5,7 and11 days post-inoculation. Inoculated primary leaves were washed under tap waterfor 2 h, and interveinal tissue was separated from the lateral veins, main vein andpetiole with the aid of a scalpel. Stems, roots and trifoliate leaves were alsoseparated. All tissues were weighed and used immediately in assays forinfectivity of viral particles and viral RNA or stored at -20 C until future use.4.2.2 ELISA and bioassayEqual weights of different separated plant parts were groundin 1 X PBS (for ELISA) or 0.01 M potassium phosphate buffer, pH 7.4 (forbioassay) and tested for the presence of SBMV-C or SBMV-B as described insections 2.2.4 and Dot blot hybridization assay for viral RNATotal leaf RNA from different leaf sections of inoculated andhealthy plants was extracted following the method of Siegel et al. (1976). Leafmaterial (100-500 mg) was ground to a fine powder in liquid nitrogen using amortar and pestle and transferred to a microfuge tube to which was added 400 ulphenol/chloroform! octanol (25/24/1) and 400 ul lox TNE (100 mM Tris-HC1 pH487.5, 100 mM NaCl and 10 mM EDTA), 0.1% SDS and 5% 2-mercaptoethanol.After mixing vigorously, the suspension was centrifuged (10,000 X g) for 2 mmand the aqueous phase was removed and re-extracted two times (second time withchioroform/octanol only). One tenth volume of 2 M sodium-acetate pH 5.8 and 2-2.5 volumes of absolute alcohol were then added to the aqueous phase and theRNA was precipitated at -70 C for 10 mm. The precipitate was pelleted bycentrifugation, washed with 70% ethanol and resuspended in sterile, deionizedwater.For further separation of high molecular weight ssRNA,instead of washing and resuspending the pellet in water, it was resuspended inTE (10 mlvi Tris-HCL pH 7.5 and 1mM EDTA), 100 p.1 of 8 M LiCl was added andthe solution was incubated on ice for 1-3 h. After spinning for 10 mm, the pelletwas resuspended in autoclaved water, re-precipitated with NaAc and ethanol asdescribed above, and resuspended in sterile, deionized water. The supernatantfluid remaining after LiCl precipitation was removed and 2.5 volumes of absoluteethanol were added and the suspension was centrifuged in a microfuge for 10 mm.The pellet containing dsDNA, dsRNA and low molecular weight ssRNA waswashed with 70% ethanol and resuspended in sterile H20. All samples werestored at -70 C.Presence of SBMV-C RNA was assessed by dot- blotting using49a nick-translated (Rigby et al., 1977)32P-labeled probe (107108 cpm/p.g) of SBMVcloned DNA designated “slorf 18” (kindly provided by Dr. Claire A Rinehart,University of Wisconsin, Madison, WI).Dot-blot hybridization analysis was done on nitrocellulosemembranes (Schleicher and Schuell, Keene, NH), which were cut to 8 x 10 cm,soaked sequentially in deionized H20 for 5 mm, and in 20X SSC (20X SSC is 3 MNaCl, 0.3 M sodium citrate), and dried. Each sample of total nucleic acid orssRNA was thawed and 2 .tl or a volume equivalent to approximately 20 ng and40 ng of nucleic acid was deposited on the filters. Membranes were allowed to dryand were then baked under vacuum for 1 h at 80 C. The filters were thentransferred to a Seal-a-Meal plastic bag and were prehybridized at 42 C, for 1 h,in hybridization buffer (50% [v/v] deionized formamide, 50 mM Tris-HC1 pH 7.5,1 M NaCl, 0.2% bovine serum albumin [BSAJ, 0.2% PVP, 0.2% Ficoll, 0.1% sodiumpyrophosphate, 10% sodium dextran sulphate and 250 pg/ml sheared salmonsperm DNA).Hybridizations were carried out in the same solution used for prehybridization. The labelled probe was denatured by adjusting to 0.1 M NaOH andthen boiling for 5 mm. After quickly cooling the probe, it was added to thesolution in the bag and allowed to hybridize with the RNA immobilized on thefilters overnight, at 42 C with continuous agitation.50Following hybridization, the filters were rinsed with 2 X SSCplus 0.1% SDS, then washed for 15 mm, at 55-60 C successively in each of thefollowing three solutions: 2 X SSC plus 0.1% SDS; 0.5 X SSC plus 0.1% SDS andfinally in 0.2 X SSC. Membranes were blotted lightly to remove excess moisture,wrapped in plastic film and exposed to X-OMAT film (Kodak) for 2, 4 and 12 hrat -70 in an X-ray cassette with Lightning Plus (Dupont) intensif,ing screens.4.2.4 Tissue print-immunoblottingTissue prints were performed essentially as described byCassab and Varner(1987). The nitrocellulose paper (Schleicher & Schuell, #BA85,Mandel Scientific, Edmonton, AB) was pre-treated with 0.2 M CaCl2 for 30 mm,dried on paper towels and placed on Whatman filter paper on a flat glass plate.Infected and healthy leaves, petioles and stems were cut witha razor blade, rinsed briefly in distilled H20 and dried on Kimwipes facial tissue.The freshly exposed surfaces were blotted onto the nitrocellulose paper byapplying light pressure on the membrane for 10-20 s. The tissue print was thendried with warm air and either processed immediately or kept at 4 C. Theremaining free binding sites, on the nitrocellulose paper, were blocked bysubmerging the sheet in 2% BSA and 1% Triton X-100 in 0.01 M Tris pH 7.4,0.85% NaC1 (TBS) for 2 h at 37 C or overnight at 4 C.51Total leaf and viral proteins transferred by blotting weredetected by staining the blot with India ink (Hancock and Tsang, 1983). DrawingIndia ink for fountain pens (Pelikan AG, D-300, Hanover 1, Germany) was dilutedto 1 uL!ml of PBS-Tween and the blot was left overnight in stain. They wererinsed in deionized water for 5 mm and then dried.Virus antigens adsorbed on the blots were detected usingalkaline phosphatase-conjugated antibody as described by Blake et al (1984).Primary antibody against SBMV-C was diluted 1/1000, 1/2000 or 1/4000 inantibody solution (1% BSA, 0.3% Triton-X100, 0.05% Tween in TBS) andincubated for 1 h at 37 C in a shaker. The membrane was washed at roomtemperature, 10 times, 5 mm each time, in TTBS (TBS, 0.05% Tween).Anti-rabbit IgG (Fc) alkaline-phosphatase conjugate wasdiluted 1/10,000 in antibody solution and the membrane was incubated for 1.5 hat 37 C. Membranes were washed as above except that the last two washes werein substrate buffer (0.1 M NaCl, 5 mM MgCl2 in 0.1 M Tris, pH 9.5). Substratesolutions, BCIP (5-bromo-4-chloroindoxyl phosphate) and NBT (nitrobluetetrazolium), (both from GIBCO BRL Immunoselect Cat. No. 8280 SA), wereprepared just prior to use. To 10 ml of substrate buffer, 22 p1 of NBT were addedand gently mixed by inversion of the tube, then 17 p1 of BCIP were added andmixed. The membranes were incubated in substrate solution until purple prints52appeared. The reaction was stopped by addition of 5 mM EDTA in 20 mM TrispH 7.5 for 10 mm, membranes were washed in distilled H20 and dried betweenseveral layers of Whatman filter paper.4.2.5 Cryosectioning and immunofluorescent labelling of thicksectionsSamples from mixed infected and healthy tissue were preparedaccording to Griffiths (1984) post-embedding immunocytochemical technique forhigh resolution immunofluorescence.Tissue was fixed in 2 or 4% formaldehyde in 0.05 M phosphatebuffer, pH 7.0, for 1 h on ice. After rinsing with buffer, sections were infiltratedinto sucrose through a series of 20,40,60,80 and 100% sucrose (in phosphatebuffer, vlv). Samples were mounted on small aluminum stubs and immediatelyfrozen in liquid N2. At this point, samples could be stored at -70 C for furtherprocessing. Samples were sectioned on tungsten-coated glass knives with aReichert Ultramicrotome E in a cryobox (FC4E). Sections were picked up on dropsof sucrose and placed on Poly L Lysine (Sigma P1274 mw 100,500)-coated glassslides. Slides were rinsed in TBS and stored overnight at 4 C. All of the followingprocedures were done at room temperature.53Free aldehydes were blocked with NH4C1, 50 mM, for 10 mm., thenslides were left in BLOTTO for 30 mm to block nonspecific protein binding sites.Following blocking, SBMV antibody was added at different concentrations (1:100,1:500 and 1:1000, diluted in BLOTTO) and samples were incubated for 3 h. Slideswere rinsed in TBS-Tween several times, anti-rabbit IgG-FITC conjugate wasadded and samples were incubated for 1 h. After thorough rinsing, the backs ofslides were washed and samples were mounted in glycero]Jphosphate buffer (80:20v/v) and examined with a fluorescence microscope.4.2.6 Immunosorbent electron microscopyTo facilitate visualization of viral particles from homogenatesof infected plants, grids were pre-treated with protein-A and SBMV antiserum.Copper grids, 400 mesh, covered with Formvar and coated with a carbon layerwere floated on 15 .tl drops of Protein A (10 ug/ml of distilled water) for 10 mm.Grids were washed once in PBS, floated for 10 mm in 15 ul drops of SBMVantiserum diluted 1110 in PBS and washed one more time in PBS.Samples of inoculated and healthy plants were homogenizedin PBS with a mortar and pestle and pre-treated grids were floated on dropletsof sap and incubated at room temperature for 1 h. After incubation, grids werewashed by floating in five drops of PBS, 2 mm on each drop, and finally on one54drop of Bacitracin for 1 mm. Samples were stained with a 2% solution of uranylacetate (UA) for 1 mm and dried. Preparations were observed with a Hitachi 600transmission electron