Iowa State University Capstones, Theses and Retrospective Theses and Dissertations Dissertations
1996 Biology of RPV barley yellow dwarf virus satellite RNA Lada Rasochova Iowa State University
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Biology of RPV barley yellow dwarf virus satellite RNA
by
Lada Rasochova
A dissertation submitted to the graduate faculty in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY
Major Molecular, Cellular and Developmental Biology
Major Professor W. Allen Miller
Iowa State University
Ames, Iowa
1996 XJMI Number: 9712592
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Lada Rasochova has met the dissertation requirements of Iowa State University
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TABLE OF CONTENTS
ABSTRACT v
CHAPTER I. GENERAL INTRODUCTION 1 Dissertation Organization 1 Introduction 1 Research Goals 8
CHAPTER! LITERATURE REVIEW 9 Luteoviruses 9 Qassification 9 Virion properties 9 Symptoms 12 Cytopathological effects 13 Tissue specificity 13 Host range 14 Serological relationships 15 Luteovirus transmission 15 Genome organization and replication 17 RNA agents associated with luteoviruses 21 Satellites 23 Classification 23 Requirements for replication of plant satellites 28 Helper virus specificity and host range of plant virus satellites 31 B type mRNA satellites 32 C ty^ linear satRNAs 33 D type circular satRNAs 35 Effects of satellites on disease symptoms 37 Possible mechanisms 37 Satellite viruses 41 B type mRNA satellites 41 C ty^ linear satRNAs 42 D ty^ circular satRNAs 46
CHAPTER 3. SATELLITE RNA OF BARLEY YELLOW DWARF-RPV VIRUS REDUCES ACCUMULATION OF RPV HELPER VIRUS RNA AND ATTENUATES RPV SYMPTOMS IN OATS 49 Abstract 49 Introduction 50 Results 52 Lack of mechanical transmission of satRPV RNA 52 Aphid transmission of satRPV RNA-containing virus from protoplasts to plants 53 SatRPV RNA affects helper virus RNA accumulation in oats but has no effect on symptoms caused by mixed infection of RPV andPAV 54 SatRPV RNA attenuates symptoms and decreases helper virus RNA accumulation in oats infected with pure RPV 55 SatRPV RNA decreases helper virus RNA accumulation in oat protoplasts 57 Discussion 56 Lack of mechanical transmission of satRPV RNA 57 iv
Helper vims titer is a limiting factor for satRPV RNA transmission by aphids from infected protoplasts extract 58 Lack of effect of satllPV I^A on symptoms caused by RPV+PAV 59 Possible mechanisms of reduction of virus symptoms severity and RNA accumulation 60 Acknowledgments 62 Materials and Methods 62 In vitro transcription 62 Whole plants: inoculation, virus propagation and purification, and extraction of virion and total RNAs 63 Protoplasts: inoculation, partial virus extraction, and total RNA isolation 63 RNA analysis 64 Aphid feeding on artificial membranes and inoculation of plants 64 References 65
CHAPTER 4. SPECLFICrrY OF RPV BARLEY YELLOW DWARF VIRUS SATELLrTE RNA IS ASSOCIATED WrrH POLYMERASE GENE OF SUBGROUP n LUTEOVIRUSES 76 Abstract 76 Introduction 77 Materials and Methods 79 In vitro transcription 79 Virus propagation, purification, and extraction of virion RNA 80 Electroporation of protoplasts and extraction of total RNA 80 Encapsidation assay 81 Purification of virions from tobacco protoplasts 82 Aphid feeding on artificial membranes, inoculation of plants, and isolation of total leaf RNA 82 RNA analysis 82 Results 83 Subgroup n BWYV supports satRPV RNA replication in tobacco cells 83 Subgroup I ST9a RNA failed to replicate satRPV RNA in tobacco protoplasts 84 SatRPV RNA is encapsidated in mostly circular form in BWYV capsids but as a linear form in B YDV particles 85 Aphid transmission of satRPV RNA from protoplasts to shepherd's purse plants required both BWYV and ST9a RNA 86 Effect of satRPV RNA on accumulation of BWYV and ST9a iRNA and disease symptoms in shepherd's purse plants 87 Discussion 88 Specificity of satRPV RNA - helper virus polymerase interaction 88 Effect of satRPV RNA on symptoms and viral RNA accumulation in plants 90 Host plant requirements of satRPV RNA 91 Acknowledgments 92 References 92
CHAPTER 5. GENERAL CONCLUSIONS 109 Conclusions 109 Recommendation for Future Research 111
REFERENCES CrrED 117 V
ABSTRACT Satellite RNAs (satRNAs) depend on their helper viruses for replication, encapsidation and spread. The goal of the research was to determine helper virus specificity and host range of the RPV barley yellow dwarf luteovirus (B YDV-RPV) satRNA (satRPV RNA) and assay satRPV RNA for the ability to affect the accumulation and modulate disease symptoms of its helper viruses in protoplasts and plants. Although similar in structural genes, subgroup I and n luteoviruses have very different polymerases. BYDV-RPV and beet western yellows virus
(BWYV), members of subgroup II luteoviruses, supported satRPV RNA replication m monocotyledonous and dicotyledonous hosts, respectively. In contrast, BYDV-PAV and ST9 associated RNA (ST9a RNA), subgroup I luteoviruses, failed to replicate satRPV RNA.
However, the stimulation of BWYV by ST9a RNA resulted in an increased accumulation of satRPV RNA progeny in BWYV + ST9a RNA + satRPV RNA inoculated tobacco protoplasts.
BWYV encapsidated satRPV RNA, but in a form different from that found in BYDV (RPV +
PAV) particles. SatRPV RNA was transmitted to plants by aphids that acquired virus from infected protoplasts. The concentration of encapsidated helper and satRPV RNA had to be above a threshold level to facilitate aphid transmission. Oat plants infected with BYDV-RPV and satRPV RNA had milder symptoms than those infected with BYDV-RPV alone. SatRPV
RNA reduced BYDV-RPV helper RNA accumulation in oat plants and protoplasts. However, it did not affect symptoms caused by the severe mixed infection of RPV and PAV BYDVs and had no effect on PAV RNA accumulation in oats. In contrast, satRPV RNA reduced the accumulation of both BWYV helper RNA and nonhelper ST9a RNA in shepherd's purse plants but attenuated BWYV and ST9a RNA symptoms only slightly. SatRPV RNA symptom modulation seems to be determined by competition between the satRPV RNA and its helper virus for replication and encapsidation. The results showed that satRPV RNA can replicate in both monocotyledonous and dicotyledonous protoplasts and plants and suggested that the vi specificity determinants of satRPV RNA replication are contained within the polymerase genes of supporting viruses rather than in structural genes or host plants. 1
CHAPTER 1. GENERAL INTRODUCTION
Dissertation Organization
This dissertation contains two manuscripts. The manuscript "Satellite RNA of barley yellow dwarf RPV virus reduces accumulation of RPV helper virus RNA and attenuates RPV symptoms in oats" is formatted for publication in Molecular Plant-Microbe Interactions and was published in a shorter version. The manuscript "Specificity of RPV barley yellow dwarf virus satellite RNA replication is associated with polymerase gene of subgroup n luteoviruses" is formatted and will be submitted for publication in Virology. I have been principally involved in the research (i.e., experimental design, performing all the experiments, data analysis and interpretation) and preparation of both the manuscripts. The roles of co-authors on the second manuscript were following: Boni Passmore taught me the techniques involving tobacco protoplasts isolation and total and encapsidated RNA analysis. Bryce Falk showed me the techniques used in the aphid transmission experiments, photographed plants, observed symptoms and collected plant tissue for subsequent RNA analyses performed by me. W. Allen
Miller was involved in the experimental design, interpretation of results and preparation of manuscripts. The dissertation also includes the general introduction chapter, in which I state the research problem and address the background and significance of the research topic; literature review chapter, and general conclusion chapter in which I summarize the results and make suggestions for additional research. The references cited in these three chapters are placed at the end of the dissertation.
Introduction
Satellite RNAs (satRNAs) are defined as sub-viral agents composed of RNA molecules which do not encode a capsid protein and depend for their productive multiplication on co- 2 infection of a host cell with a helper virus (Mayo et al., 1995). SatRNAs are encapsidated by the coat proteins of their helper viruses, either separately or in association with the RNA(s) of the virus and such particles are antigenically identical to those of the helper virus (Francki,
1985). Thus, satRNAs differ firam satellite viruses, which encode their own coat protein and form particles that are antigenically and usually morphologically distinct from virions of their helper viruses (Mayo et al., 1995). SatRNAs have substantially distinct nucleotide sequences from genomes of either helper virus or host and thus differ from defective interfering RNAs that are fully derived from their helper viruses (Mayo et al., 1995). The Qrst satRNAs were discovered because of their effect on disease symptoms. Depending on the satRNA-helper virus-host plant combination, some satRNAs can attenuate, exacerbate or produce new symptoms that are not associated with helper virus alone (Collmer and Howell, 1992). The features of the helper virus that allow it to replicate certain satRNA are not well defined for most satRNAs (Andriessen et al., 1995). Nothing is known about the host plant components required for satRNA replication and very littie about the interactions between the satRNA, helper vims and host required for satRNA to modulate symptom development (Masuta et al.,
1993; Sleat et al., 1994; Kong et al., 1995). The object of this research is the satellite RNA of RPV barley yellow dwarf virus
(satRPV RNA). It is a small (322 nt. Figure 1) noncoding RNA that was discovered in the process of cloning and sequencing of the RPV barley yellow dwarf luteovirus (BYDV-RPV;
Miller et al., 1991). The RNA was abundant in plants infected with an Australian isolate of
BYDV-RPV and in the mixture of RPV and PAV BYDVs. It has no significant sequence similarity to BYDV-RPV or BYDV-PAV genomic RNA and is encapsidated within BYDV-
RPV capsids predominantly as a linear monomer (Miller et al., 1991). Circular and multimeric forms of satRPV RNA were also detected in infected tissues. Isolated multimers can self- cleave into monomers in vitro in a protein-free reaction. Sequences that are involved in the formation of hammerhead ribozyme and responsible for self-cleavage were found in both (+) 3
ACAGAGCGCGUACUGUCUGACGACGUAUCCGCGCGGACUAGAAGGCUGGUGC 10 20 30 40 50
CUCGUCCAACAAAUAGAUACAGAAAUCCACCGAAGUAAAGAUCUCCAAUUGUG 60 70 80 90 100
GCACCACCAGGUGGCCACCACUCUUUGAAGUGAGGAGACUUGCUUUACGUGU 110 120 130 140 150
UUGUUCAGCCCGAGCUUUCGCUCGCACUGGAACACUGGUGUUUCGUCCUU 160 170 180 190 200
UCGGACUCAUCAGUCAAGGUACGCACCUUGA'GACACCGGGAAACAAUCGAUCA 210 220 230 240 250 260
AUCUUUCACAGAGCAACGAGUUCGCUACUCXJUGCAAAAGAUCGACUUCCUAUU 270 280 290 300 310
UCGUGGAUA 320
Figure 1. Nucleotide sequence of satRPV RNA. Sequences comprising proposed self- cleavage structures of the minus strand (complementary to bases 194-244) and plus strand
(bases 310-389) are underlined. Minus strand cleavage occurs between the complements of bases 238 and 239 (apostrophe). 4 and (-) strands of satRPV RNA (Miller and Silver, 1991). The (+) strand hammerhead is an unusual derivation from a consensus ribozyme. It can form an additional base-pairing which could generate a slow cleaving pseudoknot. The biological role of the altemative structures of satRPV RNA is unknown. Miller and Silver (1991) proposed that the formation of altemative stmctures serves as a molecular switch between the self-cleaving hammerhead and the inactive pseudoknot structure that may perform other fimctions in the satRPV RNA life cycle including an origin of replication, an origin of assembly, circularization of linear monomers, and other functions.
We demonstrated that satRPV RNA meets the definition of a true satRNA (Silver et al., 1994). We electroporated infectious satRPV RNA transcripts (Figure 2) into oat protoplasts in the presence or absence of BYDV-RPV RNA and showed that satRPV RNA depends on
BYDV-RPV genomic RNA for replication but BYDV-RPV replicates independently of satRPV
RNA. However, BYDV-PAV did not support satRPV RNA replication. We also demonstrated that satRPV RNA replicates by a symmetrical rolling circle (Figure 3B) during which the (+) strand linear monomers found in virions are first circularized. Circular monomeric forms of satRPV RNA are then copied by an RNA dependent RNA polymerase to give rise to multimeric (-) strand RNA. Newly foraied (-) strand multimers are processed to linear monomers. The processing occurs by self-cleavage. The linear (-) strand RNA monomers are then circularized and copied to produce (+) strand multimers which are processed into unit-lcagth linear RNAs encapsidated in virus particles. Linear monomeric and multimeric replication intermediates of both strands (Silver et al., 1994) and (+) strand circular monomers (Miller et al., 1991), which are formed during a symmetrical rolling circle, were detected in satRPV RNA progeny. The (-) strand circular monomers could not be identified because ±e gel system used in the experiment (denaturing 1% agarose) does not allow resolution of linear and circular fonns of satRPV RNA (Silver et al., 1994). The replication mechanism of satRPV RNA resembles that of satRNA of tobacco ringspot virus (satTRSV 5
A
pT7Sat
B
Self-cleaved transcript:
nts: 269 3 22 122
Figure 2. Map of permuted dimeric clone (pT7Sat) and transcripts of satRPV RNA.