microscope.4.2.7 Tissue fixation and embedding for electron microscopySamples from inoculated and healthy bean and cowpea werefixed according to standard glutaraldehyde and osmium tetroxide embeddingprocedure, and two other procedures: Hatta and Francki’s (1981) procedure todigest ribosomes, and an enhanced method for fixation and gold labelling(Berryman and Rodewald, 1990).For the standard fixation procedure, leaves and petioles takenat different times post-inoculation were cut into 1 mm2 pieces and placed in 4%glutaraldehyde in 0.1 M cacodylate buffer, pH 7.0, for 12-18 h (first hour at roomtemperature and the remaining time at 4 C). Samples were washed twice for 15mm in 0.1 M cacodylate buffer pH 7.0. Specimens were post-fixed in 1% (or 0.5,or 0.25%) 0s04 in cacodylate buffer for 1 to 3 h, rinsed in distilled water and thenwashed twice, 15 mm each time, in distilled water.Fixed tissue was then dehydrated by washing twice, 15 mmeach time, in each of 30%, 50%, 70%, 95% and absolute ethanol and then three55times (15 mm each time) in propylene oxide. A solution of 50/50 propylene oxideand Epon 812 mixture (10 g Epon 812, 0.36 g of 2,4,6-tri dimethyl-aurinomethylphenol [DMP], 4.46 g of nadic methyl anhydride [NMA] and 3.76 g of dodecenylsuccinic anhydride [DDSAI), was used to begin infiltration. Samples were leftovernight in uncovered vials, then transferred to flat embedding molds filled withfresh Epon 812 mixture. Blocks were left to dry at 55 C for 48 h.Alternatively, fixed tissue was dehydrated by washing once for45 mm at 4 C in each of 50%, 70% and 90% acetone. It was then placed, for 60mm at 4C in vials with 1:1 LR White medium/acetone, followed by 7:3 LR Whitemedium/acetone. Tissue was finally infiltrated with 3 changes of 100% LR Whitemedium for 60 mm each, then transferred into flat embedding molds which wereleft at -20C under UV light to allow for polymerization of the medium.To avoid interference from ribosomes when observing infectedtissue, these were digested with RNase according to Hatta and Francki (1981).Tissue was cut in pieces as described above and then fixed for 16 hours, at 4 C ina mixture of 4% paraformaldehyde and 1% glutaraldehyde in 0.13 M phosphatebuffer pH 7.3. Sections were rinsed in 2 X SSC (SSC: 0.15 M sodium chloride and0.0 15 M sodium citrate, pH 7.0) and then washed in several changes of 2 X SSCfor 6 h at room temperature. Samples were then digested with pancreatic RNase(2 jig/mi) (type III, Sigma Chemical Co. St Louis, MO) in 2 X SSC for 16 h at 2556C. After digestion, specimens were post-fixed in 0.4% osmic acid for 1-2 h,washed repeatedly in distilled H20 and then dehydrated and embedded asdescribed for samples above.To avoid problems of osmium interference withimmunolabelling of antigenic sites, an alternative fixative without this substancewas used (Berryman and Rodewald, 1990). Samples were cut and fixed, at roomtemperature for 2-3 h, in 1% glutaraldehyde, 0.2% picric acid, 4% formaldehyde,and 0.05 mM CaCl2 in 0.1 M potassium phosphate buffer pH 7.4. A solution of3.5% sucrose in 0.1 M phosphate buffer with 0.5 M CaC12 was used as a washfollowing fixation. After washing for 2 h in several changes of the sucrosesolution, free aldehydes were quenched with 50 mM ammonium chloride insucrose:phosphate buffer for 1 h at 0 C. Phosphate ions were removed by rinsingthe samples in cold 0.1 M maleate buffer, pH 6.5 containing 3.5% sucrose.Specimens were then dehydrated and infiltrated in Epon and LR White mediumfollowing the procedures described above.Ultrathin sections were cut with a Reichert OMU2ultramicrotome using glass knives (made with an LKB Knifemaker, type 7801B).Sections were collected on 100 mesh copper or nickel grids, covered with Formvarand carbon. Sections were stained with a 4% aqueous uranyl acetate solution.574.2.8 Immunogold labelling of antigensColloidal gold was prepared by reduction of hydrochloroauricacid (HAuC14)with sodium citrate (Frens, 1973; Slot and Geuze, 1981) whichresulted in suspensions of colloidal gold particles with average diameters of 5, 10and 17 nm. The colloidal gold suspensions were then complexed with Protein Aas described by Horisberger and Rosset (1977).The affinity of the gold-Protein A complex for viral antibodywas tested on grids which had been floated on drops of tissue homogenate frominfected plants. Grids, which had been incubated on drops for 10 mm, were thensequentially washed with six drops, 5 mm each, of BSA-Tween (0.5%-0.05%) inPBS. Antiserum or purified IgG was diluted 1/100 and 1/500 in PBS-BSA-Tween(as above) and grids were floated on the drops for 1 h after which they werewashed as before. For gold labelling, drops of 5, 10 and 17 nm gold-Protein Adiluted in PBS-BSA-Tween were placed on paraflim and grids were incubated onthem for 30-60 mm. Grids were washed with 6 drops of PBS-BSA-Tween for 5mm, rinsed in PBS and incubated in 2% glutaraldehyde for another 5 mm. Beforestaining, grids were washed twice for 10 mm. All reactions were carried out atroom temperature.58Labelling of antigens on ultrathin sections mounted on gridswas essentially as described above except that grids were not floated on sap frominfected plants but were pre-treated with PBS-lysine (0.1%) for 5 mm, before thefirst PBS-BSA-Tween wash. To stain sections, grids were submerged in a 1:1dilution of lead citrate in 0.01 M NaOH for 3 mm, washed in distilled water, thenstained with UA for 5 mm., washed again and incubated one more time in leadcitrate for 3 mm. Grids were rinsed in distilled water and examined in an Hitachi600 transmission electron microscope.4.2.9 Electrophoresis and Western blotting of viral proteinsViral proteins from purified virus preparations (section 2.2.1)and from soluble protein preparations from infected plants, were analyzed bysodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) through12% gels using a discontinuous Laemmli buffer system (Laemmli, 1970) asdescribed by the manufacturer (BioRad). Following electrophoresis, gels weretransferred to 3 changes of fixative (30% methanolll0% acetic acid), and agitatedgently for 30 mm in each change. Gels were stained with Coomassie bluestaining solution (50 ml methanol, 50 ml H20, 10 m1 acetic acid and 0.2%Coomassie blue G250) for 1 h and destained overnight with destaining solution (40ml methanol, 50 ml H20 and 10 ml acetic acid). Alternatively, proteins weretransferred to teflon membranes (Millipore Immobilon P, Mississauga, Ont.) using59a wet blot apparatus (TE series transphor electrophoresis unit, Hoefer Sci.,Minnesota). SBMV-C proteins were detected using both polyclonal anti-SBMV-Cand monoclonal (kindly provided by Dr. D. Mackenzie, Ag. Can. Res. St.) antiSBMV-C antibodies, following the method described in section 4.2.4, except anti-mouse IgG (Fc) alkaline-phosphatase conjugate was used for monoclonal antibodydetection.Soluble proteins from infected and healthy plants were isolatedaccording to Gegenheimer (1990). Fresh plant material (500 jig) was homogenizedin 1:5 (wlv) ice-cold extraction buffer (50 mM Tris pH 8.0, 5 mM borate, 1.5%insoluble PVP, 5% glycerol, 20 mM DIECA and 0.0 1% mercaptoethanol). Thehomogenate was centrifuged in a microfuge for 5 mm at 4 C, the supernatant fluidwas decanted and 1:1 (v/v) 2 X SDS sample buffer (0.08 M Tris HC1 pH 6.8, 0.13%glycerol, 2.6% SDS, 0.06% 2-beta-mercaptoethanol and 0.0 16% bromophenol blue)was added. Samples were stored at -20 C until needed.4.3 Results4.3.1 Distribution of SBMV-C capsid protein compared to that ofSBMV-B in double infections with SHMVTo establish the course of infection and distribution of SBMV60C, in the presence of SHMV, in bean plants, antigen accumulation in theinterveinal tissue, lateral veins, midrib, petiole, stem, roots and trifoliate leaveswas assessed at different times after inoculation Parallel tests were done withSBMV-B in the presence of SHMV, and accumulation of viral antigen wascompared for both strains (Fig.8). In the early stages of infection, estimates ofvirus concentration in the interveinal tissue were similar for both strains,although SBMV-C was slightly higher in most of the repeated experiments. Theseresults are likely accounted for by the fact that primary leaves infected withSBMV-B and SHMV suffered severe wilting and yellowing as early as 4 days post-inoculation.Further dissection of the leaf and petiole revealed that theconcentration of SBMV-C decreased from the lateral veins toward the mid- anddistal parts of the petiole, where virtually no virus could be found. In later stagesof infection, the bean strain of SBMV was present in the roots and trifoliateleaves, whereas the cowpea strain remained confined to the inoculated primaryleaves, and no antigen was detected in stem, roots, or trifoliate leaves even at 2months post-inoculation. The absence of infective SBMV-C outside the inoculatedleaves was confirmed by bioassay of extracts of plant parts on a local lesion host.Thus, in the presence of SHMV, the cowpea strain of SBMVappeared to move through the epidermal and mesophyll cells of inoculated61Fig. 8. Accumulation of SBMV-C (D) and SBMV-B (•) capsid, detected by ELISA, inmixed infections with SHMV in: a) roots, b) stem, c) petiole, d) main vein, e) lateralvein, f) mesophyll and g) trifoliates of bean ( vulgaris cv. Bountiful). Graphs A, Band C show accumulation profiles 5,7 and 11 days post inoculation, respectively. Thesymbol “X” indicates the tissue was necrotic at sampling time and “*“ indicates thatvalues for these samples were <0.025.62070.60.504,]D.5J0.44gO.3.j ri..D 0.2 .4Oil I 1.I.Io.oI0. c d e. f 9VQrious pIQn pQrtsFig. 