(A) Map of satRPV RNA sequence in tiie transcription vector, pTTSat. Shaded boxes indicate satRPV RNA dimerjoined at 5g/I site (B/S). Large arrows indicate cleavage sites. (B)
Expected RNAs were obtained after self-cleavage of (+) sense RNA derived by transcription of
EcoRl -cut pT7Sat with T7 polymerase (begiiming at right-angled arrow in panel A). The boldest line indicates the 322-nt infectious monomer. Abbreviations: H = MndDI; S = SauSAl; B = BglR; R = EcoRI. 6
(-)
B
|(-) I self-cleavage virus (-) particle |(+)| self-cleavage i (-) © |(+) I self-cleavage
Figure 3. Rolling circle mechanism of satellite RNA replication (modified from Symons,
1991). Linear (+) monomeric RNA circularized and is copied to produce multimeric (-) strand which acts as a template directly (A), or self-cleaves, circularizes and is copied to produce multimeric (+) strand (B). After self-cleavage (+) strand linear or circular monomeric RNA is encapsidated. 7
RNA; Bruening et al., 1991), viroid-Iike satellite of lucerne transient streak virus (satLTSV RNA; Foster and Symons, 1987), avocado sunblotch viroid (Hutchins et al., 1986) and
carnation stunt associated RNA (Hernandez et al., 1992), where both (+) and (-) RNAs self-
cleave and the circular (-) RNAs act as templates for (+) RNA transcription and vice versa. In
contrast, the satRPV RNA replication differs from an asymmetrical model proposed for satellite
RNA of velvet tobacco mottie virus (VTMoV) and subterranean clover mottle virus (SCMoV;
Hutchins et al., 1985; Davies et al., 1990) in which the multimeric (-) strand is not processed
into monomers, but is copied to give multimeric (+) strands which then self-cleave (Figure
3A). SatRPV RNA is the only known satellite of luteoviruses. Luteoviruses can cause significant losses in a variety of crops in many countries of the world (Rochow and Duffus, 1981). BYDV is the most widespread of all cereal viruses. It is the most economically important virus of wheat, barley and oats (D'Arcy, 1995). BYDV has been estimated to cause annual losses of 1-3% in the United States (Dufflis, 1977) and in the midwest, it may cause greater losses than any other disease in oats. The prevalence and severity of BYDV in oats has increased in recent years, despite the use of more tolerant oat varieties. Other luteoviruses infect a wide range of dicotyledonous crop plants. For example, beet western yellows luteovirus (BWYV) is a serious pathogen in sugar beet, lettuce, spinach and many other dicotyledonous crops (Rochow and Duffus, 1981). Overall, the reduction in yield caused by luteoviruses has a significant economic value. Before I started my project, nothing was known about the effect of satRPV RNA on symptom induced by BYDV-RPV or mixmre of RPV and PAV BYDVs. No information was available about the helper virus requirements for satRPV RNA replication, effects of satRPV
RNA on helper virus replication and accumulation and helper virus or host plant range of satRPV RNA. The research involving an RNA associated with luteoviruses is directiy relevant to luteovirus biology because the identification of elements which are required for satRNA 8
replication can contribute to better understanding of virus replication in general and perhaps
lead to new means of controlling viral diseases.
Research Goals
Long-term goals of the research involving satRPV RNA are to fully understand the
complicated three-way relationship between satRNA, helper virus, and host plant. This
includes defining the essential fimctions required for satRNA replication that are supplied by
(i) the satRNA itself,
(ii) the helper virus, and (iii) the host plant. The goals also include determining the effects of satRNA on replication and/or accumulation of
its helper virus(es), disease symptoms, and understanding the mechanisms behind these effects.
My project addressed two specific objectives. The first objective was to determine the
effect of satRPV RNA on BYDV (RPV ± PAV) RNA replication in oat protoplasts, deliver
satRPV RNA to plants and determine the effect of satRPV RNA on viral RNAs replication,
virus accumulation and disease symptoms caused by BYDV in oats. The second objective was
to determine the range of helper viruses and, to some extent, host plants that allow the replication of satRPV RNA and to study the effects of satRPV RNA on replication and/or accumulation of these viruses and symptom development in divergent monocotyledonous and dicotyledonous hosts. 9
CHAPTER 2. LITERATURE REVIEW
Luteoviruses
Classification Luteoviruses are phloem-limited, obligately aphid transmitted plant viruses. The name
luteovirus was derived from Latin luteus, "yellow", as the infection with viruses of this group
induces yellowing symptoms in plants (Shepherd et al., 1976). Luteoviruses have been classified on the basis of symptoms, host range, serological relationships, aphid vector specificity, cytopathological effects, and genome organization into two subgroups (Miller, 1994). BCnown luteoviruses, divided into two subgroups, and their most important vectors are listed in Table 1. A list of tentative species is also shown (Randies and Rathjen, 1995).
Virion properties
Virions of luteoviruses are 25 to 30 nm in diameter, hexagonal in outline with no envelope or surface features. They contain no lipids or carbohydrates (Randies and Rathjen, 1995).
Virions exhibit icosahedral symmetry (T=3) and contain 180 subunits of a 22 to 23 kDa coat
protein. A few copies of the coat protein may contain an additional polypeptide fiised to carboxy-terminus of the coat protein. This polypeptide is produced by translational readthrough of the coat protein stop codon (Brown et al., 1996). Particles S20W is 104-127
(Randies and Rathjen, 1995). Virions are moderately stable, survive freezing and thawing, and are insensitive to chloroform and non-ionic detergents. The thermal inactivation points range from 65 to 70 ^C (Rochow, 1970). The ability to use frozen tissue makes it possible to overcome some of the disadvantages of low virus concentration in infected plants by accumulating frozen tissue. It also aids purification and facilitates serological tests of crude samples (Rochow and Duffiis, 1981). The virions are 21 to 28% RNA by weight (Rochow, 1970). 10
Table 1. List of luteoviruses and their major vectors (Miller, 1994; Randies and Rathjen, 1995)
Virus Abbreviation Vector
Subgroup I barley yellow dwarf-MAV BYDV-MAV Sitobion avenae barley yellow dwarf-PAV BYDV-PAV Rhopalosiphum padi, S. avenae barley yellow dwarf-SGV BYDV-SGV Schizaphis graminum soybean dwarf SDV Acyrthosiphon solani subterranean clover redleaf SCRLV A. solani
Subgroup n barley yellow dwarf-RPV BYDV-RPV R. padi barley yellow dwarf-ElMV BYDV-RMV R. maidis barley yellow dwarf-RGV BYDV-RGV R. padi beet westem yellows BWYV Myzus persicae potato leafroll PLRV M. persicae tobacco necrotic dwarf TNDV M. persicae tomato yellow top ToYTV Macrosiphum euphorbiae
Subgroup uncertain bean leaf roll BLRV A. pisum carrot red leaf CRLV Cavariella uegogpodii groundnut rosette assistor CRAY Aphis craccivora
Indonesian soybean dwarf ISDV Aphis glycines
Solanum yellows SYV M. persicae 11
Table 1. (Continued)
Virus Abbreviation Vector
Tentative Species beet yellow net BYNV celery yellow spot CeYSV chickpea stunt CpSV cotton anthocyanosis CAY grapevine ajinashika GAY millett red leaf MRLY
Physalis mild chlorosis PhyMCY Physalis vein blotch PhyYBY raspberry leaf curl RLCY tobacco vein distorting TYDY tobacco yellow net TYNY tobacco yellow vein assistor TYYAY 12
Symptoms Symptoms induced by luteoviruses are often difficult to distinguish from symptoms caused by other pathogens or from nutritional disorders. Characteristic features of luteoviral infection include stunting of infected plants, yellowing, reddening, rolling, and brittieness of infected leaves (Rochow and Duffus, 1981). Symptoms vary with the plant species, crop variety, the age and physiological condition of the plant at the time the infection occurs
(Rochow and Duffus, 1981). For example, BYDV induces stunting in all its hosts and can cause yellowing in some lines of barley, bright reddening of leaf tips in oats, com and other grasses. Leaf elongation and initiation may be inhibited, tillering reduced and heading suppressed in infected plants. BYDV infection results in marked increase in sterile florets in oats and reduced kernel weight (D'Arcy, 1995). Beet western yellows (BWYV) induces bright yellowing of leaves on sugar beets, lettuce and spinach. Symptoms induced by most luteoviruses including BYDV and BWYV are usually relatively mild. The severity of symptoms is usually greatest when plants are infected young.
Environmental factors also affect the expression of symptoms. Temperature and light intensity are critical for symptom development (Rochow and Duffus, 1981). Most severe symptoms usually appear in plants grown in conditions of high Ught intensity and cool temperature. Another factor involved may be the concentration of virus inoculated to plants (D'Arcy, 1995). Increasing the number of virus-transmitting aphids feeding on a host plant often leads to increased severity of symptoms and greater yield loss whereas increasing the number of virus-free aphids has Uttle or no effect.
The extemal symptoms are basically indirect, secondary expression of phloem collapse.
Phloem necroses spread from infected sieve elements, and cause symptoms by reducing translocation that results in carbohydrate accumulation, slowing plant growth, inducing loss of chlorophyll and inhibiting photosynthesis by feedback (Jensen and D'Arcy, 1995; Randies and
Rathjen, 1995). Luteoviruses do not change organization of phloem but kill cells in phloem. 13
Cytopathological effects
The effects of luteoviral infection on ultrastructure of the host cells have been determined for BYDVs and BWYV. It seems that subgroup I and II luteoviruses cause different structural changes in infected cells (Gill and Chong, 1979). These include:
(i) Interaction with the nucleus:
Subgroup I luteoviruses show extreme distortion of the nucleus, aggregation and
accumulation of densely staining, heterochromatin-like material. Subgroup H-
infected cells have relatively normal nuclei at first, until heterochromatin slowly
disintegrates. (ii) Site of the first occurrence of virus progeny:
New virus particles of subgroup I luteoviruses accumulate in cytoplasm, those of subgroup n around nucleus.
(iii) Appearance of membranes, vesicles and fibrils:
Single-membraned vesicles containing fibrils are found in the cytoplasm of
subgroup I-infected cells but double-membraned vesicles in cells infected with subgroup n luteoviruses.
Tissue specificity
A characteristic feature of luteoviruses is their restriction to phloem tissue. Virions have been found in phloem parenchyma, companion cells, and sieve elements (Waterhouse et al., 1988). Moreover, luteoviruses can be transmitted by aphids but not by manual inoculation of sap. Therefore, it is possible that:
(i) the conditions necessary for luteoviruses to multiply are found within phloem cells only, or
(ii) viruses are unable to move from cell-to-cell but can spread along phloem sieve elements only. 14
However, it has been demonstrated that mesophyll, epidermis, xylem, and undifferentiated cells can be infected with luteoviruses (Young et al., 1989; Dinesh-Kumar et al., 1992; Veidt
et al., 1992; Silver et al., 1994). Moreover, some replication of PLRV in mesophyll cells was
detected in Nicotiana clevelandii infected with PLRV and potato virus Y (Barker, 1989).
Double infection with PAV-Iike and RPV -like BYDVs was also reported to result in
breakdown of tissue specificity and invasion of xylem (Jensen and D'Arcy, 1995). Therefore,
the tissue specificity of luteoviruses is not likely caused by the inability of luteoviruses to
replicate outside the phloem but rather results from their inability to move from cell-to-cell.
Host range
Most luteoviruses have natural host ranges largely restricted to one plant family. For example, hosts of BYDV and SDV are confined to the family Poaceae and Leguminosae,
respectively (Mayo and Ziegler-Graff, 1995). However, hosts of BWYV include plant species
from 23 dicotyledonous families (Rochow and Duffris, 1981). Since luteoviruses are
transmitted by aphids only, host range restrictions can reflect the host range of the aphid vector
rather than that of the virus. For example, Myzus persicae, the major vector of BWYV, is
occasionally able to transmit BWYV to oats under laboratory conditions (Duffus and Rochow,
1978; Rochow and Duffus, 1978) but such transmission was not reported to occur in the field. On the contrary, BWYV and PLRV which have the same aphid vector, Myzus persicae, infect different host plants. Moreover, none of the shepherd's purse plants which are hosts of
BWYV could be infected with BYDV using Myzus persicae, Rhopalosiphum padi or Sitobion
avenae as vectors although these aphids were able to transmit BYDV to oats (Rochow and
Duffus, 1978). Therefore, the host range of luteoviruses is not simply determined by the range
of the vector. The host determinants of the virus infectivity are virtually unknown. 15
Serological relationships Luteoviruses are strongly immunogenic. All are serologically related to some degree and based on their cross-reactivity can be clustered into the following groups (Waterhouse et al., 1988; Martin and D'Arcy, 1995):
(i) BYDV-MAV, BYDV-PAV, BYDV-SGV;
(u) BYDV-RPV, BYDV-RMV, BYDV-RGV;
(iii) SDV, SCRLV, BLRV; and
(iv) TNDV,PLRV,andToYTV.
BWYV cross-reacts with nearly all luteoviruses at least slightly.
Luteovinis transmission
Luteoviruses are transmitted in a circulative non-propagative maimer by aphids
(Randies and Rathjen, 1995). Most luteoviruses have a single major vector but, for example,
BYDV-PAV is transmitted efficiendy by Rhopalosiphum padi and Sitobion avenae (Table 1).
In addition, the heterologous encapsidation in mixed infections known to occur among luteoviruses can allow transmission of one virus by the vector of another (Wen and Lister,
1991; Rochov, 1970). The mechanism of virus transmission and factors influencing transmission efficiency have been examined (Duffiis and Gold, 1967; Gray et al., 1991; van den Heuvel et al., 1991; Gildow, 1993; Gildow and Gray, 1993; van den Heuvel et al., 1993;
Jolly and Mayo, 1994; van den Heuvel et al., 1994; Brault et al., 1995; Power and Gray,
1995; Garret et al., 1996; Chay et al., 1996). The virus is acquired by phloem feeding and accumulates in the hindgut of the aphid. It is transported across the gut lining into the hemocoel and then crosses into the accessory salivary glands of the aphid. The virus exits the aphid along with salivary secretions via salivary duct during the feeding. The particles cross both the hindgut cells and the accessory salivary gland membranes via coated vesicles and the passage occurs by receptor-mediated endocytosis (Gildow, 1993; Gildow and Gray, 1993). In 16
addition, the virus must move across the basal lamina surrounding the accessory salivary glands. The mechanism of this transport is unknown. The basal lamina may selectively filter viruses or it may simply concentrate virions and thus increase the efficiency of transport into the accessory salivary glands. Vector specificity of luteoviruses is probably determined by some features of their protein coats (Mark Young, pers. comm.). It is thought that different domains of both coat and readthrough proteins interact with putative receptors on the aphid membranes.
Multiple factors have been shown to affect the efficiency by which aphids can transmit different luteoviruses (Power and Gray, 1995). These include environmental factors (temperature), biotype, developmental stage, morphological form of a single aphid species and others. The length of acquisition and inoculation access periods required for successful transmission varies among different virus-vector combinations but it has to be at least 15 min
(Gray et al., 1991). The efficiency of transmission is gready increased when acquisition and inoculation access times increase to approximately 24 to 48 hrs (Power and Gray, 1995).
There is a latent period of 10 to 48 hrs after acquisition. The vector continues to transmit the virus for many days and retains the ability to transmit after the moulting. However, the virus does not multiply in its vector and is not transmitted to the progeny of the aphid (Power and
Gray, 1995). The effect of virus concentration on transmission efficiency was studied using aphid feeding on preparations of purified virus through artificial membranes. The positive correlation between the virus titer and transmission efficiency was reported for BYDV (Pereira and Lister, 1989) and PLRV (van den Heuvel et al., 1991). For example, Pereira et al. (1989) reported no transmission of BYDV-PAV at the virus concentration of 5 ng/ml but 50% transmission when the titer of BYDV-PAV was increased to 70 (ig/ml. 17
Genome organization and replication Virions of luteoviruses contain a single molecule of infectious, linear, positive sense single-stranded RNA (Miller et al., 1995; Mayo and Ziegler-Graff, 1996). The genome size is fairly uniform among the members and range from 5,641 for BWYV to 5,982 nt for PLRV
(Mayo and Ziegler-Graff, 1996). A small protein (VPg) covalently linked to the 5' end of the genomic RNAs (gRNAs) has been reported for PLRV (9 kDa; Mayo et al., 1982) and BYDV-
EIPV (17 kDa; Murphy et al., 1989). No VPg was detected at the 5' end of BYDV-PAV (Ed
Allen, pers. comm.). However, it is not yet clear if other luteoviruses also possess a VPg.
There is no 3'-terminal poly(A) track or a tRNA-like structure in any luteovirai genome (Miller,
1994).
Luteoviruses contain 5 or 6 open reading frames (ORFs) which encode proteins of between 4 and 72 kDa (Randies and Rathjen, 1995). The organizations of ORFs on genomic
RNAs (gRNAs) of subgroup I and n luteoviruses are shown in Figure 4. The sizes of ORFs of different luteoviruses and possible functions of their products are listed in Table 2.
Luteoviruses use many different strategies to express their genes (Miller et al., 1995; Mayo and
Ziegler-Graff, 1996). These include: (i) production of subgenomic RNAs (sgRNAs) to express downstream ORFs;
(ii) translational frmneshift between the overlapping ORFs; (iii) leaky scanning by ribosomes to translate ORFs downstream of the first AUG;
(iv) readthrough of termination codons to express downstream ORFs as fusion
proteins;
(v) cap independent translation; and possibly
(vi) proteolysis of a precursor protein to produce more than one product from one
ORF. Genome organization of subgroup I luteoviruses differs from that of subgroup n (Figure 4).
Subgroup I luteoviruses contain a much longer nucleotide sequence 3' of the ORF 18
I Subgroup I rfitaa- m ' gRNA ' agRNAI •gRNA2 ST9a RNA *gRNA3 & ams::-; w gRHA
Subgroup II 66.^icee-i ^ aac cS^ irx-^poLX^ »• sn Figure 4. Diagram of the genome organization of the luteovirus subgroups. Boxes indicate ORFs, numbered as in Martin et al. (1990), with coding capacity shown in kilodaltons (K). Black-shaded ORFs are conserved between subgroups. CP indicates coat protein gene, POL indicates putative polymerase gene. Cross-hatched POL ORFs have homology to Carmovirales. Stripped POL ORF are homologous to Sobemovirales. Unshaded ORFs have no significant similarity to ORFs of any other virus, with the exception of protease motif in ORFl of subgroup n. Known positions of subgenomic RNAs are shown below the genomes. 19 Table 2. Proteins encoded by different luteoviruses with their sizes (kDa) and possible functions (modified from Randes and Rathjen, 1995) ORF BYDV BYDV PLRY BWYY BYDY SDY Function MAY PAY RPV — 0 — 28 29 29 — unknown 1 39 39 70 66 71 40 unknown 2 61 60 69 70 72 59 putative RdRp 3 22 22 23 23 22 22 coat protein 4 17 17 17 20 17 21 putative movement protein 5 51 43 56 52 50 48 put. transmission factor 6 4 7 — — — — unknown 5. This sequence is about 600 to 867 nt long for subgroup I viruses but only about 150 nt long for viruses of subgroup n (Miller et al., 1995). In subgroup I luteoviruses this sequence at 3" end may encode a 4 to 7 kDa ORF 6 of an unknown function. Such ORF was identified in BYDV-PAV and BYDV-MAV but never in members of subgroup II (Mayo and Ziegler-Graff, 1996). In contrast, subgroup n luteoviruses have ORF 0 at the 5' end of the genome that is absent in subgroup I luteoviruses (Miller et al., 1995). The function of PO is unknown and there have been no reports of detection of this protein in vivo. Little significant homology has been found between PO proteins of subgroup n luteoviruses (Mayo and Ziegler-Graff, 1995). PO has been shown to be dispensable for infectivity (Veidt et al., 1992). Potato plants expressing PO of PLRV had disease-like symptoms which suggests that PO may be involved in symptom expression (van der Wilk et al., 1993). It has also been speculated that PO determines host range of a particular luteovirus (Veidt et al., 1992). 20 All luteoviruses have putative RNA-dependent RNA polymerase (RdRp) that is encoded by ORF 2 in both subgroups. ORF 2 is expressed as a fusion with ORF 1 by ribosomal frameshifting (Di et al., 1993). P2 contains a conserved amino acid motif GXXXTXXXN(X25-40)GDD that is present in all known RdRps and thus can be found in all RNA viruses (Koonin, 1991). The polymerase sequences are very divergent between the subgroups. Subgroup I polymerases are similar to those of the diantho-, carmo-, tombus-, and necroviruses which are classified as Carmovirales (Koonin and Dolja, 1993). In contrast, subgroup n polymerases are more similar in amino acid sequence to those of sobemoviruses and therefore are placed in the order Sobemovirales (Koonin and Dolja, 1993). Habili and Symons (1989) identified possible helicase motifs in PI and P2 of SDV and other luteoviruses of subgroup I. The PI sequences of all subgroup n luteoviruses contain motifs characteristic of chymotrypsin-like serine proteases (Miller et al., 1995) but no proteolytic activity has been described for any luteoviral proteins. However, the replication proteins of subgroup n luteoviruses are similar to those of viruses containing sequence domains VPg-protease- polymerase. Thus, proteolytic activity of PI in subgroup n luteoviruses may exist. Moreover, the VPg of subgroup II luteoviruses may be generated by proteolytic cleavage in PI (Miller et al., 1995; Mayo and Ziegler-Graff, 1995). ORFs 1 and 2 were shown to be the only virus- encoded genes required for replication of BWYV RNA (Reutenauer et al., 1993) and BYDV- PAV (Mohan et al., 1995) in protoplasts. In contrast to ORF 2, the block of ORFs 3,4 and 5, located within the 3' half of luteoviral genomes, is the most highly conserved among luteoviruses. ORFs 3,4 and 5 are expressed from a major subgenomic RNA (sgRNA) of about 2.9 kb. This sgRNA was found in cells infected with BYDV-PAV (Dinesh-Kumar et al., 1992; Kelly et al., 1994), BWYV (Falk et al., 1989), PLRV (Tacke et al., 1990) and other luteoviruses. Two smaller sgRNAs of 0.8 kb and 0.3 kb have been detected in PAV-infected cells (Miller et al., 1995). The 0.7 kb SgRNA was found in BWYV-infected plants (Falk et al., 1989) and 0.4 kb sgRNA was 21 identified in plants infected with BWYV and ST9a RNA (Passmore et al., 1993). None of these smaller sgRNA except for 0.8 kb sgRNA of BYDV-PAV appears to have messenger activity. No sgRNAs of luteoviruses are encapsidated (Miller et al., 1995). ORF 3 of all luteoviruses encodes 22 to 23 kDa coat protein (CP; Miller, 1994). It was identified by comparing its deduced amino acid sequence to that of peptides present in purified virions, using anti-CP antibodies to P3 expressed in E. coli, by immunoprecipitation of cell- free translation products and by western blotting (Miller et al., 1988; Veidt et al., 1988). Coat protein is required for formation of intact particles but unnecessary for viral RNA replication (Reutenauer et al., 1993; Mohan et al., 1995). It plays a role in virus transmission and vector specificity but the mechanism of the interaction between the viral coat and the vector is unclear. ORF 4 is contained completely within ORF 3 and is expressed using leaky scarming mechanism of translation initiation (Dinesh-Kumar and Miller, 1993). It has biochemical properties of cell-to-cell movement protein including ability to be phosphorylated and bind nucleic acid nonspecifically (Tacke et al., 1991 and 1993). Chay et al. (1996) demonstrated that P4 is involved in systemic infection and therefore possibly movement in plants. ORF 5 is expressed as a C-terminal extension of coat protein by readthrough of the coat protein termination codon (Miller et al., 1995). It has been detected in infected protoplasts, plants and, in a shortened form, in purified virus particles (Cheng et al., 1994; Brault et al., 1995; Wang et al., 1995). P5 is not necessary for the assembly of virions (Reutenauer et al., 1993; Brault et al., 1995; Mohan et al., 1995) but is required for aphid transmission (Brault et al., 1995; Chay et al., 1996). RNA agents associated with luteoviruses A 322 nt single-stranded satellite RNA (sat RPV RNA) has been found in an Australian isolate of BYDV-RPV (Miller et al., 1991). Its known properties, replication strategy, and helper virus specificity are described in Chapter 1. 22 Another example of an association between a luteovirus and a second unrelated RNA is the ST9 isolate of BWYV (Falk et al., 1989). Particles of BWYV ST9 isolate contain two prominent RNAs, the 5,641 nt BWYV gRNA and a second, 2,843 nt long associated RNA, that lacks any significant sequence homology to BWYV gRNA (ST9a RNA). ST9a RNA encodes its own polymerase (Chin et al., 1993; Figure 4). The polymerase contains two regions of significant homology to putative RdRp of subgroup I luteoviruses but no homology to BWYV polymerase (Chin et al., 1993). Therefore, ST9a RNA is assigned to subgroup I luteoviruses. Because it encodes a polymerase, ST9a RNA is capable of independent replication in protoplasts (Passmore et al., 1993). However, it does not encode a coat protein (Chin et al., 1993) and depends on BWYV for encapsidation, systemic spread and aphid transmission. In BWYV and ST9a RNA-infected plants, both BWYV and ST9a RNA become encapsidated separately by BWYV capsid protein. Both types of particles are then transmitted by Myzus persicae (Falk and Duffus, 1984; Sanger et al., 1994). The ST9a RNA exacerbates symptoms caused by BWYV. Plants infected with BWYV and ST9a RNA exhibit more severe stunting and yellowing, compared to those infected with BWYV only (Sanger et al., 1994). They also contain about 10 times more virions per gram of tissue (Falk and Duffus, 1984). The ST9a RNA was also shown to stimulate accumulation of BWYV gRNA in protoplasts and plants (Passmore et al., 1993; Sanger et al., 1994). However, ST9a RNA does not alter the phloem limitation of BWYV and does not cause subsequent invasion of mesophyll tissue (Sanger et al., 1994). Although ST9a RNA has some properties of satellite RNAs, such as helper-dependency for encapsidation and transmission, it is not a true satellite as it does not depend on its supporting virus for replication (Mayo et al., 1995). 