8-ttiUx L63primary leaves at the same rate as the bean strain. However, movement ofSBMV-C into the vascular tissue was limited or nonexistent. The antigendetected in lateral veins and midrib could be due to the presence of SBMV-Cinfected mesophyll tissue that could not be completely eliminated duringdissection and also to its presence in parenchyma cells bordering the conductingsystem.4.3.2 Distribution of SBMV-C RNA in doubly inoculated plantsAlthough the absence of infective SBMV-C outside theinoculated leaves was determined by bioassay, it was further confirmed bydetermining the distribution of SBMV-C RNA. Dot blot hybridization analysisrevealed a greater amount of SBMV-C RNA in the interveinal tissue, smalleramounts in the lateral veins and main vein, and no SBMV-RNA in the petiole orstem (Fig.9). No signal in these two plant parts was observed even after a 32 hexposure of the filter. However, the presence of very small amounts of SBMV-CRNA cannot be completely excluded by these detection methods.4.3.3 Viral antigen distribution in thick sections of leaf and petioleby tissue blot analysisEfficient transfer of leaf and viral proteins was achieved by64A Bi-.2—3-.4—5—6—7—Fig. 9. Detection of the cowpea strain of southern bean mosaic virus (SBMV-C)RNA in different parts of bean plants (Phaseolus vulgaris “Bountiful”) by dot-blothybridization using a nick-translated 32P c-DNA probe of SBMV-C. Total RNAwas extracted from each plant part and 100 ng added for each sample. 1,mesophyll; 2, extraction buffer; 3, lateral veins; 4, main vein; 5, healthy tissue; 6,petiole; 7, stem. A and B, 4 and 12 h exposure, respectively.65pressing freshly sectioned material onto nitrocellulose membranes, as shown byresults of India ink staining (Fig. 10).Samples were taken from cowpea leaves inoculated withSBMV-C and Bountiful bean leaves inoculated with a mixture of SBMV-C andSHMV. Healthy bean tissue was used as negative control. A dilution series ofpurified SBMV-C was blotted on one side of the filter to be used as a guide forcolour change when substrate was added.Of approximately 90 prints of each sample, only 14 of thosefrom bean leaves inoculated with SBMV-C + SHMV showed a positive signal.These positive strips belonged to leaves that had been inoculated 7 and 12 daysprior to sampling; none of the strips from leaves collected 21 or more days afterinoculation showed a positive signal. This contrasted with approximately 60positive signals for SBMV-C in cowpea, from samples collected 7, 12 and 21 dayspost-inoculation.Positive controls of SBMV-C in cowpea primary leaves showeda deep purple stain throughout the entire section (Fig. hA, B), or in certain areasof the mesophyll and the vascular tissue (Fig. 12A, B). Sections from beanprimary leaves infected with SBMV-C + SHMV, on the other hand, presented adeep purple colour in mesophyll and epidermal cells of the leaf but no positive66Fig. 10. Transfer of proteins from leaf sections and tissue pressed ontonitrocelitilose membranes. Proteins were stained with India ink as described inMaterials and Methods. A) Cross section of petiole from a primary bean leaf. B)Cross section of leaf blade and lateral veins of a primary bean leaf. C) Crosssection of stem of a bean plant. D) Sap from primary leaf.A B C D67Fig. 11. SB1VIV-C antigen detected in SBMV-C-infected cowpea primary leaf bladeand vein tissue by tissue print immunoblotting. A. Cross section of leaf blade andlateral vein, (magnification, bOX). B. Cross section of leaf and lateral vein,(magnification, 200X). Deep purple coloring represents areas where antigen ispresent.6869tiVFig. 12. SBMV-C antigen detected in SB1VIV-C-infected cowpea primary leaf bladeand vein tissue by tissue print immunoblotting. A. Cross section of leaf blade andlateral vein, (magnification, bOX). B. Cross section of leaf and lateral vein,(magnification, 200X). Dark purple areas indicate viral (SBMV-C) antigen ispresent.70ILaVsignal was detected in the vascular bundles of the vein (Fig. 13A, B). The lightpurple zones represent background but are clearly distinguishable from thepositive dark purple areas, as can be seen by comparing prints from mixedlyinfected primary bean leaves (Fig. 13A, B) to the healthy bean tissue prints (Fig.14A, B).4.3.4 SBMV-C antigen distribution in thick sections bycryosectioning and innnunofluorescent labellingThick sections of bean tissue inoculated with SBMV-B +SHMV, SB1VW-C + SHMV or mock-inoculated were fixed and infiltrated insucrose. Samples mounted on pins and frozen in liquid N2 were cut into 3 jimsections and treated as described in section 2.4.5.In three replicated experiments, mesophyll cells and phloemcompanion cells in tissue infected with SBMV-B + SHMV fluoresced strongly (Fig.15A, B). At antiserum dilutions of 1/100 some autofluorescence and backgroundcould be seen in healthy controls (Fig. 16A, B) while very little background wasfound at antiserum dilutions of 1/500 (Fig. 17A, B). No fluorescence was found intissue infected with SBMV-C + SHMV. In all of three replications of thisexperiment, this tissue presented great difficulties in cutting, much more so thanthe SB1VIV-B + SHMV infected tissue. The reason for this is unknown.72Fig. 13. SBMV-C antigen detected in “Bountifu1 bean primary leaf tissue doublyinoculated with SBMV-C and SHMV by tissue print inimunoblotting. Dark purplestain is present in epidermis. mesophyll and vascular parenchyma but not inconducting bundles. A. Cross section of leaf blade and lateral vein (magnification,bOX). B. Cross section of leaf blade and lateral vein (magnification, 200X).73VLVFig. 14. Tissue prints of healthy “Bountifult’primary leaf tissue showing a lightpurple background, clearly distinguishable from the positive dark purple coloringthat results when the antigen is present. A. Cross section of leaf blade andlateral vein (magnification, bOX). B. Cross section of leaf blade and lateral vein(magnification, 200X).75(Ci-1Fig. 15. Immunofluorescent labelling of a thick section of a primary “Bountiful”leaf doubly-inoculated with SBMV-B and SHMV. A) Mesophyll and vasculartissue cells under bright field microscope B) Same section under fluorescentmicroscope; mesophyll and companion cells fluoresce strongly when antiserum wasdiluted 1/500. (Magnffication 430 X).77SL9!5TVFig. 16. Immunofluorescent labelling of a thick section of “Bountiful” beanprimary leaf mock-inoculated with phosphate buffer. A) Mesophyll and vasculartissue cells under bright field microscope B) Same section examined underfluorescent microscope; some background fluorescence can be seen when antiserumwas used at 1/100 dilution. (Magnification 430 X).79089!5T.VFig. 17. Immunofluorescent labelling of a thick section of “Bountiful” beanprimary leaf mock-inoculated with phosphate buffer. A) Mesophyll and vasculartissue cells under bright field microscope B) Same section examined underfluorescent microscope; practically no background was present when antiserumwas used at 1/500 dilution. (Magnification 430 X).81CoZ444.3.5 Viral particle distribution in thin sections of leaf and petioleby standard osmium tetroxide fixationUltrathin sections of tissue embedded in LR White medium orEpon were observed with the electron microscope. Samples from three differenttreatments were compared; SBMV-B + SHMV- and SBMV-C + SHMV-infectedBountiful primary leaves and healthy Bountiful primary leaves. An average of 32different grids for each treatment was examined.Both viruses are clearly visible in mesophyll and phLoemcompanion cells in sections of SBMV-B + SHMV-infected material (Fig. 18A, B).SBMV-B was found in great amounts in vesicles in the cytoplasm as well as in thenucleus. In sections of SBMV-C + SHMV-infected tissue, only SHMV was clearlyvisible in mesophyll and phloem companion cells, but no vesicles with SBMV-Cparticles were found in the cytoplasm (Fig. 19). Some “virus-like” particles couldbe seen alongside the plasmalemma, but these were very much like ribosomes alsofound in healthy tissue (Fig. 20A, B).In an attempt to distinguish virus particles from ribosomes, thetissue was treated with RNase. The RNase destroyed the ribosomes withoutharming the virus particles. In SBMV-B + SHMV-infected tissue, some83Fig. 18. Thin section of Epon-embedded ‘Bountiful” primary leaf tissue doublyinocuLated with SHMV and SBMV-B. Both viruses were found in large amountsboth in cytoplasm and in vesicles in mesophyll and companion cells. A) View of acell with vesicles and cytoplasm containing both viruses; ye, vesicles; m,mitochondria; cw, cell wall; cl, chloroplast; v, virus. B) Close-up of two vesicles,one with spheres and another one with rods corresponding to SBMV-B and SH1VW,respectively. (Bars represent 1 um).84/--*---•*••—.i;-••--•-M3_____‘•••.•••••--••_%•1••-•-I•,:8!5T-.