23 Satellites Classification Satellites are sub-viral agents composed of nucleic acid molecules that depend for their productive multiplication on co-infection of a host cell with a helper virus (Mayo et al., 1995). Satellite nucleic acids have substantially distinct nucleotide sequences fix)m those of the genomes of either helper virus or host. When a satellite encodes the coat protein, in which its nucleic acid is encapsidated, it is referred to as a satellite virus. The particles of satellite viruses are anitgenically and morphologically distinct fix>m those of the helper virus (Mayo et al., 1995). Satellites have been classified into following groups (Mayo et al., 1995; Table 3): (i) dsDNA satellites, (ii) ssDNA satellite viruses, (iii) dsRNA satellites, (iv) ssRNA satellite viruses, and (v) ssRNA satellites (satRNAs). There appears to be no taxonomic correlation between the viruses that are associated with satellites. For example, plant satRNAs, that represent the largest group among the satellites, have been found associated with viruses from at least nine virus groups: the cucumo-, tombuS", nepo", enamo-, sobemo-, carmo-, luteo-, umbra- (Roossinck et al., 1992), and potexviruses (Lin and Hsu, 1994). The majority of viruses that support plant satRNAs have host range restricted to dicotyledonous plants and are encapsidated in icosahedral particles. In addition, some viruses have more than one satellite and one satellite may depend on the helper virus for replication but on a second satellite for encapsidation (Francki, 1985; Roossinck et al., 1992). The only example of a dsDNA satellite is the satellite bacteriophage P4 in the family Myoviridae (Bertani and Six, 1988). The ssDNA satellite viruses are members of the %tn\isDependovirus in the family Parvoviridae (Bems, 1990). The dsRNA satellites have 24 Table 3. List of known and tentative species of ssRNA satellites (Mayo et al., 1995) Helper virus Abbreviation Reference ssRNA satellite viruses subgroup 1: chronic bee-paralysis virus associated satellites chronic bee-paralysis SCBPV Overton et al., 1982 subgroup 2: tobacco necrosis virus satellites maize white mosaic SMWMV Zhang etal., 1991 Panicum mosaic SPMV Masutaetal., 1987 tobacco mosaic STMV Mirkov et al., 1989 tobacco necrosis STNV Ysebaertet al., 1980 satellite RNAs subgroup 1: genus deltavirus hepatitis B HDV Taylor, 1992 subgroup 2: B type mRNA satellites Arabis mosaic large satAMV Liu et al., 1990 bamboo mosaic satBaMV Lin and Hsu, 1994 chicory yellow mottle satCYMV Rubino et al., 1990 grapevine Bulgarian latent satGBLV Fritsch et al., 1993 grapevine fanleaf satGFLV Fuchs et al., 1989 myrobalant latent ringspot satMLRV Frisch et al., 1993 pea enation mosaic satPEMV Demler and Zoeten, 1989 strawberry latent ringspot satSLRV Kreiah et al., 1993 tomato black ring satTBRV Hemmer et al., 1987 25 Table 3. (Continued) Helper virus Abbreviation Reference tentative species: groundnut rosette satGRV Murant et al., 1988 subgroup 3: C type linear satRNAs cucumber mosaic satCMV CoUmer and Howell, 1992 Panicum mosaic small satPMV Masuta et al., 1987 peanut stunt satPSV Naidu et al., 1991 turnip crinkle satTCV Simon and HoweU, 1986 tentative species: Cymbidium ringspot satCyRSV Rubino et al., 1990 tobacco necrosis small satTNV CoUmer and Howell, 1992 tomato bushy stunt satTBSV Gallittelli and Hull, 1985 subgroup 4: D type circular satRNAs Arabis moscaic small satAMV Kaper et al., 1988 E^V barley yellow dwarf satRPV Miller et al., 1991 luceme transient streak satLTSV AbouHaidar and Paliwal, 1988 Solanum nodiflorum motde satSNMV Francki, 1990 subterranean clover mottle satSCMV Davies et al., 1990 tobacco ringspot satTRSV Buzayan et al., 1986 velvet tobacco mottle satVTMV Gould et al., 1981 tentative species: rice yellow mottle satRYMV Sehgal et al., 1993 26 been found in association with viruses of the family Totiviridae (Wickner, 1992). The list of ssRNA satellite viruses and satRNAs is shown in Table 3. Two subgroups of ssRNA satellite viruses are known (Mayo et al., 1995): (i) chronic bee-paralysis virus associated satellite, and (ii) tobacco necrosis virus satellite. SatRNAs have ssRNA genomes and are encapsidated into the capsid protein of the supporting virus. Thus, the particles containing satRNA are antigenically identical to those of the helper. SatRNAs have been classified into four different subgroups (Mayo et al., 1995; Table 3): (i) genus deltavirus, (ii) B type mRNA satellites, (iii) C type linear satRNAs, and (iv) D type circular sat RNAs. Hepatitis delta virus (HDV), a member of genus deltavirus subgroup, has circular RNA genome. It encodes proteins used during its replication. It replicates via a rolling circle mechanism and is capable of self-cleavage at a ribozyme structure (Taylor, 1992; Jeng et al., 1996). Its catalytic activity resembles that of plant associated D type circular satRNAs and certain viroids (Jeng et al., 1996). The B type mRNA satellites are 0.8 to 1.5 kb, encode a non-structural protein that may be essential for their replication and exhibit some sequence homology to their helpers (Mayo et al., 1995). The B type mRNA satellites rarely modify symptoms caused by their helper viruses in plants (Collmer and Howell, 1992). SatRNA of bamboo mosaic potexvirus is one of the three known satRNAs that are associated with a virus that infects monocotyledonous plants (Lin and Hsu, 1994). Moreover, unlike other satRNAs, this satellite is encapsidated in rod-shaped virus particles (Lin and Hsu, 1994). 27 The C type linear satRNAs contain ssRNA genomes of less than 0.7 kb. They do not encode any functional protein and do not form circles at any stage of the infection (Mayo et al., 1995). They can substantially modify symtoms caused by their helper viruses (Collmer and Howell, 1992). The D type circular satRNAs exhibit the following properties (Francki, 1985; Roossinck et al., 1992; Mayo et al., 1995): (i) They are about 350 nt long, (ii) have a high degree of secondary structure, (iii) contain no significant ORFs, (iv) exist as a circular as well as linear molecules, (v) replicate via symmetrical or asymmetrical rolling circle (Figure 3), and (vi) the replication involves self-cleavage of circles and multimeric forms by an RNA-catalyzed reaction (Prody et al., 1986). At least three other plant pathogens, avocado sunblotch viroid (Hutchins et al., 1986), peach latent mosaic viroid (Hemandez and Flores, 1992) and carnation stunt associated viroid-like EINA (Hemandez et al., 1992) can also self-cleave in the absence of proteins. SatRNA of barley yellow dwarf luteovirus (satRPV RNA) is another satRNA associated with a virus that infects monocotyledonous plants (Miller et al., 1991). A small RNA that is associated with rice yellow mottle sobemovirus (satRYMV RNA) and shows homology to satRNA of lucerne transient streak sobemovirus may be another example of a type D circular satRNA supported and encapsidated by a virus restricted to monocotyledonous (Sehgal et al., 1993). However, the information about the satRYMV RNA sequence, structure and mechanism of replication is lacking. The specificity of satellite replication occurs at the level of both the helper virus and host plant (Roossinck et al., 1992). In addition, satellites can have drastic effects on helper virus replication and accumulation and may alter the symptoms induced by their helper viruses 28 (Francki, 1985; Collmer and Howell, 1992). The details of the complex three-way interaction between the satellite, helper virus and host have not been determined but some information is available. Requirements for replication of plant virus satellites Satellites depend completely on their helper viruses for replication (Francki, 1985). Satellites do not encode their own polymerases and, unlike viroids, do not rely exclusively on the replication machinery of the host (Flores, 1995). The replication of satellites presumably requires the presence of the replicase, which includes helper virus-encoded RdRp and host components (Quadt et al., 1993). However, it remains to be proven that the viral polymerase is the only component required for satellite replication that is supplied by the helper virus. The following experiments suggested that the viral polymerase is actually involved in the satellite replication: (i) Partially purified cucumber mosaic virus (CMV) RdRp isolated fix)m tobacco protoplasts inoculated with RNA 1 and 2 of CMV was shown to replicate satCMV in vitro (Wu et al., 1991; Hayes et al., 1992). RNAs 1 and 2 each contain one ORF, encoding proteins of 111 kDa and 97kDa, and were shown to be required for the replication of viral genomic RNAs (Palukaitis, 1992); (ii) Satellite tobacco necrosis virus (STNV) can replicate in tobacco protoplasts transfected with a vector expressing tobacco necrosis virus (TNV) ORFs 1 and 2 under the control of the cauliflower mosaic virus 35S promoter, and in the protoplasts derived from transgenic Nicotiana tabacum expressing the same genes. ORFs 1 and 2 encode the putative RdRp of TNV (Andriessen et al., 1995). However, putative CMV and TNV RdRps utilized the satellite template with much lower efficiency when compared to plants or protoplasts inoculated with helper virus RNA and its satellite (Hayes et al., 1991; Andriessen et al., 1995). Several explanations can account for the 29 lower efficiency of satellite replication. It is possible that replication of satellite does not always involve the same mechanisms as that required for viral RNAs replication and that other factors, which are specific for satellite replication, may be involved. It seems likely that parts of the helper virus genomes that may stimulate satellite multiplication are lacking from the RdRp complexes. In addition, the replicase transcript level in protoplasts transiently or stably transformed with putative RdRp genes of TNV is much lower compare to the level of TNV genomic RNA that serves as polymerase mRNA in infected cells. It is also possible that TNV replicase encoded by a nuclear gene does not form proper structures which are required for efficient replication (Andriessen et al., 1995). SatTRSV RNA was shown to replicate efficiendy in protoplasts inoculated with tobacco ringspot virus (TRSV) genomic RNA and satTRSV RNA even in the presence of actinomycin D (Buckley and Bruening, 1990). This indicates that synthesis of host RNA is not required for satellite replication. However, pre-inoculation of the protoplasts with actinomycin D 24 hrs prior to inoculation inhibited replication of satTRSV RNA. This suggests that a host factor is involved in satTRSV RNA replication. However, replication of TRSV genomic RNA was not monitored in these experiments, and thus specific conclusions cannot be drawn. The RdRp of helper virus must be able to recognize specific sequence(s) or secondary structure(s) within the (+) and (-) surands of satRNA (replication origin) to initiate satRNA synthesis. It is conceivable that efficiency of satRNA replication depends upon the efficiency of these interactions (Roossinck et al., 1992). However, such sequences or secondary structures on a satRNA have not been identified. Linear satellites presumably replicate by a mechanism similar to that of their helper viruses. The RdRp must first bind to the 3' end of the (+) strand RNA to initiate synthesis of (-) strand, and then to the 3' end of the (-) strand to initiate synthesis of the (+) strands (Roossinck et al., 1992). Therefore, the 3' and 5' ends of both helper virus genomic RNA and 30 satRNA should possess sequence motifs or secondary structures able to interact with the same viral replicase. There is usually very little appreciable sequence homology at 5' and 3' termini of linear satellites and their helper viruses (Francki, 1985). However, some chimeric satRNAs have been described. For example, the common 5' end sequences in the satellite and genomic RNAs were identified in satCyRSV RNA (Rubino et al., 1990). Similarly, the 3' half of the satRNA C of turnip crinkle virus (TCV), composed of 166 bases, is nearly identical to two regions at the 3' end of the TCV helper virus genome (Simon and Howell, 1986). In addition, some satRNAs, such as satCMV RNA, do not require perfect 5' ends or cap structure for the infectivity, whereas infectious transcripts of genomic RNAs usually do not tolerate extra sequences at the 5' ends of their genomes (Masuta et al., 1988). Oncino et al. (1995) showed that both 5' and 3' noncoding regions of large satTBRV RNA are important for replication. A large reduction in the accumulation of satTBRV RNA was observed when minor modifications were introduced into the 5' region and chimeras with a heterologous 5' noncoding regions failed to repUcate (Oncino et al., 1995). The 5' noncoding region of satTBRV RNA can possibly form a hairpin structure that could be involved in the recognition by the viral replicase but the actual existence and fiinction of such conformation remains to be determined. The D-type circular satRNAs replicate by the rolling circle replication pathway that can be either symmetrical or asymmetrical (Figure 3). SatTRSV RNA (Passmore and Bruening, 1993), satLTSV RNA (Foster and Symons, 1987) and satRPV RNA of RPV BYDV (Miller et al., 1991) appear to rely on the symmetrical model (Figure 3B) because both strands are capable of self-cleavage and discrete monomeric and multimeric forms of the (-) strand accumulate in infected cells (Davies et al., 1990; Hutchins et al., 1985; Keifer et al., 1982; Silver et al., 1994; Symons, 1989). In contrast, the (-) strands of satRNAs of some sobemoviruses lack self-cleavage structures, and accumulate only in high molecular weight forms, suggesting that they employ the asymmetrical strategy (Figure 3A; Davies et al., 1990). The particular strategy used may not be crucial to the replication of a sateUite, as Sheldon and 31 Symons (1993) showed that self-cleavage of the (-) strand was not necessary for satLTSV RNA repUcation. However, Van Tol et al., (1991) showed that the formation of circles is an essential step in the replication of satTRSV RNA. SatTRS V RNA mutated at either of two distant sites (nt 51 + 52 and nt 212 + 213) that has normal rate of self-cleavage but reduced spontaneous circularization of (-) strand failed to replicate. Double mutant satTRSV RNA with restored ability to form circular intermediates replicated efficiently (Van Tol et al., 1991) Similarly to linear satellites, circular satRNAs must also contain replication origin in both (+) and (-) strands. The identity of such sequences