‘b4;-—•••I.Fig. 19. Thin section of Epon-embedded Bountiful primary leaf tissue doubly-inoculated with SHMV and SBMV-C. No spheres in vesicles or cytoplasm arevisible, but large amounts of rods are present in some cells and are clearly visiblewhen aligned in rows (v, virus). cc, companion cell; ci, chioropiast; v, virus; Se,sieve element. (Bar represents 1•_Fig. 19ap0y rI -‘U1¶ rrt‘9• aFig. 20. Thin sections of Epon-embedded “Bountiful” primary leaf tissue. A) Cellin leaf doubly inoculated with SHMV and SBMV-C; only rods are clearly visible,although ribosomes could be confused for viral spheres. B) Cell in leaf (mockinoculated) showing ribosomes in the cytoplasm; r,ribosomes; cw, cell wall. (Barsrepresent 250 nm).8889Fig. 20)r.i% •;.‘rn f’aI• r’rdeterioration of the tissue was observed after the treatment (general view ofseveral cells in Fig. 21A, with details in B,C), but there were virtually noribosomes present and both viruses were clearly visible (Fig. 21B, C). Tissueinfected with SBMV-C + SHMV was very deteriorated; once again SHMV waspresent yet there was no evidence of SBMV-C particles in the cells (Fig.22A, B).Healthy controls also showed membrane disruption after treatment yet there wereno ribosomes in the cells, proving that the enzyme treatment was effective (Fig.23A, B).4.3.6 Viral particle distribution in thin sections of leaves byimmunogold labellingGold labelling was first tested on grids which had been floatedon a homogenate of SBMV-C- or SBMV-B-infected tissue. Whole antisera as wellas purified IgG were used; dilutions of 1/100 gave satisfactory results (Fig. 24 A,B).Tissue from many different SBMV-C + SHMV-infected beanleaves was observed over a period of nine months. During these observations, noSBMV-C could be seen in any of the sections, although SBMV-B could be readilyseen in SBMV-B + SHMV-infected bean and SBMV-C could be seen in SBMV-Cinfected cowpea tissue.90Fig. 21. Thin sections of RNase-treated “Bountiful” primary leaf tissue doubly-inoculated with SHMV and SBMV-B. A) General view of a group of cells withlarge vacuoles. The granular matrix in the vacuoles (bottom left cell) as well asthe granular material in the nucleus (top right cell) are representative of massesof virus particles shown at higher magnifications in B and C. B) Nucleus withSBMV-B and cytoplasm with SB1VW-B and SHMV, C) Higher magnification (10 Xthat of B) of cytoplasm in B, both spheres and rods are present. (Bars represent 1urn).91‘c:’I.‘-Ic”JZ4cq.Q-Fig. 22. Thin section of RNase-treated “Bountiful” primary leaf tissue doublyinoculated with SBMV-C and SHMV. A) Degraded ribosomes can be seen asclumps of amorphous material, vacuoles and cytoplasm contain large amounts ofS}IMV. v, virus; cc, companion cell; se, sieve element. B) Higher magnification ofcytoplasm with virus and degraded ribosomes. (Bars represent 500 urn).93‘5’A5,‘S94Fig. 22‘.9”., A5..-:.‘.q. ‘ •-f :L;c;•‘:“i1r’.,•.4se=‘ ;: ••4S:”,Fig. 23. RNase-treated “Bountiful’ primary leaf tissue mock-inoculated withphosphate buffer. A) Ribosomes in treated tissue can be seen as amorphousmasses or in clumps, their distinct shape lost. B) }{igher magnification of cellswith degraded ribosomes. cl, chioroplast. (Bars represent 1 urn).95H(Y)cNCDC)‘-Fig. 24. Specific gold-labelling of SBMV in leaf-clips of homogenates of infectedtissue A) Homogenate of SBMV-B + SHMV-infected bean, spheres specifically arelabelled. B) Homogenate of SBMV-C + SHMV-infected cowpea, spheres arespecifically labelled. g, gold; v, virus. (Bars represent 100 nni).97-H114a)Given these results, tissue was fixed for gold labelling after ithad been blotted on nitrocellulose paper and tested for the presence of antigen, asdescribed in section 4.2.5. Strips which showed positive signals on the filters werefurther treated and embedded in Epon and L.R. White resins. The fixationprocess involved the use of formaldehyde-glutaraldehyde and picric acid, andavoided the use of osmium tetroxide in order to maintain the antigenicity of theviral proteins.After sectioning the blocks, several antibody and goldconcentrations were assessed to determine optimal conditions for labelling. Wholeantiserum and purified IgG were used at a 1/100 dilution and 15 mn and 10 nmgold particles were used at a dilution of 1/5 (v/v). Positive controls included beantissue inoculated with SBMV-B + SHMV and cowpea tissue inoculated withSBMV-C. In both cases, large amounts of gold-labelled virus particles could befound in the mesophyll (Fig. 25A, B, C and Fig. 26A, B, C) and vascularparenchyma. No SBMV-C particles were found in bean tissue inoculated withSBMV-C + SHMV (Fig. 27A, B), but in some of the mesophyll cells, clumps ofamorphous material appeared heavily labelled (Fig. 28 A, B, C and 29A, B).These clumps were seen in large vacuoles where SHMV particles were clearlypresent. These clumps were never observed in tissue infected with SBMV-B +SHMV, yet similar amorphous material has been observed in protoplastsinoculated with cowpea chiorotic mottle virus (van Lent, 1988).99Fig. 25. Gold-labelling of SBMV-B antigen in thin sections of “Bountiful” primaryleaf tissue doubly-inoculated with SBMV-B and SHMV. A) Partial view ofmesophyll cell with SBMV-B virus particles in nucleus and cytoplasm labelledwith protein A-gold. Cl, chloroplast; N, nucleus; n, nucleolus. B) Highermagnification of virus particles in nucleus; g, gold particles; v, virus. C) Highermagnification of nucleus and nucleolus. (Bars represent 500 nm).100rLFig. 26. Gold-labelling of SBMV-C antigen in thin sections of cowpea primary leaftissue doubly-inoculated with SBMV-C and SHMV. A) Partial view of mesophyllcell with SBMV-C virus particles in the cytoplasm. B) and C) Highermagnifications of the same area showing specific-labelling of the virus particles. v,virus; g, gold; cl, chloroplast. (Bars represent 500 nm).102coI/.;c...:I.J%.;Y•,J:ir,/;L.(I.)•Fig. 27. Thin section of “Bountiful” primary leaf doubly-inoculated with SBMV-Cand SHMV. A) No gold label was found in the nucleous, yet great amounts ofSHMV could be seen in most cells. N, nucleous; cl, chioroplast. B) Somebackground labelling could be found in the cell wall, but no virus was labelled inthe cytoplasm. cw, cell wall; c, cytoplasm; v, virus. (Bars represent 500 nm).104105Fig. 28. Gold-labelling of thin section of “Botif” primary leaf doublyinoculated with SBMV-C and SHMV. A, B and C are different magnifications ofclumps of amorphous material found in vacuoles in some of the mesophyll cells. g,gold; v, virus (rod); p, protein. (Bars represent 500 nm).10644.t•ii3,61.0cc:LOI1Stt-2’t-*:c;.1 44-lc1’-4. -I-jA?:I-am.•‘I’Fig. 29. Gold-labelling of thin sections of “Bountiful primary leaves doublyinoculated with SBMV-C and SHMV. A and B are different magnifications ofclumps of heavily-labelled amorphous material found in vacuoles of mesophyllcells. v, virus (rods); g, gold; p, protein. (Bars represent 200 nm).108•C.4t.4.I••.cSVV:VW’b•VVV•••‘V••:—.VV_V••V•*—••.•••••S.V-.•VSV-•VV•V?VVVV••S.—a.•.•••..SV5•VI.4.3.7 Electron microscopy observations after trapping ofSBMV-C in mixed-infected leavesImmunosorbent electron microscopy of extracts from primary beanleaves doubly inoculated with SB1V1V-C and SHMV resulted, in some cases, ingrids covered with small particles of about 15-20 nm (Fig. 30A). These particlescorrespond to the T=1 structure which is assembled from 60 coat protein subunitsinstead of 180 subunits in typical virions. Homogenates from cowpea infectedwith SBMV-C and trapped with the same antibody resulted in grids covered withparticles of about 28-30 nm (Fig. 30B), the size of typical virions (Hull,1987). Novisible particles were seen in extracts of crude sap from healthy plants treated inthe same manner (Fig. 30C).4.3.8 Electrophoresis and Western blottingTotal soluble proteins from both infected and healthybean and cowpea were electrophoresed on a denaturing SDS-polyacrylamide gel.When gels were transferred onto nylon membranes, incubated with either apolyclonal (not shown) or monoclonal SBMV-C antibody and then developed withNBT-BCIP substrate, no differences were found in the migration of SBMV-C coatprotein (29 TWa) from cowpea or bean SBMV-C + SH1VIV-infected primary leaves(Fig. 31). The monoclonal antibody reacted with one other protein of110Fig. 30. SBMV-C-specific trapping of antigen from homogenates of infected tissue.A) Homogenate from “Bountiful” primary leaves doubly-inoculated with SBMV-Cand SHMV. Trapped particles have a diameter of approximately 18 nm. B)Homogenate from cowpea primary leaves doubly-inoculated with SBMV-C andSHMV. Trapped particles have a normal diameter of approximately 30 nm. C)Homogenate from healthy plants; no particles can be seen. (Bars represent 100nm).11140SS4*,‘I’0rLl4-$I0II.. .+ .E .99I97,40066,200.-45,0007’____31,00021,500..-”14,400.-”Fig. 31. Western blot analysis of SBMV-C coat protein migration from cowpea andbean primary inoculated leaves. Partially purified leaf proteins, extracted asdescribed in Material and Methods, were electrophoresed through 12% (w/v) SDSpolyacrylamide gel, transferred to a nylon membrane, reacted with anti-SBMV-Cspecific monoclonal antibody and developed as described in Materials andMethods. Molecular weight markers were stained with Coomassie blue. A proteinof approximately 32 KDa in both healthy and SBMV-B-infected bean reacted withmonoclonal antibody (upper arrow). SBMV-C coat protein migrated to the sameposition as SBMV-C coat protein from cowpea (lower arrow), indicating that thesize of the protein produced in SHMV + SBMV-C-infected primary bean leaveswas the same as that produced in SBMV-C-infected cowpea primary leaves.113approximately 32 KDa, present in healthy bean (Fig. 31) and cowpea tissue (notshown). The unexpected reaction of the monoclonal antibody with a protein inhealthy tissue could be due to the presence of a similar epitope, in an endogenousplant protein, to that against which the monoclonal was raised. The sameantibody did not react against SBMV-B coat protein (Fig. 31). These resultsindicate that the SBMV-C coat protein produced in SBMV-C-infected bean was thesame size as that produced by SBMV-C-infected cowpea.114Chapter 5DiscussionThe results of these experiments provide an example of a system inwhich one virus (SHMV) enables a second virus (SBMV-C) to move from cell to cellwithin the inoculated leaf of a plant that is not normally a host of SBMV-C.However, SHMV does not enable SBMV-C to move systemically through the plant.The cowpea strain of SBMV replicated only subliminally in bean, havingno effect on the appearance or development of inoculated plants. Conversely, whenSBMV-C and SHMV were inoculated either simultaneously or sequentially (atintervals not exceeding 24 h), SBMV-C replicated and produced large amounts ofinfectious material and, in bean, cultivar ‘Pinto”, caused characteristic necrotic pinpoint local lesions. It is interesting to note that the bean strain of SBMV, in singleinoculations, produced very similar local lesions in “Pinto” bean. This phenomenonseems to indicate that once SBMV-C concentration exceeds a certain threshold, itthen triggers the defence mechanism in bean in a dose-dependent fashion, in amanner similar to that for SBMV-B. The host-virus reaction appeared to besomewhat specific, since SHMV alone failed to induce these symptoms in spite of thefact that it reached high titres in the leaf. The specificity of this particular host-virusreaction was confirmed by the fact that, upon mixed inoculation with SBMV-C and115SHMV, the other three bean cultivars (“Top Crop”, “Tendergreen” and “Bountiful”),developed the same symptoms as those induced by single inoculation with SHMV.The observed increase of infectious SBMV-C in mixed infections withSHMV could be due to either complementation of some replication function, whichwould augment the susceptibility of infected cells, or to complementation of amovement factor/function which would allow the dependent virus to move intoneighbouring cells and replicate. The first hypothesis was tested by examining thebehaviour of SBMV-C in bean protoplasts. Results showed that SBMV-C replicatedand produced infective particles in bean protoplasts to the same degree as SBMV-B.This suggests that the limited recovery of SBMV-C from inoculated bean leaves wasnot due to the inability of the cells to support viral replication. Further experimentsshowed that SHMV had no effect on the accumulation of SBMV-C in beanprotoplasts. Thus, SHMV infection most likely augmented SBMV-C concentration inwhole leaves by complementation of a movement factor. This could be a plant andlorviral factor, since spread of a virus within a plant depends on both the genome of theplant (Zaitlin and Hull, 1987) and the translation products of the viral genome(Atabekov and Dorokhov, 1984). One possible factor is the “transport protein” whichis important in determining the host range of plant viruses (Taliansky et al, 1982b).Mixed infection of bean with SBMV-C and SII1V1V produced high levelsof SBMV-C antigen and infectious material only in inoculated primary leaves. Thus,116in the SBMV-C/SHMV system, the helper effect provided by SHMV was limited toshort distance (cell-to-cell) transport. This phenomenon was not temperature-dependent, since the same results were obtained with inoculated plants grown ateither 32 C or 25-28 C (Fuentes and Hamilton, 1988, 1991). Malyshenko et al (1989)obtained similar results in complementation experiments between different viruseswithin the tobamovirus group and between unrelated viruses from differenttaxonomic groups. The complementation of transport function was limited to spreadof the dependent virus only in the inoculated primary leaves.Comparison of the accumulation of SBMV-C and SBMV-B viral antigenin different parts of inoculated plants showed that, in mixed infections with S}{MV,SBMV-C moved at the same rate as SBMV-B between epidermal and parenchymacells of the leaf blade (Fig.8). However, the comparatively low quantities of SBMV-Cin the veinal system of inoculated leaves and its absence in petioles, stems, roots andtrifoliate leaves suggested that the virus was not being transferred from parenchyinainto the conducting system. These results were further confirmed when virusantigens were detected in thick leaf sections after transfer onto nitrocellulose paper.When tissue from cowpea inoculated with SBMV-C was compared to bean tissueinoculated with a mixture of SBMV-C and SHMV, it was evident that, in the cowpeatissue, viral antigen could be found in the mesophyll and the conducting bundle,whereas in the bean tissue SBMV-C could only be found in epidermal and mesophyllcells of the leaf (Figs.11 to 14). The absence of infectious material in tissues other117than those in the inoculated leaves was also confirmed by bioassay and Southern-blothybridization analysis (Fig.9).The fact that SHMV, in both single and mixed infection, movedsystemically in the host, but did not complement the long distance movement ofSBMV-C, implies that a specific virus-plant interaction regulates viral movementbetween parenchyma cells and vascular tissue. Recent reviews suggest that viralmovement is probably regulated at several levels by different host and viral genes(Atabekov and Taliansky, 1990; Hull, 1989). It is generally accepted that virusesmove through plasmodesmata (for reviews see Esau, 1968; Meshi and Okada, 1987;Zaitlin and Hull, 1987; Hull, 1989; Robards and Lucas, 1990) and plasmodesmatalconnections function differently in different symplastic domains (symplasm meaninga community ofliving plant cell protoplasts interconnected by plasmodesmata) (Erweeand Goodwin, 1985). Thus, it is likely that different plant and/or viral genes regulatecell-to-cell and long distance viral movement (eg. between parenchyma and vasculartissues).One mechanism to explain cell-to-cell movement, proposed by Citovskyet al (1990) and Citovsky and Zambryski (1991), is based on their finding that theTMV 30 KDa movement protein is both an RNA- and single-stranded DNA-bindingprotein. They proposed that cell-to-cell transport of plant viruses occurs by theformation of an unfolded nucleic acid-movement protein complex which is targeted118to the plasmodesmata. The movement protein then interacts with theplasmodesmata, changing their size-exclusion limits, thus allowing for translocationof the viral nucleic acid.Since the binding of RNA by the 30 KDa protein is non-specific (Citovskyet al, 1990), an analogous transport protein of SH1VW could bind SBMV-C single-stranded RNA and transport it from cell to cell. In fact, Malyshenko et al (1988)showed that SHMV facilitated cell-to-cell transport of the B component RNA of redclover mottle comovirus (RCMV), which does not encode either the transport 58KDa/48 KDa protein or the coat protein, in cowpea. This provides evidence that avirus may act as a helper for a viral RNA component which does not encode its owntransport or coat protein. On the other hand, a more recent study by Taliansky etal (1992) suggested that the presence of the full length TMV genome or a certainTMV-encoded product(s) other than the 30 KDa protein is required forcomplementation of the RCIVW-B RNA. Thus, the interaction of the movementprotein with the viral RNA and with the plasmodesmata are not enough to promoteviral movement.Another factor that the 30 TWa protein-RNA binding model does notaccount for is the relative host specificity of movement complementation. Whencowpea plants were simultaneously inoculated with SB1V]EV-B and SHMV,complementation of SB1VIV-B movement by SHMV did not occur. Thus, SHMV will119act as a helper to SBMV-C in bean but will have no effect in the reciprocal situationin cowpea; SHMV infects both hosts systemically. Citovsky and Zambrysky (1991)explain the relative specificity of viral RNA translocation as a result ofcompartmentalization of viral replication which would increase the probability of the30 KDa protein associating with viral nucleic acid rather than with host RNA. Thesame could be true for the interaction of the 30 TWa protein of a helper virus withthe RNA of a dependent virus. The probability of the 30 KDa protein of a helpervirus associating with the RNA of a dependent virus would be greater if both virusesreplicated in the same cell compartment. For example, compartmentalization ofreplication and