or secondary structures that are specifically recognized by the helper virus RdRp is unknown. The majority of plant virus satellites do not encode any proteins. Therefore, their nucleotide sequence and the resultant secondary or tertiary structure determine the interactions between the satRNA and the components of replication complexes. The B type mRNA satellites encode a non-structural protein that may also be essential for the replication (Mayo et al., 1995). For example, Henmier et al. (1993) showed that the entire 48K protein encoded by the satTBRV RNA is needed for the satRNA replication. The multiplication of large satRNA of arabis mosaic virus was also shown to require the 39kDa protein encoded by the satellite (Liu and Cooper, 1993). It was suggested that these proteins may interact with the repUcation complexes to allow replication to proceed. They may divert the replicases from their normal substrates and target them to the satRNAs. Helper vinis specificity and host range of plant virus satellites The features of the helper virus or host plant, which are required for the replication of certain sateUite, are not well defined and the mechanisms controlling these interactions are not understood in most cases. Related viruses may or may not replicate the same satRNA in some hosts but not in the others. The same helper virus may support the replication of a 32 heterologous satellite. In addition, the efficiency of satellite replication may vary widely with the strain of helper virus and the host plant (Francki, 1985; Roossinck et al., 1992). B type mRNA satellites Some, but not all, B type mRNA satellites can be exchanged among different helpers (Mayo et al., 1995). For example, isolates of tomato black ring nepovirus (TBRV) can be divided into two main serotypes: S and G. SatRNAs from S and G serotypes are about 60% homologous and multiply with viruses of homologous serotype but not with viruses of heterologous serotype (Fritsch et al., 1993). However, satellite from TBRV G multiplies when co-inoculated with a pseudorecombinant helper virus comprising RNA 1 from TBRV G and RNA 2 from TBRV S. Grapevine chrome mosaic nepovirus (GCMV) or a pseudorecombinant comprising of GCMV RNA 1 and TBRV G RNA 2 does not support replication of satellite from TBRV G (Mayo and Hemmer, 1993). GCMV or a pseudorecombinant comprising of GCMV RNA 1 and TBRV S RNA 2 supports the multiplication of TBRV S satRNA but the yield of the satellite progeny is significantly lower than with TBRV S as a helper (Oncino et al., 1995). Analysis of chimeric satRNAs, obtained by exchanging different regions between satRNAs from G and S isolates of TBRV, showed that the 5' and the 3' noncoding regions of satRNA, although important for replication, are not determinants of helper virus specificity. Oncino et al. (1995) suggested that the 48kDa satRNA-encoded protein plays the important role in the specificity of satTBRV replication. The proteins encoded by satRNAs from either G or S serotypes of TBRV are about 90% identical but satTBRV RNA from G serotype is only about 60% identical to that from S serotype TBRV (Fritsch et al., 1993). The proteins are most conserved in central part and most different in their N-terminal 120-130 amino acids (Fritsch et al., 1993). No particular signals within their sequence were identified and the role of the proteins in the specificity of satTBRV RNA replication remains speculative. In addition to helper virus, the host plant was 33 also shown to play a role in the satTBRV RNA replication. SatTBRV RNA replicates very efficiently in N. clevelartdii and Petunia hybrida but poorly in Chenopodium guinoa (Roossinck et al., 1992). Large satRNA of arabis mosaic nepovirus (AMV) from lilac multiplies when co- inoculated with serologically similar ash or ivy strains of AMV (Liu et al., 1991). However, no satellite progeny was detected in cells co-inoculated with AMV strains from hop or sugar beet which have few, if any, different antigenic determinants. This suggests that determinants for helper virus specificity are not located in the capsid-coding region of RNA 2 (Liu et al., 1991). More serologically distinct nepoviruses, such as dogwood mosaic, strawberry latent ringspot, grapevine fanleaf (GFLV) or cherry leaf roll virus also do not support satAMV RNA replication. In contrast, AMV was able to replicate satRNA of GFLV (Hans et al., 1992). Large satRNA associated with bamboo mosaic potexvirus (BaMV) can be replicated by BaMV helper in bamboo (Bambusa vulgaris), barley (Hordeum vulgare) and Chenopodium guinoa (Lin and Hsu, 1994). Unrelated viruses, such as potato virus X, tobacco mosaic virus, and C!MV, failed to support satBaMV RNA replication (Lin and Hsu, 1994). The ability of satBaMV to multiply in both monocotyledonous and dicotyledonous host plant suggests that both monocots and dicots can supply all components required for satRNA infectivity. C type linear satRNAs The most extensively studied interactions are those between the different strains of satCMV RNA, strains of CMV helper or related cucumoviruses and the host plants. Various satCMV RNA associated with different strains of CMV can replicate in both solanaceous and cucurbit host plants (Roossinck and Palukaitis, 1991) and the majority of satCMV RNAs replicates to high levels in solanaceous plants but generally poorly in cucurbits. The WL^- satRNA is an exception because it replicates to high levels in both solanaceous (tobacco) and cucurbit (zucchini squash) plants with most CMV strains, including Fny-CMV, as helpers. 34 The Sny-CMV strain is not able to replicate WLj-satRNA to detectable levels in zucchini squash but supports WLj-satRNA replication efficiently in tobacco (Roossinck and Palukaitis, 1991). The Fny strain of CMV helper differs less than 1% in nucleotide sequence from Sny- CMV (Gal-On et al., 1995). The poor replication of WL^-satRNA in zucchini squash was mapped to RNAl of Sny-CMV (Roossinck and Palukaitis, 1991). This implies a host-virus interaction involving RNAl or the la protein encoded by RNAl. The la protein contains conserved motifs of a helicase and is required for replication of viral RNAs but its precise role in the replication of satRNA is unknown (Roossinck and Palukaitis, 1991; Gal-On et al., 1995). Tomato aspermy cucumovirus (TAV) that is closely related to CMV was shown to serve as a helper for satCMV RNA (Moroines et al., 1992; Moroines et al., 1994). As is the case for CMV, the nature of the interactions of TAV with satCMV RNA depends on the strain of TAV, strain of satCMV RNA and the species of the host. Some TAV strains were shown to serve as efficient helpers of some satCMV EU^A, such as B2-satRNA, B3-satRNA, G-satRNA and WL2-sat RNA but they do not support the accumulation of others, such as Ix-satRNA in systematically infected leaves of tobacco or tomato. Moroines et al. (1992) demonstrated that the ability of TAV strains to serve as effective helpers of satCM RNA may be controlled by the extent of satRNA spread or encapsidation rather than by the efficiency of satRNA replication. Moroines et al. (1994) showed that helper virus pseudorecombinants which have RNAs 1 and 2 from CMV supported the systemic movement and accumulation of Ix-satRNA as efficiendy as CMV, whereas pseudorecombinants having RNAs 1 and 2 from TAV supported replication of B2-satCMV RNA very poorly. Thus, the ability to support the accumulation and systemic movement of satCMV RNAs depends primarily on RNA 1 and 2 instead of RNA 3, which is presumed to encode movement fiinctions. The mechanism of the interaction between the satCMV RNA and the RNA 1 and/or RNA 2 of the helper virus or their encoded proteins is not known (Moroines et al., 1994). Bemal and Garcia-Arenal (1994) identified at least four 35 nucleotides, scattered over the Ix-satRNA molecule, that determine satRNA phenotypes defective for movement and suggested that the ability of different strains of satCMV RNA to move systematically may depend on conformational changes of the satRNAs. In contrast to TAV, peanut stunt cucumovirus (PSV) that is related to CMV and TAV and often contains its own satRNA (satPSV RNA) does not support replication of satCMV RNA (Kaper et al., 1978). In addition, satPSV RNA does not replicate in the presence of CMV as a helper (Naidu et al., 1991). This could be explained by little sequence homology between satPSV and satCMV RNA. SatRNAs of tombusviruses, such as satTBSV RNA, satAMCV RNA, satCIRSV RNA, satCyRSV RNA, satPAMV RNA and satPLCV RNA, are closely related to each other. They can be supported by tomato bushy stunt tombusvirus (Galliteli and Hull, 1985). The eggplant mottle crinkle tombusvirus (EMCV), which unlike other tombusviruses does not contain satellite of its own, can also replicate satTBSV RNA. However, satTBSV RNA is not packaged in EMCV particles (Galliteli and Hull, 1985). This suggests, that encapsidation is not necessary for satRNA infectivity and that at least some satRNAs may move from ceil-to- cell in a naked form. On the other hand, unrelated viruses, such as mmip crinkle virus, do not act as helper viruses of satTBSV RNA (Galliteli and Hull, 1985). In addition, the host plant plays an important role since the replication of satTBSV RNA is more efficient m Nicotiana benthamiana than in N. clevelandii. D type circular satRNAs Among the small circular satRNAs of sobemoviruses, the ability of heterologous viruses to replicate certain satRNA varies greatly. A list of different helper viruses as well as nonhelpers that were identified for known satRNA of sobemovimses is shown in Table 4 (Gould et al., 1981; Jones et al., 1983; Jones and Mayo, 1984; Francki, 1985; Davies et al., 1990; Francki, 1990; AbouHaidar and Paliwal, 1988; Roossinck et al., 1992; Sehgal et al., 36 Table 4. Helper virus specificity of satRNAs of sobemoviruses satRNA Helper^ Nonhelper^ satLTSV RNA LTSV SNMV SoMV SBMV TRosV CfMV satSNMV RNA SNMV VIMV LTSV SCMV satVTMV RNA VIMV SNMV SoMV satSCMV RNA SCMV SoMV LTSV ^Abbreviations: SoMV, sowbane mosaic virus; SBMV, soutliem bean mosaic virus; TRosV, mmip rosette virus; CfMV, cocksfoot mottle virus, for other abbreviations see Table 3, page 24. 37 1993). Several interesting interactions between the satRNA, helper and host plant were observed among sobemoviruses. For example, LTSV supports the replication of satSNMV RNA but SNMV does not replicate satLTSV RNA. LTSV supports the multiplication of satSCMV RNA and SCMV acts as a helper for satSNMV RNA (Jones et al., 1984; Dall et al., 1990). The replication of satLTSV RNA is also supported by sobemoviruses that are normally devoid of satRNAs. These include sowbane mosaic virus, turnip rosette virus (TRosV), southern bean mosaic virus and cocksfoot mottle sobemovinis (CfMV; Sehgal et al., 1993). The replication of satLTSV RNA by TRosV is host-dependent. TRosV supports satLTSV RNA in Brassica rapa, B. raphanistrwn and Sinapis arvensis, but not Thlaspi arvense and Nicotiana bigelovii ( Sehgal et al., 1993). Furthermore, satLTSV JiNA replicates effectively and is encapsidated in the presence of CfMV in Triticum aestivum and Dactylis gleomerata. Host species, in which sobemoviruses have been shown to support satLTSV RNA, thus include members of both dicotyledonous and monocotyledonous families, namely Leguminosae, Chenopodiaceae, Solanaceae, Brassicaceae, and Poaceae. The ability of satLTSV RNA to replicate in plant species belonging to Brassicaceae co-infected with TRosV and in those of Poaceae family co-infected with CfMV is significant because these plant species are not hosts for LTSV (Sehgal et al., 1993). This indicates that the host plant requirements for the replication of helper virus and satRNA may not be the same and that satRNA may interact with a suitable helper in divergent monocotyledonous and dicotyledonous plants. Effects of satellites on disease symptoms Possible mechanisms The disease outcome is an interaction between the strain of the helper virus, strain of the satellite and the species and/or cultivar of the host plant (Roossinck et al., 1992; CoUmer and Howell, 1992). Most satellites have no effect on or attenuate disease symptoms. However, some satellites can exacerbate symptoms or produce new symptoms that are not 38 associated with the helper virus alone. Moreover, the modulation of disease symptoms by the same satellite can range from no effect on symptoms to attenuation or exacerbation. The type of response is determined by the strain of satellite, the strain of helper virus, the species of host plant and environmental conditions. Known satellites and their reported effects on disease symptoms induced by their helper viruses are listed in Table 5. The symptom alteration by viral satellites probably involves more than one mechanism, none of which is fully understood. The attenuation of disease symptoms is generally, but not always, accompanied by a substantial decrease in accumulation of helper virus RNA and helper virus titer (Harrison et al., 1987; Piazzolla et al., 1982; Kaper and Collmer, 1988; Hanada and Francki, 1989; Wu and Kaper, 1995; Kaper and Tousignant, 1977; Naidu et al., 1991, Collmer and Howell, 1992; Moroines et al., 1992; Roossinck et al., 1992). This leads to the suggestion that one mechanism of satellite mediated attenuation of symptoms is a competition between satellite and helper virus for shared, replicase-related factors (Roossinck et al., 1992; Collmer and Howell, 1992). In addition to competition for replication, satRNA may compete with the helper virus RNA for encapsidation in helper virus-encoded coat proteins, which leads to a reduction of virus titer. Satellites may also interfere with helper virus movement, both long distance and cell-to-cell (Naidu et al., 1991). Alternatively, the satellite itself may induce host defense response but there is no experimental evidence for this mechanism (Roossinck et al., 1992). However, Kong et al. (1996) showed that satRNA C, that a normally intensify symptoms of TCV, may be indirectly involved in induction of host defense response when co-infected with TCV containing heterologous coat protein. Satellites usually attenuate symptoms caused by their helpers only. For example, satCMV RNA, while ameliorating the symptom of CMV, does not attenuate symptoms of nonhelper tobacco mosaic virus, potato virus X, or potato virus Y in mixed infections (Tien et al., 1987). Exacerbation of disease symptoms in the presence of satRNA was never reported to be accompanied by an increase in the accumulation of helper virus RNA (Roossinck et al., 1992). 