translation of SBMV-C and SHMV may differ in bean and cowpea,with both viruses replicating in the same cell compartment in bean. This modelwould by no means explain all helper-dependent virus phenomena, since certain virusgroups such as comoviruses have different strategies for cell-to-cell movement(Wellink and van Kammen, 1989), and the nature of the infectious agent can bedifferent for different viruses (Maule, 1991).Repeated observations of thin sections, treated and stained as describedin Materials and Methods, failed to reveal the presence of intact SBMV-C virions inleaf tissue doubly infected with SHMV and SBMV-C (Fig. 19, 20 and 22). Results ofimmunosorbent electron microscopy ofleaf dips showed that although whole SBMV-Cparticles were present, they were fewer in number than those from SBMV-B-infectedbean and occasionally the antibody-coated grids were covered with particles of a120diameter corresponding to a T=1 configuration instead of the normal T=3 pattern(Fig. 30). Furthermore, sections from SBMV-C + SHMV-infected leaves labelled withgold revealed the presence of heavily labelled clumps or strands of amorphousmaterial, but no whole particles (Fig. 28 and 29). The presence of this amorphousmaterial, which appears to be viral, is correlated with the high levels of antigendetected by ELISA and suggests that SBMV-C virions are either unstable or fail toassemble efficiently in bean.Studies on the movement of free TMV RNA mutants or mutants withdeletions or insertions in the coat protein have shown that their cell-to-cell movementis just as efficient as wild type TMV, yet long distance spread occurs inefficiently andsporadically compared to that of the wild type (Siegel et al., 1962; Dawson et al,1988). Saito et al (1990), showed that a mutant with modifications in the origin ofassembly displayed reduced capacity for both long distance movement and assemblyin tobacco. On the other hand, a hybrid virus containing an exact exchange of thecoat protein open reading frame of odontoglossum ringspot virus (ORSV) for that ofTMV was able to replicate and efficiently move from cell to cell in tobacco leaves.However, although virions accumulated to a high level, the chimeric virus was unableto move systemically (Dawson and Hilf, 1992). Thus, it appears that, in some cases,a compatible capsid protein may be one of the factors required for virion entry intothe conductive tissue.121In the case of SBMV-C, several factors could be responsible for the lackof systemic movement of the virus in bean. One of these factors may be the absenceof the compatible capsid protein. Due to the absence of proper recognition sites in thecapsid protein of SBMV-C, it would be unable to interact with the host factor thatwould allow modification of the cell and subsequent movement of the virus into thevascular system. Since the bean strain of SBMV does move systemically in bean, oneway of investigating the role of the coat protein would be to replace the gene codingfor the SBMV-C coat protein with that from SBMV-B and then observe the behaviourof this hybrid in bean.A second cause of the lack of SBMV-C long-distance mobility in beancould be the fact that SBMV-C virions seem to be unstable and occur in lowconcentration in mixedly infected plants. Although gel electrophoresis of coatproteins of SBMV-C from bean plants and SBMV-C from cowpea plants showed nodifference in their migration (Fig. 31), and hence their molecular weight, results ofthe electron microscopy showed that whole virions were either in very lowconcentration, in an unstable configuration or completely absent. SBMV-C particlesare stabilized by three types of bond: 1) divalent cations; 2) pH-dependent contacts;and 3) salt linkages between protein and RNA (Hull, 1977). It is possible that inbean, one or several of these SBMV-C structural contacts is altered, resulting inunstable particles. SBMV could belong to the group ofviruses that requires assemblyofvirus particles or formation ofa specific ribonucleoprotein-complex for efficient long122distance movement, as occurs with TMV (Saito et al., 1990,) brome mosaic virus(Sacher and Ahiquist, 1989) and beet necrotic yellow vein virus (Quillet et eL, 1989).A third cause for the lack of systemic movement of SBMV-C in bean,which would not exclude either one of the mechanisms proposed above, could berelated to differences in the routes by which viruses move systemically. Gergerichand Scott (1988) examined the movement ofpurified virus particles in the xylem andsubsequent initiation of primary infection in non-wounded cells using a modificationof the steamed-stem techniques described by Caidwell (1930) and Schneider andWorley (1959). Beetle-transmissible viruses, such as SBMV-B, were translocatedthrough the steam-killed stem sections and initiated infection above the steam-killedarea in bean. Viruses not transmitted by beetles, such as SHMV., did not movethrough the steam-killed sections and were not able to infect non-wounded cells abovethe steam-killed sections. These results and those from previous work (Bennett,1940; Schneider and Worley, 1959; Roberts, 1970), would indicate that SBMV istranslocated mainly in the xylem while SHMV moves through the phloem. Thus,although SBMV-C is able to move from cell to cell with the aid of SHMV, once itreaches the cells neighbouring the conducting tissue, other requirements for itstranslocation are not met.The above observations would imply: 1) that the nature of the infectiousSBMV-C agent that moves from cell-to-cell in SBMV-C/SHMV infected bean is123probably different from that in long-distance movement; and 2) that these types ofmovement are distinct processes controlled by different factors. Long distancemovement may involve one or more interactions of host factors with viral products,different from, or in addition to, those involved in cell-to-cell movement. In the firstplace, loading of the mobile form of the virus from the mesophyll into the lumen ofthe sieve element may occur through plasmodesmata of the phloem parenchyma (inthe symplasm), or it could involve an apoplasmic step. In either case, it would seemthat the cellular control process which regulates the properties of plasmodesmata inphloem parenchyma is different from that of plasmodesmata in the mesophyll(Robards and Lucas, 1990). Thus, virus products other than, or in addition to, themovement protein, may be required to allow virai translocation into the conductingelements (Hull, 1989). It can be speculated that in addition to modifications of theplasmodesmata in the parenchyma cells surrounding the phloem, other virusproducts, such as the coat protein, interact with plant factors in order to overcome,or be protected against, the plant’s natural defences present in the vascular system.These plant factors could be components in the cell wall which bind to virus particlesor mobile forms and allow them to be released into the conducting stream. Theavailability of these complementary plant factors could be the result of a long processof co-evolution between a specific plant species and virus, and would determinewhether a particular plant is a host for a virus.In a preliminary study, BYMV, a potyvirus, and BRMV, a124comovirus, were found to act as helpers for SBMV-C replication in inoculated primaryleaves of “Bountiful” bean. These results, together with the results from other virus-host combinations for which complementation of the movement function has beendemonstrated (Taliansky et al., 1982 a,b; Carr & Kim, 1983; Barker, 1987,1989;Malyshenko et aL, 1987, 1988, 1989), have been interpreted to suggest that transportcomplementation is relatively non-specific (Taliansky et al. 1982 b, Atabekov andTaliansky, 1990). However, there are many examples for which complementationbetween viruses is not known to occur (Barker, 1989, Malyshenko et al, 1989). Threeviruses that infect bean systemically, alfalfa mosaic virus, tomato ringspot virus andbean golden mosaic virus, which were also tested as part of this study, failed tofacilitate SBMV-C movement in bean. Currently there is no simple explanation forthis phenomenon. Viruses have been grouped according to amino acid sequencesimilarities of their putative transport proteins, yet no correlation was found amongsequence similarities in transport proteins ofviruses and their ability to complementeach other (Melcher, 1990).This study provides a unique example in which the “helper effect” of onevirus is limited to cell-to-cell movement of the “dependent” virus in the inoculatedleaf. Thus, it allows for a clear separation between the processes of short and longdistance movement. This division, which has been widely used before to describeviral movement, supports the hypothesis that short and long-distance movement ofviruses require different mechanisms involving different viral and plant factors. One125of these factors in the SBMV-C/SHMV complementation system seems to be properassembly of the SBMV-C coat protein, which is apparently not required for cell-to-cellmovement complementation but presumably necessary for complementation of long-distance movement.Systemic invasion ofplants by viruses is usually accompanied by diseaseand