39 Table 5. Effect of satellites on disease symptoms (Roossinck et al., 1992; CoUmer and Howell, 1992) Satellite^ No effect Attenuation Exacerbation NR'* satellite viruses SMWMV + SPMV + + STMV + + + STNV + + B type mRNA satellites large satAMV RNA + + satBaMV RNA satCYMV RNA + satGBLV RNA satGFLV RNA + + satMLRV RNA + satPEMV RNA + satSLRV RNA + satTBRV RNA + + satGRV RNA + + + C type linear satRNAs satCMV RNA + + + small satPMV RNA 40 Table 5. (Continued) Satellite^ No effect Attenuation Exacerbation NR^ satPSV RNA + + satTCV RNA + + satCyRSV RNA + + satTNV RNA + satTBSV RNA + D type circular satRNAs small satAMV RNA + satRPV RNA + satLTSVRNA + + satSNMV RNA + + satSCMV RNA + satTRSV RNA + + satVTMV RNA + satRYMV RNA + ^For abbreviations see Table 3, page 24; t>Not reported. 41 This suggests that the mechanism of satellite-mediated symptom intensification involves direct interactions between the satellite and host plant similar to those induced by helper virus. In addition, exacerbation of symptoms may be due to a dosage effect because the satellite titer is generally extremely high in cells infected with satellite and helper virus. Satellite viruses SMWMV, SPMV, STMV and STNV have usually limited effect on symtomatology of their helper viruses (Table 5). However, SPMV was reported to exacerbate symptoms induced by PMV on two hosts (Collmer and Howell, 1992). In addition, STMV caused increased chlorosis in the form of bright-yellow patches relative to the mosaic caused by TMV-U2 helper virus in certain cultivars of pepper (Rodriguez-Alvarado et al., 1994). In contrast, in Jalapeno peppers, the presence of STMV ameliorated severe leaf distortions causes transiently by TMV- U2 and this was accompanied by significant reduction of virus titer (Rodriguez-Alvarado et al., 1994). Moreover, in some cell types STMV was shown to induce specific subcellular cytopathological effects which are different from these caused by TMV (Kim et al., 1989). There was no apparent correlation between cytopathological changes and disease symptoms. STNV, which usually has no disease modulating effect, was reported to reduce the number and size of local lesions in Phaseolus vulgaris plant (Kassanis, 1981). B type niRNA satellites In general, large satellites of nepoviruses have little or no effect on symptoms caused by their helper viruses (Table 5; Fritsch et al., 1993). No effect on symptom expression was reported for satPEMV RNA (Demler and de Zoeten, 1989). However, large satAMV RNA was reported to intensify symptoms in three species of Leguminoseae and ameliorate symptoms in 10 host species from families Apocyanaceae, Chenopodiaceae, Cucurbitaceae, Rosaceae, and Solanaceae (Liu et al., 1991). The satGRV RNA was shown to be largely 42 responsible for disease symptoms in groundnut (Murant et al., 1988; Kumar et al., 1991). Groundnut rosette virus (GRV), which is a helper for satGRV RNA, produces only transient mottle on groundnut plants. Groundnut rosette assistor luteovirus (GRAV), which is, in addition to satGRV RNA itself, necessary for aphid transmission of GRV and satGRV RNA, infect groundnut symptomlessly. Different forms of satGRV RNA have been identified, which are responsbile for different forms of rosette (chlorotic, green, mosaic) on groundnut (Murant and Kumar, 1990). Despite their ability to cause various symptoms in groundnut, these variants of satGRV RNA ameliorated symptoms induced by GRV alone in other plant species, including Nicotiana benthamiana (Murant and Kumar, 1990). However, one variant of satGRV RNA induced brilliant yellow blotch mosaic symptoms, instead of the usual veinal chlorosis and mild mottle which are characteristic for GRV alone (Kumar et al., 1991). Moreover, a cross-protection between satGRV RNA isolates has also been observed (Kumar et al., 1991) but the mechanism of this is unclear. C type linear satRNAs SatCMV is the best characterized among the satellites in respect to symptom modulating properties. The effect of satCMV RNA on disease symptoms was reviewed extensively by Collmer and Howell (1992). Different satCMV RNAs can produce different effects on different hosts which can range from necrosis or chlorosis to disease attenuation. Sequences controlling chlorosis and necrosis have been mapped. Necrogenic domain of both D-satRNA and Y-satRNA is located within 150 nucleotides at the 3' end of their molecules (Kurath and Palukaitis, 1989; Devic et al., 1989; Masuta and Takanami, 1989). Additional sequences outside necrosis domain influence the necrogenicity of Y-sat in both quantitative and qualitative manner (Wu and Kaper, 1992). SatCMV RNAs, that are similar in the necrogenic region but opposite in phenotype, can be converted from ameliorative to necrogenic and vice versa by exchanging as little as one to three nucleotides between the satellites (Masuta and Takanami, 43 1989; Devic et al., 1980; Sleat and Palukaitis, 1990). It seems likely that not only primary but also secondary and tertiary structure at the 3' end of the satRNA is responsible for eliciting lethal necrosis on tomato. However, no correlation between the 3' end structure and pathogenicity was observed using currently available computer models (Collmer and Howell, 1992). The role of helper virus is not well characterized in most studies of the pathogenicity of satCMV RNAs. Y-satRNA induces necrosis on tomato when supported by the subgroup n CMV strain KIN-CMV (Devic et al., 1989) and by the subgroup IY-CMV, but does not induce necrosis when supported by the subgroup 11-CMV (Wu and Kaper, 1992). D4- satRNA induces necrosis in tomato when supported by CMV strain of either subgroup I or EI, whereas WLM2-satRNA induces necrosis only in the presence of subgroup n CMV strains (Sleat et al., 1994). Moreover, WLM2-satRNA ameliorates the symptoms induced by subgroup I CMV strains without inducing necrosis. The co-inoculation of WLM2 satRNA with pseudorecombinant helper viruses formed between subgroup I and subgroup n CMV strains showed an association of the necrosis induction phenotype with RNA2 of the subgroup n strains (Sleat et el., 1994). In addition, environmental factors, such as temperature, were shown to influence necrogenicity of satCMV RNAs (Wu and Kaper, 1992; Wu et al., 1993, Kaper et al., 1995). Sequences controlling chlorosis were found in a different region of the satCMV RNAs. Chlorosis-inducing domain of Y-satRNA in tobacco is located within nucleotides 150-219 (Devic et al., 1989; Masuta and Takanami, 1989). Chlorosis domain of B-satRNA in tomato are located within 185 nucleotides at the 5' end of the satellite genome (Kurath and Palukaitis, 1989). Kuwata et al. (1991) showed that phenotypes of satCMV RNAs inducing either chlorosis or symptom attenuation can be altered reciprocally by changing die sequences in a limited region (nucleotides 191-193 for Y-satRNA). Although sequences controlling chlorosis in Y- and B5-satRNAs are localized to a similar and highly homologous region, it is not clear if 44 chlorosis is induced by each satRNA via a similar mechanism. A nucleotide change in B5- satRNA (ntl53) to one present in Y-satRNA will, for example, destroy chlorotic phenotype in B5-satIiNA (Sleat and Palukaitis, 1992). Furthermore, host specificity of chlorosis is apparently determined by whether nucleotide 149 is a U (tomato) or C (tobacco) (Sleat and Palukaitis, 1992). The nucleotide 153 is also important since deletion of this nucleotide does not affect satRNA replication but renders it non-pathogenic on either tobacco or tomato (Sleat and Palukaitis, 1992). Moreover, the mutation of nucleotides 185/186 on Y-satRNA destroys its ability to induce yellow mosaic in combination with CMV but not with tomato aspermy virus (Jaegle et al., 1990). Another interesting observation is that different satCMV RNA can induce either white or bright yellow chlorosis depending on host plant and strain of helper virus (CoUmer and Howell, 1992). In addition, in some cases satellite-induced chlorosis is phenotypically different from CMV-induced chlorosis and temperature-sensitive (CoUmer and HoweU, 1992). Furthermore, Masuta et al.(1993) identified a single incompletely dominant gene in wild Nicotiana species that controls yellow mosaic symptoms induced by Y-satRNA on tobacco plants. Attenuation of disease symptoms by certain satCMV RNAs is usuaUy accompanied by a reduction in the accumulation of CMV RNAl and 2 and decrease in the specific infectivity (CoUmer and HoweU, 1992). This is consistent with a competition between the satRNA and helper virus for repUcation. The abUity of some strains of satCMV RNA to attenuate CMV symptoms was used to engineer transgenic plants expressing satCMV RNA (Harrison et al., 1987; Pena et al., 1994; Komari et al., 1982; Jacquemond et al., 1988; McGarvey and Montasser, 1994). The disease attenuation is not always accompanied by decrease in accumulation of viral RNAs. An example of this is an interaction between satCMV RNA and TAV (Moroines et al., 1992, Harrison et al., 1987). Some satRNA of PST (G-satRNA, WC-satRNA) strongly suppress systemic symptom development in noninoculated leaves of Nicotiana tabacum (Naidu et al., 1991). This suggests 45 that in addition to competing for viral RNA replicase and cellular factors essential for replication, G- and WC-satRNAs interfere with movement and systemic spread of PS V. On the contrary, V-satRNA of PSV that differs from G-satRNA in six nucleotides only, has no effect on symptoms induced by PSV in tobacco (Naidu et al., 1991). Substitutions in two nucleotide positions were required to change the V-satRNA phenotype to ameliorative (Naidu et al., 1992). SatCyRSV RNA does not appear to impact CyRSV symptoms directly but modulates symptoms indirecdy. Infection of Nicotiana benthamiana with CyRSV leads to apical necrosis and death of plants. When satCyRS V RNA was expressed in transgenic plants no protection from apical necrosis was observed (Rubino et al., 1992). The lack of symptom attenuation in the presence of satCyRSV RNA was correlated with reduced production of DI RNAs which, in the absence of satCyRSV RNA, act as an attenuator of virus replication and disease. Rubino et al. (1992) demonstrated that satCyRSV RNA indeed inhibits DI formation and/or replication. The mechanism probably involves competition for some factors necessary for replication and/or packaging in virions. On the other hand satRNAs associated with other tombusviruses, including satTBSV RNA and satCyRSV RNA, were reported to reduce the number of local lesions caused by TBSV in Chenopodium amaranticolor (Gallitelli and Hull, 1985). Systemic symptoms induced by TBSV in N. clevelandii were also attenuated (Gallitelli and Hull, 1985). TCV is naturally associated with a number of subviral RNAs, including small satRNAs (satRNA D, satRNA F), chimeric satRNA (satRNA C), and DI RNAs (DI RNA G). SatRNAs D and F are avirulaent but satRNA C is able to intensify symptoms of TCV infection in Arabidopsis thaliana leading to plant death (Li and Simon, 1990). SatRNA C was shown to exacerbate symptoms on all hosts where TCV induced visible symptoms but did not produce any new symptoms on hosts that were symptomless when infected with TCV in the absence of satRNA C (Li and Simon, 1990). Lack of symptoms was not correlated with decrease in viral 46 or satRNA C accumulation. Similarly, the exacerbation of symptoms was not accompanied with an increase of TCV genomic RNA levels (Li and Simon, 1990). It was suggested that satRNA C mediated symptom intensification may be due to interactions of satRNA C sequences with a host component(s) and/or dosage effect brought about by the presence of large amounts of satellite molecules in infected ceils (Li and Simon, 1990). SatRNA C sequences that are involved in symptom exacerbation were localized to the region at the 3' end of satRNA C genome that is homologous to TCV and to the region comprising nt 79 to 100 that is homologous to satRNA D (Simon et al., 1988). Interestingly, satRNA C was able to eliminate or greatly attenuate symptoms induced by TCV derivative that contained a heterologous coat protein ORF fix)m related cardamine chlorotic fleck carmovirus (Kong et al., 1995). The inhibition of symptom development was accompanied by the reduction of viral genomic RNA accumulation (Kong et al., 1995). This suggests that the disease outcome is determined by the coat protein of TCV or its ORF in a way that is affected by satRNA C. Kong et al. (1996) proposed that 3' end of satRNA C interacts directly or indirectly with the TCV coat protein. In the absence of wild type coat protein the 3' end of satRNA C is involved in the induction of host defense response which results in restriction of virus movement and symptom attenuation (Kong et al., 1996). D type circular satRNAs Among the rolling circle satellites, satTRSV RNA is the only one known to ameUorate symptoms induced by the helper virus. SatTRSV RNA was shown to protect Blackeye 5 cowpeas against the severe necrotic and other symptoms induced by budblight strain of TRSV (Gerlach et al., 1986). SatTRSV RNA also reduces severe stunting induced by TRSV in Black Valentine beans (Gerlach et al., 1986). The symptom attenuation correlates with reduction in the accumulation of encapsidated TRSV genomic RNAs 1 and 2. The titer of TRSV is also generally lowered in plants co-inoculated with satTRSV RNA (Van Tol et al., 1991). Mutated 47 satTRSV RNA transcripts that did not form (-) strand circles and therefore did not repUcate failed to protect cowpea against co-inoculated TRSV (Van Tol et al., 1991). Analysis of progeny revealed that these mutants although incapable of efficient replication induced the accumulation of an apparently latent endogenous form of satTRSV RNA (Van Tol et al., 1991; Buzayan et al., 1995; Passmore et al., 1995). The latent form of satTRSV RNA differed from the wild type at one position (nt 54) only (Buzayan et al., 1995) but, unlike the wild type, had no symptom attenuating properties. On the other hand, non-replicating satTRSV RNA restricted the usual increase of cherry leafn)ll nepovirus (CLRV) in inoculated leaves of cowpea and ameliorated symptoms caused by CLRV although CLRV did not support satTRSV RNA replication (Ponz et al., 1987). Ponz et al. (1987) showed that satTRSV RNA reduced accumulation of CLRV translation products in vitro and altered the pathway of polypeptides produced and speculated that CLRV symptom attenuation was caused by inhibition of CLRV translation in vivo . The attenuating properties of satTRSV RNA were used for engineering virus resistant transgenic plants. Transgenic Nicotiana tabacum expressing satTRSV RNA developed primary lesions on TRSV inoculated leaves 1-2 days later than non-transgenic plants (Gerlach et al., 1987). Majority of new leaves were symptomless while others developed mild systemic reaction only 5-6 weeks after inoculation (Gerlach et al., 1987). Transgenic plants expressing (-) strand satTRSV EINA developed necrotic ringspot lesions initially but recovered and later developed resistant phenotype. Transgenic plants expressing an intemal fragment of satTRSV RNA were not protected and developed symptoms similar to those observed on TRSV infected untransformed plants. The symptom attenuation was accompanied by reduction in TRSV accumulation (Gerlach et al., 1986) which suggests competition between the satTRSV RNA and its helper virus for replication and possibly encapsidation. In contrast to satTRSV RNA, small satAMV RNA exacerbates disease symptoms in Chenopodium quinoa and hops and is responsible for hop nettlehead disease (Liu et al., 1991; Collmer and Howell, 1992). Likewise, satRNAs of sobemoviruses generally exacerbate 48 symptoms induced by their helper viruses (Table 5). For example, co-infection of satLTSV RNA and LTSV or TRosV in Chenopodium amaranticolor results in an apparent change of lesion type, from chlorotic to necrotic (Jones et al., 1983, Jones and Mayo, 1984). No information about the mechanism(s) responsible for disease exacerbation is available. Nothing is known about the sequences or structures within the rolling circle satRNA molecules that are required for disease modulation or about the interactions between the satRNA and helper virus. Satellite-mediated S5anptom alteration is clearly a complex process that is determined by interactions between the host, helper virus and satellite. 49 CHAPTER 3. SATELLITE RNA OF RPV BARLEY YELLOW DWARF VIRUS REDUCES ACCUMULATION OF RPV HELPER VIRUS RNA AND ATTENUATES RPV SYMPTOMS IN OATS A paper published in a shorter version in the Molecular Plant-Microbe Interactions Lada Rasochova and W. Allen Miller Abstract Satellite RPV RNA (satRPV RNA) is dependent on its helper virus (RPV barley yellow dwarf virus) for replication and encapsidation in protoplasts. The effect of satRPV RNA on disease symptoms and helper virus replication in plants has not been studied previously. We show that satRPV RNA transcripts are not mechanically transmissible, but they can be transmitted to oat plants by aphids {Rhopalosiphwn padi) that were allowed to feed on partially purified virus extracts from oat protoplasts. Plants infected with RPV and satRPV RNA developed milder symptoms than those infected with RPV helper virus alone. SatRPV RNA significantly decreased RPV RNA accumulation in plants and protoplasts. However, satRPV RNA did not attenuate symptoms caused by the severe mixed infection of RPV and PAV B YDVs and had no effect on PAV RNA accumulation. The concentration of encapsidated helper virus had to be above a threshold level to facilitate aphid transmission of itself and satRPV RNA from protoplasts to plants. This suggests that competition for both replication and encapsidation is involved in the reduction of helper virus RNA accumulation that results in this amelioration of symptoms in oats. 50 Introduction Barley yellow dwarf luteovirus (BYDV) infection results in stunting, yellowing or reddening of leaves, delayed or no heading and marked increase in sterile florets in its hosts (Gramineae). Symptoms vary with plant species, crop cultivar, age of plant, isolate of virus and environmental conditions (Rochow and Duffus, 1981). In plants, BYDV is confined to phloem (Jensen and D'Arcy, 1995). However, protoplasts from other tissues and undifferentiated cells can be infected by viral RNA by electroporation or polyethylene glycol treatment (Young et ai, 1989; 1991). BYDV is obligately transmitted from plant to plant by aphids in a persistent, circulative manner (Power and Gray, 1995). In order to transfer infectious genomic RNA transcripts into plants, partially purified virions from protoplasts infected with RNA transcripts can be fed to the aphids through artificial membranes. These aphids can then transmit virus to plants (Young et aL, 1991; Sanger et ai, 1994; Wang et al., 1995; Chayera/., 1996). At least five serotypes of BYDV have been defined by their specific aphid vectors (Power and Gray, 1995). Based on the genome organization, serological relationships, and cytopathological properties, luteoviruses and BYDV strains have been divided into two subgroups (D'Arcy, 1986; Miller, 1994; Mayo and Ziegler-Graff, 1996; Miller etal, 1995). Subgroup I includes the PAV and MAV serotypes of BYDV and others; subgroup n includes the RPV, SGV and RMV serotypes of BYDV, beet western yellows virus (BWYV), potato leafroU virus (PLRV), and others. The RPV and PAV serotypes of BYDV, subjects of this report, are so different from each other that they are considered to be separate viruses (D'Arcy, 1986; Miller, 1994) and we refer to them as such in this manuscript. In oats, PAV causes more severe symptoms than RPV (Martin and D'Arcy, 1995). These two viruses can act synergistically in mixed infections, resulting in more severe symptoms than either virus alone. Genomic RNAs of both viruses can be heterologously encapsidated in doubly infected plants (Wen and Lister, 1991). 51 Satellite RNAs (satRNAs) are sub-viral RNAs that depend for their productive multiplication on co-infection of a host cell with a helper vims (Francki,1985; Mayo et ai, 1995). They have no sequence similarity with the helper genome, yet generally accumulate to much higher levels than the helper. They can be co-encapsidated with the helper virus' genome or encapsidated separately (CoUmer and Howell, 1992). In either case, the virion proteins are encoded by the helper. SatRNAs are dispensable for helper vims replication, and can decrease helper virus genomic RNA accumulation (Roossinck et al, 1992). SatRNAs can attenuate or exacerbate symptoms, or produce new symptoms that are not caused by the helper virus alone (Roossinck et al., 1992; Collmer and Howell, 1992). Symptom production can be altered by changing any part of the combination of satRNA, helper virus, host plant, and environmental conditions (Wu et al., 1993; Kaper et ai, 1995). BCnown rolling circle satRNAs, including small satRNA of arabis mosaic nepovirus, satRNA of lucerne transient streak sobemovims, satRNA of Solarium nodiflorum mosaic sobemovirus and satRNA of velvet tobacco mottle sobemovims, usually exacerbate symptoms associated with their helper viruses (Roossinck et al., 1992; Collmer and Howell, 1992). The exception is satRNA of tobacco ringspot virus (satTRS V) that greatly attenuates symptoms caused by TRSV and reduces helper virus replication (Gerlach et al., 1986). The protective effect of satRNAs can be explained by competition between viral and satRNAs for replicase or shared replicase-related factor(s) (Hanada and Francki, 1989). SatRNA sequences have been expressed in transgenic plants to suppress development of disease symptoms caused by helper vimses (Gerlach et ai, 1987; Jacquemond et ai, 1988; Saito etal., 1992; Garvey and Montasser, 1994; Pena etal., 1994). However, satRNAs do not necessarily ameliorate disease symptoms simply by reducing helper virus replication (Harrison et ai, 1987; Moroines et al., 1992). A small molecular weight RNA (satRPV RNA) associated with an Australian RPV-like isolate (Miller et al., 1991) was shown to have all the properties of satellite RNA in oat 52 protoplasts (Silver et al., 1994). SatRPV RNA is 322nt long with no significant sequence homology to its helper virus (Miller et al., 1991). It replicates by a rolling circle mechanism in both (+) (encapsidated) and (-) strands (Silver et al., 1994). Circular and multimeric forms capable of self-cleavage at hammerhead ribozyme structures were identified in both strands (Miller and Silver, 1991). Full-length satRPV RNA transcripts from a cDNA clone were infectious in oat protoplasts in the presence of RPV helper virus (Silver et al., 1994). In contrast, the PAV virus of BYDV is unable to support satRPV RNA replication. One of die most obvious unanswered questions about the satellite has been its effect on disease symptoms development in plants. Until now, it was not possible to inoculate plants with a virus preparations that differ only by the presence or absence of the satellite RNA and have no difference in the helper virus. Here, we deliver satRPV RNA to whole plants and observe the effect on helper virus replication and disease symptoms. Results Lack of mechanical transmission of satRPV RNA. Although luteoviruses are obligately aphid transmitted, it was conceivable that satRPV RNA could be mechanically transmissible for the following reasons: (i) similar satRNAs such as satTRSV RNA are extremely infectious in plants (Gerlach et al., 1986); (ii) satRPV RNA has a high specific infecdvity in oat protoplasts (Silver et al., 1994); and (Hi) an RNA that is normally dependent on a luteovirus helper, BWYV ST9-associated RNA, is mechanically transmissible (Passmore et al., 1993). To attempt mechanical transmission, one week following aphid-inoculation of oats with an RPV+PAV mixture, one [ig of monomeric satRPV RNA transcript was rubbed on the first leaves with RNase-free abrasive. Plants were observed for symptoms and assayed for satRPV RNA replication in inoculated and uninoculated leaves by northern blot hybridization. In all experiments in this paper, linear, monomeric satRPV RNA was used as inoculum, so accumulation of abundant mul timers or circular forms was 53 considered evidence of replication. Although monomeric (+) strand satRPV RNA was detected in inoculated leaves one week after inoculation (data not shown), no replication, as indicated by multimeric or (-) strand satRPV RNA in inoculated leaves was detected, nor was satRPV RNA present in uninoculated leaves, despite several attempts. Therefore, we conclude that the satRPV RNA detected in inoculated leaves was simply the highly stable inoculum RNA, and that satRPV RNA is not mechanically transmissible under the conditions used here. Aphid transmission of satRPV RNA-containing virus from protoplasts to plants. We next attempted to transmit satRPV RNA to plants by feeding aphids on partially purified virus extracts of protoplasts that had been inoculated with satRPV RNA plus helper virus, and then transferring the viruliferous aphids to plants. This approach had been employed previously with infectious genomic RNA transcripts in protoplasts (Young et al., 1991, Chay et al., 1996). Initially, we used a mixture of RPV and PAV viruses (RPV+PAV) as helper for satRPV RNA, because RPV in the presence of PAV accumulates to higher titer than RPV alone and the mixture supports satRPV EINA in protoplasts (Silver et al., 1994). Aphid transmission to plants requires that both helper and satellite RNA are encapsidated. To test for satRPV RNA encapsidation, we compared total with encapsidated RNAs from protoplasts. Large quantities of encapsidated satRPV RNA, predominantly as a linear (+) stranded monomer (data not shown), were detected in agreement with the original preparations of encapsidated satRPV RNA from plants (Miller et al., 1991). The level of encapsidated RPV genomic RNA was almost undetectable (data not shown), and moderate quantities of PAV RNA were present (data not shown). Using ELISA we estimated that PAV virions were at least five-fold more abundant than RPV virions (data not shown). The latter were present in concentrations of less than 10|j.g/ml. 54 Aphids (Rhopalosiphum padi) were fed via Parafilm™ membranes on ttie above virus exttact from protoplasts for a 24 hr acquisition access period and then transferred to oat plants for a four day transmission access period. Northern blot hybridization of total RNA extracted from plants two weeks later revealed PAV RNA but neither RPV genomic nor satRPV RNA (data not shown). We suspected that titer of RPV virus was too low to facilitate aphid transmission to plants. Absence of transmission of RPV would preclude replication of satRPV RNA, even if the latter, which is encapsidated efficiently, were delivered efficiently to plants. In order to increase the likelihood of satRPV RNA transmission to plants, we increased the amount of encapsidated RPV RNA in the protoplast extract as follows. The above extract from infected protoplasts was spiked with purified, satellite-free virus to a final concentration of 30 [ig/ml of RPV. Following aphid acquisition and transmission of this spiked extract, satRPV RNA replicated in second leaves of all four inoculated plants as indicated by the presence of multimeric fornis (Fig. 1 A) synthesized during rolling circle replication. Therefore, the concentration of helper virus is critical for transmission of satRPV RNA to the whole plant fi-om protoplast extracts, and satRPV RNA need not be encapsidated in the same virions as helper. SatRPV RNA affects helper virus RNA accumulation in oats but has no effect on symptoms caused by mixed infection of RPV and