consequent production losses (Matthews, 1991). Long-distance movement is ofprime importance since it is the main route for the establishment of a systemicinfection and is probably related to the efficiency ofvertical spread ofviruses throughseed transmission (Maule, 1991). A system such as the one described in this workcould be used to identify the plant factors involved in blocking long-distancemovement and those viral factors necessary for long distance movement. Aconsequence of the use of a model system such as this might be the development ofdisease-resistant crops.Understanding the nature of the interactions between viruses and plantfactors which affect the movement process is required in order to categorize andpredict the behaviour of viruses in mixed infctions. In this study, the interaction ofSHMV and SBMV-C in bean has been characterized to a large extent, thus allowingcomparison with other mixed virus infection combinations.126BibliographyAbad-Zapatero, C., Abdel-Meguid, S.S., Johnson, J.E., Leslie, A.G.W., Rayment, I.Rossmann, M.G., Suck, D., Tsukihara, T. 1980. Structure of southern beanmosaic virus at 2.8 A resolution. Nature (London), 286:33.Allison, A.V., Shalla, T.A. 1974. The ultrastructure of local lesions induced by potatovirus: a sequence of cytological events in the course of infection.Phytopathology 64:784-793.Atabekov, J.G., Dorokhov, Y.L. 1984. Plant virus-specific transport function andresistance of plants to viruses. Adv. Virus Res. 29:313-364.Atabekov, J.G., Taliansky, M.E. 1990. Expression of a plant-virus coded transportfunction of different viral genomes. Adv. Virus Res. 38:20 1-248.Atabekov, J.G., Taliansky, M.E., Drampyan, A.H., Kaplan, I.B., Turka, I.E. 1984.Systemic infection by a phloem-restricted virus in parenchyma cells in a mixedinfection. Biologicheskie Nauki 10:28-31.Barker, H. 1989. Specificity of the effect of sap-transmissible viruses in increasingthe accumulation of luteoviruses in co-infected plants. Ann. appl. Biol. 115:71-78.Barker, H. 1980. Superinfection of mesophyll protoplasts with viruses. In: Tissueculture methods for plant pathologists. D.S. Ingram and J.P.Helgeson, eds. pp103-107.Barker, H. 1987. Invasion of non-phloem tissue in Nicotiana clevelandii by potatoleafroll luteovirus is enhanced in plants also infected with potato Y-potyvirus.J. Gen. Virol. 68: 1223-1227.Beier, H., Suer, D.J., Russel, M.L., Bruening, G. 1977. Survey of susceptibility tocowpea mosaic virus among protoplasts and intact plants from Vigna sinensislines. Phytopathology 67:917-921.Beier, H, Bruening, G., Russel, M.L., Tucker, C.L. 1979. Replication of cowpeamosaic virus in protoplasts isolated from immune lines of cowpeas. Virology95:165-175.Bennett, C.W. 1940. Relation of food translocation to movement of virus of tobaccomosaic. J. Agric. Res. 60:361-390.Berryman, M. A., Rodewald, R.D. 1990. An enhanced method for post-embedding127immunocytochemical staining which preserves cell membranes. J. HistochemCytochem. 38:159-170.Blake, M.S., Johnston, K.H., Russell-Jones, G.J., Gotschlich, E.C. 1984. A rapid,sensitive method for detection ofalkaline phosphatase-conjugated anti-antibodyon Western blots. Anal. Biochem. 136:175-179.Caidwell, J. 1930. The physiology of virus diseases in plants. I. The movement ofmosaic in the tomato plant. Ann. of appi. Biol. 17:429-443.Carr, R.J., Kim, K.S. 1983. Evidence that bean golden mosaic virus invades nonphloem tissue in double infections with tobacco mosaic virus. J. Gen. Virol.64:2489-2492.Cassab, G.I., Varner, J.E. 1987. Immunocytolocalization of extensin in developingsoybean seed coats by immunogold-silver staining and by tissue printing onnitrocellulose paper. J. Cell Biol. 105:2581-2588.Cassells, A.C., Gatenby, A.A. 1975. The use of lessamine rhodamine B conjugateantibody for the detection of TMV antigen in tomato mesophyll protoplasts. Z.Naturforsch. 30:696-697.Citovsky, V., Knorr, D., Schuster, G., Zambryski, P. 1990. The P30 movement proteinof tobacco mosaic virus is single strand nucleic acid binding protein. Cell60:637-647.Citovsky, V., Knorr, D., Zambryski, P. 1991. Gene I, a potential movement locus ofCaMV, encodes and RNA binding protein. Proc. Nati. Acad. Sci. USA 88:2476-2480.Citovsky, V., Zambryski, P. 1991. How do plant virus nucleic acids move throughintercellular connections? BioEssays 8:373-379.Clark, M.F., Adams, A.N. 1977. Characteristics of the microplate method of enzyme-linked immunosorbent assay for detection of plant viruses. J. Gen. Virol.34:475-483.Clark, M.F., Lister, R.M., Bar-Joseph, M. 1986. ELISA techniques. MethodsEnzymol. 118:742-766.Cocking, E.C. 1966. An electron microscope study of the initial stages of infectionofisolated tomato fruit protoplasts by tobacco mosaic virus. Planta 68:206-2 14.Culver, J.N., A.G.C. Lindbeck, Dawson, W.O. 1991. Virus-host interactions:128Induction of chiorotic and necrotic responses in plants by tobamoviruses.Annu. Rev. Phytopathol. 29:193-217.Davis, P.B., Pearson, C.K. 1978. Characterization of density gradients prepared byfreezing and thawing a sucrose solution. Anal. Biochem. 91:343-349.Dawson, W.O., Bubrick, P., Grantham, G.L. 1988. Modifications of tobacco mosaicvirus coat protein gene affecting replication, movement and symptomatology.Phytopathology 78:783-789.Dawson, W.O., Hilf, M.E. 1992. Host range determinants of plant viruses. Annu.Rev. Plant Physiol. Plant Mol. Biol. 43:527-555.Deom, C.M., Oliver, M.J., Beachy, R.N. 1987. The 30-kilodalton gene product oftobacco mosaic virus potentiates virus movement. Science 237:389-394.Deom, C.M., Schubert, K., Wolf, S., Holt, C., Lucas, W.J., and Beachy, R.N. (1990).Molecular characterization and biological function of the movement protein oftobacco mosaic virus in transgenic plants. Proc. Natl. Acad. Sci. USA 87:3284-3288.Deom, C.M., Wolf,S., Holt, C.A., Lucas, W.J., Beachy, R.N. 1991. Altered functionof the tobacco mosaic virus movement protein in a hypersensitive host.Virology 180:251-256.Dodds, J.A., Hamilton, R.I. 1972. The influence of barley stripe mosaic virus in thereplication of tobacco mosaic virus inHordeum vulgare L. Virology 50:404-411.Dorokhov, Y.L., Alexandrova, N.M., Miroshnichenko, N.A., Atabekov, J.G. 1983.Isolation and analysis of virus-specific ribonucleoprotein of tobacco mosaicvirus-infected tobacco. Virology 127:237-252.Erwee, M.G., Goodwin, P.B. 1985. Symplast domains in extrastelar tissues ofEgeriadensa Planch. Planta 163:9-19.Esau, K. 1968. Viruses in Plant Hosts. Univ. Wisconsin Press. Madison, WI. 225 pp.Esau, K., Cronshaw, J., Hoefert, L.L. 1967. Relation of beet yellows to the phloemand to movement in the sieve tube. J. Cell Biol. 32:71-87.Flor, H.H. 1971. Current status of the gene-for-gene concept. Annu. Rev.Phytopathol. 9:275-296.Frens, G. 1973. Controlled nucleation for the regulation of the particle size inmonodisperse gold suspensions. Nature, Physical Science 241:20-22.129Fuentes, A.L., Hamilton, R.I. 1988. Spread of the cowpea strain of southern beanmosaic virus in a non permissive host is facilitated by infection with sunnhemp mosaic virus. Abstr. Tnt. Congr. Plant Pathol., 5th, Kyoto, Japan. 114Fuentes, A.L., Hamilton, R.I. 1991. Sunn-hemp mosaic virus facilitates cell-to-cellspread of southern bean mosaic virus in a nonpermissive host. Phytopathology81:1302-1305.Gardiner, W.E., Sunter, G., Brand, L., Elmer, J.S., Rogers, S.G., Bisaro, D.M. 1988.Genetic analysis of tomato golden mosaic virus: the coat protein is notrequired for systemic spread or symptom development. The EMBO J. 4:899-904.Gegenheimer, P. 1990. Guide to protein purification. In Methods in enzymology. VolVol 182. Academic Press, New York. ppl74-193.Gergerich, R.C., Scott, H.A. 1988. Evidence that virus translocation and virusinfection of non-wounded cells are associated with transmissibility by leaf-feeding beetles. J. Gen. Virol. 69:2935-2938.Ghabrial, S.A., Shepherd, R.J., Grogan, R.G. 1967. Chemical properties of threestrains of southern bean mosaic virus. Virology 33:17-27.Gibbs, A.J. 1976. Viruses and plasmodesmata. In: Intercellular studies in plants:studies on plasmodesmata, B.E.S. Gunning and A.W. Robards, eds. Springer-Verlag. Berlin. pp.149-164.Griffiths, G. 1984. Immunochemistry of Cryo Sections: A How-to Guide. SorvalApplications Brief NO. 9.Goodman, R.M. and Ross, A.F. 1974. Enhancement of PVX synthesis in doublyinfected tobacco occurs in doubly infected cells. Virology 58:16-24.Hacker, D.L., Petty, I.T.D., Wei, N., Morris, T.J. 1992. Turnip crinkle virus genesrequired for RNA replication and virus movement. Virology 186:1-8.Hamilton, R.I., Nichols, C. 1977. The influence of bromegrass mosaic virus on thereplication of tobacco mosaic virus in Hordeum vulgare. Phytopathology67:484-489.Hancock, K., Tsang, V.C.W. 1983. India ink staining of proteins on nitrocellulosepaper. Anal. Biochem. 133:157-162.Hatta, T., Francki, R.I.B. 1981. Identification of small polyhedral virus particles in130thin sections ofplant cells by an enzyme cytochemical technique. J. Ultrastruc.Res. 74:116-129.Horisberger, M., Rosser, J. 1977. Colloidal gold, a useful marker for transmissionand scanning electron microscopy. J. Histochem. Cytochem. 25:295-305.Hull, R. 1977. The stabilization of the particles of turnip rosette virus and of othermembers of the southern bean mosaic virus. Virology 79:58-66.Hull, R. 1989. The movement of viruses in plants. Annu. Rev. Phytopathol. 24:213-240.Ishimoto, M., Sano, Y., Makoto, K. 1990. Increase in cucumber mosaic virusconcentration in Japanese radish plants co-infected with turnip mosaic virus(II) electron microscopic and immunohistochemical observations. Ann.Phytopath. Soc. Japan 56:63-72.Johnson, D.A., Gautsch, J.W., Sportsman, J.R., and Elder, J. H. 1984. Improvedtechnique utilizing nonfat dry milk for analysis of proteins and nucleic acidstransferred to nitrocellulose. Gene Anal. Techn. 1:3-8.Kassanis, B., Varma, A. 1975. Sunn-hemp mosaic virus. No 153 in: Descriptions ofPlant Viruses. Commonw. Mycol. Inst.! Assoc. Appi. Biol., Kew, England.Knorr, D.A., Dawson, W.O. 1988. A point mutation in the tobacco mosaic capsidprotein gene induces hypersensitivity in Nicotiana sylvestris. Proc. Natl. Acad.Sci. USA 85:170-174.Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of thehead of bacteriophage T4. Nature (London) 227:680-685.Larkin, P.J. 1976. Purification and viability determinations of plants protoplasts.Planta (Berl.) 128:213-216.Leonard, D.A. and Zaitlin, M. 1982. A temperature-sensitive strain of tobaccomosaic virus defective in cell-to-cell movement generates an altered virus-codedprotein. Virology 117:416-424.Madore, M.A., Oross, J.W, Lucas, W.J. 1986. Symplastic transport in Ipomea tricolorsource leaves. Plant Physiol. 82:432.Malyshenko, S.I., Kondakova, O.A., Taliansky, M.E., Atabekov, J.G. 1989. Plantvirus transport function: complementation by helper viruses is non-specific.J. Gen. Virol. 70:2751-2757.131Malyshenko, S.I., Lapchic, L.G., Kondakova, O.A., Kuznetzova, L.L., Taliansky, M.E.,Atabekov, J.G. 1988. Red clover mottle comovirus spreads between cells intobamovirus-infected tissues. J. Gen. Virol. 69:407-412.Mang, K.Q., Ghosh, A., Kaesberg, P. 1982. A comparative study of the cowpea andbean strains of southern bean mosaic virus. Virology 116:264-274.Matthews, R.E.F. 1991. Plant Virology. 3rd ed. Academic Press. San Diego,California. 835 p.Maule, A.J. 1983. Infection of protoplasts from several Brassica species withcauliflower mosaic virus following inoculation using polyethylene glycol. J.Gen. Virol. 64:2655-2660.Maule, A.J. 1991. Virus movement in infected plants. Crit. Rev. Plant Sci. 9:457-474.Melcher, U. 1990. Similarities between putative transport proteins of plant viruses.J. Gen. Virol. 71:1009-1018.Meshi, T., Ohno, T., Okada, Y. 1982. Nucleotide sequence of the 30K protein cistronof cowpea strain of tobacco mosaic virus. Nucleic acids Res. 10:6111-6117.Meshi,T., Okada, Y. 1987. Systemic movement of viruses. In Plant-MicrobeInteractions: Molecular and Genetic Perspectives, ed. T. Kosuge, E. W. Nester.MacMillan. New York. pp 285-304.Meshi, T., Watanabe, Y., Saito, T., Sugimoto, A., Maeda, T., Okada, Y. 1989.Function of the 3Okd protein of tobacco mosaic virus: involvement in cell-to-cellmovement and clispensibility for replication. EMBO J. 6:2557-2563.Molefe, T.L., Wakarchuk, D.A., Kalmar, G., Hamilton, R.I. 1983. Interaction ofcowpea strains of southern bean mosaic and tobacco mosaic viruses in cowpeaand bean. Abstr. Int. Congr. Plant Pathol., 4th, Melbourne, Australia. 20.Nishiguchi, M., Motoyoshi, F. and Oshima, M. 1978. Behaviour of a temperaturesensitive strain of tobacco mosaic virus in tomato leaves and protoplasts. J.Gen. Virol. 39:53-61.Petty, I.T.D, Edwards, M.C., Jackson, A.O. 1990. Systemic movement of an RNAplant virus determined by a point substitution in a 5’ leader sequence. Proc.Natl. Acad. Sci. USA 87:8894-8897.Petty, I.T.D., Jackson, A.O. 1990. Mutational analysis of barley stripe mosaic virus132RNA f3. Virology 179:712-718.Ponz, F., Bruening, G. 1986. Mechanisms of resistance to plant viruses. Ann. Rev.Phytopathol. 24:355-381.Power, J.B., Cocking, E.C. 1969. A simple method for the isolation of very largenumbers of leafprotoplasts by using mixtures of cellulase and pectinase. Proc.Biochem. Soc. Biochem. J. 111:33.Quillet, L., Guilley, H., Jonard, G., Richards, K. 1989. In vitro synthesis ofbiologically active beet necrotic yellow vein virus RNA. Virology 172:293-301.Rigby, P., Dieckmann, W.J., Rhodes, M., Berg, P. 1977. Labelling deoxyribonucleicacid to high specific activity in vitro by nick translation with DNA polymeraseI. J. of Molecular Biol. 113:237-251.Robards, A.W., Lucas, W.J. 1990. Plasmodesmata. Annu. Rev. Plant Physiol. PlantMol. Biol. 41:369-419.Roberts, D.A. 1970. Viral infection of apparently uninjured leaves as influenced byparticle morphology and host species. Phytopathology 60: 1310 (Abstract).Sacher, R., Ahiquist, P. 1989. Effects of deletions in the N-terminal basic arm ofbrome mosaic virus coat protein on RNA packaging and systemic infection. J.Gen. Virol. 63:4545-4552Saito, T., Meshi, T., Takamatsu, N., Okada, Y. 1987. Coat gene sequence of tobaccomosaic virus encodes host response determinant. Proc. Natl. Acad. Sci. USA84:6074-6077.Schneider, I.R., Worley, J.F. 1956. Upward and downward transport of infectiousparticles of southern bean mosaic virus through steamed portions of beanstems. Virology 8:230-242.Saito, T., Yamanaka, K., Okada, Y. 1990. Long-distance movement and viralassembly of tobacco mosaic virus mutants. Virology 62:329-336.Siegel, A., Han, V., Montgomery, I., Koalacz, K. 1976. A messenger RNA for capsidprotein isolated from tobacco mosaic virus-infected tissue. Virology 73:363-37 1.Siegel, A., Zaitlin, M., Sehgal, O.P.. 1962. The isolation of defective tobacco mosaicvirus strains. Proc. Nati. Acad. Sci. USA 48:1845-1851.Slot, J.W., Geuze, H.J. 1981. Sizing of protein A-colloidal gold probes for133immunoelectron microscopy. J. Cell Biol. 90:533-536.Takamatsu, N., Ishiakwa, M., Meshi, T., Okada, Y. 1987. Expression of bacterialchloramphenicol acetyltransferase gene in to tobacco plants mediated by TMVRNA. EMBO J. 6:307-311.Takebe, I. 1977. Protoplasts in the study of plant virus replication. In:Comprehensive Virology. Vol II. H. Fraenkel-Conrat and R.R. Wagner, eds.Plenum Publishing, New York. pp 237-238.Takebe, I., Otsuki, Y. 1969. Infection of tobacco mesophyll protoplasts by tobaccomosaic virus. Proc. Natl. Acad. Sci. U.S.A. 64:843-848.Taliansky, M.E., Malyshenko, S.I., Kaplan, I.B., Kondakova, O.A., Atabekov, J.G.1992. Production of the tobacco mosaic virus (TMV) transport protein intransgenic plants is essential but insufficient for complementing foreign virustransport: a need for the full-length TMV genome or some other TMV-encodedproduct. J. of Gen. Virol. 73:47 1-474.Taliansky, M.E., Malyshenko, S.L, Pshennikova, E.S., Kaplan, I.B., Ulanova, E.F.,Atabekov, J.G. 1982a. Plant virus-specific transport function. I. Virus geneticcontrol required for systemic spread. Virology 122:318-326.Taliansky, M.E., Malyshenko, S.I., Pshermikova, E.S., Atabekov, J.G. 1982b. Plantvirus-specific transport function. II. A factor controlling virus host range.Virology 122:327-332.Tomenius, K., Clapham, D., Meshi, T. 1987. Localization by inimunogoldcytochemistry of the virus-coded 30K protein in plasmodesmata of leavesinfected with tobacco mosaic virus. Virology 160:363-371.Tremaine, J.H., Hamilton, R.I. 1983. Southern bean mosaic virus. No 274 in:Descriptions ofPlant Viruses. Commonw. Mycol. Inst./Assoc. Appl. Biol., Kew,England.Tremaine, J.H., Ronald, W.P., Kelly, E.M. 1981. Comparison ofhighly basic cyanogenbromide peptides from strains of southern bean mosaic virus. Can. J.Microbiol. 27:654-663.van Lent, J.W.M. 1988. Localization of viral antigens in leaf protoplasts and plantsby immunogold labelling. PhD Thesis. Wageningen Agricultural University.van Loon, L.C. 1983. Mechanisms of resistance in virus-infected plants. In: Thedynamics of host defence, ed. J.A. Bailey, B.J. Deverall. Academic. Sidney.134pp.123-190.van Loon, L.C. 1987. Disease induction by plant viruses. Adv. Virus Res. 33:205-255.Varma, A. 1985. Sunn-hemp mosaic virus. In: The plant viruses. Ed. M.H.V. vanRegenmortel & H. Fraenkel-Conrat. Plenum Press, New York. pp 249-266.Weintraub, M., Ragetli, H.W.J., Leung, E. 1976. Elongated virus particles inplasmodesmata. J. Ultra. Res. 56:35 1-364Wellink, J., van Kammen, A.B. 1989. Cell-to-cell transport of cowpea mosaic virusrequires both the 58K/48K proteins and the capsid proteins. J. Gen. Virol.70:2279-2286.Whitfeld, P.R., Higgins, T.J.V. 1976. Occurrence of short particles in beans infectedwith the cowpea strain of TMV. 1.Purification and characterization of shortparticles. Virology 71:471-485Wolf, S., Deom, C.M., Beachy, R.M., Lucas, W.J. 1989. Movement protein of tobaccomosaic virus modifies plasmodesmatal size exclusion limit. Science, 246:377-379.Wood, K.R. 1985. Tissue culture methods in phytopathology. I. Viruses. In: Plantcell culture: a practical approach. Ed. R.A. Dixon. IRL Press. Oxford, U.K.pp.61-62Wu, S., Rinehart, C.A., Kaesberg, P. 1987. Sequence and organization of southernbean mosaic virus genomic RNA. Virology 161:73-80.Zaitlin, M., Hull, R. 1987. Plant virus-host interactions. Annu. Rev. Plant Physiol.38:291-315.135


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