PAV. Once the ability to transfer satRPV RNA to plants with a defined virus was achieved, we could observe its effect on replication of, and symptoms caused by, the helper. Genomic RNAs of RPV and PAV replicated in oat plants in the presence and absence of encapsidated satRPV RNA (Fig. 1, B and C). There was variation in levels of RPV genomic and satRPV RNAS between plant samples (each lane in Fig. 1, A and B) probably due to chance in sampling. Thus, we could not discern the effect of satRPV RNA on absolute amounts of 55 genomic RNA that accumulated, but we could compare the ratios of satRPV: genomic RPV RNA within each lane. This revealed a negative correlation between the amount of RPV (genomic and subgenomic) RNA and satRPV RNA that accumulated in any given leaf (Fig. 1A and B, compare lanes 9-10 and 11-12). RPV genomic RNA accumulated to higher levels in leaves with low satRPV RNA and to lower levels in ±e presence of high amounts of satRPV RNA. In contrast, PAV genomic RNA, which is not a helper for satRPV RNA, varied less between plant samples (Fig. IC). Thus, PAV appears to be distributed more uniformly than RPV and is not affected by the presence of satRPV RNA. In all samples, satRPV RNA accumulated to much higher levels than either RPV or PAV RNAs (note differences in exposure times between panels in Fig. 1). Oats infected with satRPV RNA in the presence of the RPV+PAV mixture showed typical B YD symptoms (Fig. 2B): severe stunting, yellowing and reddening of leaves, delayed or no heading and marked increase in sterile florets. Under these growth conditions, we observed no significant difference in symptom development between plants infected with RPV+PAV alone and those infected with RPV+PAV, and satRPV RNA (Fig. 2B) in three passages. The differences in the average height and weight of plants, number of sterile florets and ELISA values for both RPV and PAV coat protein were insignificant (data not shown). We conclude that the presence of satRPV RNA in mixed infections of RPV and PAV does not affect the already severe disease caused by the mixmre. SatRPV RNA attenuates symptoms and decreases helper virus RNA accumulation in oats infected with pure RPV. Because PAV alone (Martin and D'Arcy, 1995), or the mixture of PAV+RPV causes more severe symptoms in oats than RPV alone, the effect of satRPV RNA on disease symptoms could be masked by the presence of PAV. Thus, we tested the symptom- modulating properties of satRPV RNA in oats infected with RPV in the absence of PAV. 56 Instead of using tiie spike to increase the concentration of the helper virus, we used aphids (R. padi) that were maintained on RPV-infected, sateliite-ftee plants prior to feeding on extracts from protoplasts that were infected with RPV + satRPV RNA. After feeding for 24 hours on protoplast extracts, aphids were transferred to healthy, two-week old oat plants. Samples for total EiNA extraction were collected in quadruplicate from pooled tissue from 15 plants, instead of from individual second leaves, to avoid plant-to-plant variation. SatRPV RNA replicated in plants in the presence of RPV genomic RNA as indicated by abundant monomeric and multimeric forms (Fig. 3A). Plants infected with satRPV RNA accumulated much less genomic RNA than those infected with RPV alone (Fig. 3B, compare lanes 3-6 and 7-10). The relative amount of RPV genomic RNA in the presence of satRPV RNA was 13% ± 3% of that in the satelUte-free plants. However, satRPV RNA reduced ELISA readings by only 17% ± 6% (from 1.15 to 0.95). ELISA readings may not be quantitative with respect to helper virus concentration because satRPV RNA is encapsidated by helper virus coat protein. In summary, satRPV RNA seemed to decrease accumulation of helper virus genomic RNA substantially more than the coat protein in oat plants. SatRPV RNA strikingly reduced symptoms caused by RPV in oats under these growth conditions (Fig. 2A). Compared to plants infected with RPV alone, those which also contained satRPV RNA (i) showed less smnting, (ii) headed 5 days earlier, but one week later than healthy plants, (Hi) produced three times as many seeds, and (iv) yielded twice as much total leaf tissue. The differences between plants infected with RPV and those infected with RPV+satRPV RNA were especially apparent late in infection. These observations indicate that satRPV RNA not only decreases helper virus RNA accumulation in plants but attenuates disease symptoms caused by pure RPV virus of BYDV. 57 SatRPV RNA decreases helper virus RNA accumulation in oat protoplasts. SatRPV RNA may reduce RPV helper RNA accumulation in plants by affecting cellular levels directly, or by reducing the ability to spread through the plant. To distinguish between these possibilities, we examined the effect of satRPV RNA on RPV helper RNA accumulation in protoplasts. Earlier evidence suggested that oat protoplasts inoculated with RPV+PAV mixture accumulate less RPV genomic RNA in the presence of satRPV RNA (Silver et al., 1994). In order to avoid the synergistic relationship between RPV and PAV, we tested the effect of satRPV RNA on helper virus replication in oat protoplasts using only pure RPV virus as a helper. In the presence of pure RPV RNA helper, satRPV RNA replicated to high levels by 72 hrs as indicated by the presence of abundant monomeric and multimeric forms (Fig. 4A). In three independent samples, the amounts of RPV genomic and subgenomic RNAs were reduced to 32% ± 4% by the presence of satRPV RNA (Fig. 4B, compare lanes 1- 3 and 4-6). Therefore, satRPV RNA decreases the helper virus accumulation at the cellular as well as the whole plant level. Discussion Lack of mechanical transmission of satRPV RNA. Because B YD V is phloem-limited and obligateiy aphid transmitted, the simple explanation for the lack of mechanical transmission of satRPV RNA is that satRPV RNA did not enter the phloem and was inaccessible to helper virus. However, other mechanisms may be involved as well. For example, a satellite-like molecule of BWYV luteovirus, ST9a RNA, which is encapsidated in BWYV coat protein, can be transmitted mechanically, even though BWYV is phloem-limited and not mechanically transmissible. ST9a RNA was also shown not to alter phloem limitation of BWYV that would result in the ability of virus to spread to mesophyll cells (Sanger et al. 1994). 58 The high stability of satRPV RNA inoculum is consistent with other satellites. Satellite RNA of cucumber mosaic virus (satCMV RNA) can survive in vivo for at least 10 days without replication and can then multiply when the plant is inoculated with helper virus (Mossop and Francki, 1978; Mossop and Francki, 1979). Helper virus titer is a limiting factor for satRPV RNA transmission by aphids from the infected protoplast extract The inability to transmit satRPV RNA directly from protoplast extracts infected with RPV+PAV+satRPV RNA may be due to low RPV titer brought about by satRPV RNA out- competing RPV genomic RNA for encapsidation, especially if satRPV RNA is present in great molar excess. Northern blot hybridization indicated that, although infected protoplasts contained high quantities of RPV, PAV and satRPV RNA, satRPV RNA was the major RNA encapsidated. High ratios of satRPV RNA to helper virus RNA in virus particles have been reported for other satellites. In mixed infections of satTRSV and its helper, satTRS V RNA may comprise more than 90% of encapsidated RNA (Buzayan et ai, 1986). SatRNA of velvet tobacco mottle virus (VTMoV) can account for up to 50% total and over 80% encapsidated RNA (Randies etal., 1981). PAV RNA may have sequestered additional E^V coat protein through transencapsidation, resulting in even lower quantities of available RPV coat protein and further explaining low RPV titer. Wen and Lister (1991) observed that NY-PAV RNA was heterologously encapsidated in RPV particles, but there was no evidence of encapsidation of RPV in the coat protein of PAV. This suggests that RPV had to compete not only with its satellite, but also with PAV genomic RNA for encapsidation. Virus titer also significantly influences the efficiency of transmission by aphids (Pereira et ai, 1989; Pereu^a and Lister, 1989; Van den Heuvel etal., 1991; Power and Gray, 1995). Although Gray et a/. (1991) reported that virus titer was less important than acquisition access 59 period, we observed that feeding of aphids on protoplast extracts for longer than 24 hours resulted in high mortality of aphids. This can be explained by presence of feeding inhibitors in cell extracts (Duffiis and Gold, 1967). The fact that the titer of <10 ^ig/ml of RPV was not aphid transmissible, but 30 ^ig/ml was readily transmitted, is consistent with observations of Periera etal. (1989) who reported no transmission of PAV at 18 Hg/ml and 50% transmission at 70 ng/ml. Thus, using the spike to increase the concentration of helper resulted in successful transmission of RPV and satRPV RNA and supports our observation that helper virus titer was limiting factor for aphid transmission. Lack of efTect of satRPV RNA on symptoms caused by RPV+PAV. Because infections with RPV and PAV viruses have been observed to cause more severe symptoms in mixed infection than either virus alone, the effect of the satellite is probably masked. However, we did not compare these symptoms directly with those caused by PAV alone, so it is possible that the severe symptoms in the mixture were due entirely to PAV. Levels of satRPV RNA in infected leaves correlated negatively with the accumulation of RPV helper but PAV levels were unaffected. Similar observations were made previously in oat protoplasts (Silver et al., 1994). If PAV alone is responsible for the symptoms seen in the mixture, then it is not surprising that satRPV RNA had no affect on symptoms. The variability in RPV genomic RNA levels in satellite-free plants can be explained by observation of Pereira and Lister (1989). They showed that virus titers vary widely among different leaves on the same plant, and also among leaves of the same age in the same position on different plants. These variations were more pronounced with Clintland 64 oats than with other barley and wheat cultivars. Despite this variation it appears that, within a particular leaf, high levels of satRPV RNA correlate with lower levels of helper virus RNA in these mixed infections. 60 Possible mechanisms of reduction of virus symptom severity and RNA accumulation. The effect of satRPV RNA on helper virus RNA accumulation was compared more directly when pure RPV was used as helper. The use of pooled samples of several leaves from the different treatments allowed us to conclude that satRPV RNA reduces helper virus RNA accumulation. This probably explains the reduced symptom severity in the presence of satRPV RNA. Because satRPV RNA accumulated to high levels in the mixed infection, but had no effect on symptoms, it is likely that symptom reduction in the absence of PAV was attributable to diminished RPV RNA levels, rather than to high accumulation of satRPV RNA per se. Thus, satRPV RNA is not likely to ameliorate symptoms by actively inducing a host defense response. The small reduction in virus titer detected by ELIS A, relative to the large reduction in viral RNA indicates that the determining factor in coat protein steady state levels is not the amount of its mRNA. The correlation of reduced symptoms with reduced helper RNA levels, but not coat protein levels suggests that coat protein is not the major symptom determinant. Concomitant reduction of helper virus RNA levels and disease symptoms has been observed for many other satellite RNAs. SatTRSV RNA protects cowpeas against the severe necrosis and ringspots induced by the budblight strain of TRSV (Gerlach et al, 1986), and protects Black Valentine beans from stunting caused by the same virus (Gerlach et al., 1986). Symptoms and TRSV levels were also reduced in transgenic tobacco plants expressing satTRSV RNA (Gerlach et al., 1987). SatCMV RNAs that attenuate disease in many hosts also reduce virus accumulation (Piazzolla et al., 1982). However, some satCMV RNAs can intensify symptoms induced by CMV (Masuta and Takanami, 1989; reviewed by Collmer and Howell, 1992). SatRNA of peanut stunt virus (PSV) (G-satRNA) strongly suppresses systemic symptom development and helper virus RNA replication in noninoculated leaves in tobacco. On the contrary, V-satRNA of PSV that differs from G-satRNA in six nucleotide positions has no effect on symptoms in tobacco (Naidu et al., 1991). Changes in two 61 nucleotide positions were required for suppression of systemic symptoms (Naidu et aL, 1992). SatRNA C associated with turnip crinkle virus (TCV) that normally intensifies the symptoms of TCV attenuated symptoms in Arabidopsis when supported by TCV chimeric virus that had coat protein of cardamine chlorotic fleck virus (Kong et al, 1995). Thus, unlike our above conclusion with RPV, the coat protein (or its ORF) of TCV may determine symptoms in a way that is affected by satRNA C. Like satRPV RNA and its helper, symptom attenuation by satRNA C was correlated with a reduction m TCV RNA levels in plants. Thus, accumulation of satellite RNAs can reduce symptoms, by reducing helper virus levels, but this is not always the case. It is likely that satRPV RNA reduces RPV genomic RNA levels by competing for a component involved in the virus life cycle. The most obvious candidate would be the replicase, components of which could be host or viraly encoded. The inhibition of helper RNA accumulation in protoplasts by satRPV RNA are consistent with this hypothesis, but competition for encapsidation in coat protein can not be ruled out. As discussed previously, RPV coat protein encapsidates PAV RNA, but not vice versa (Wen and Lister, 1991), so it may be less specific, and more amenable to encapsidation of satRPV RNA, perhaps even more than its own genomic RNA. The small reduction in ELIS A readings with antibodies against RPV virions, relative to the larger reduction in RPV genomic RNA is consistent with this. Lack of encapsidation alone reduces genomic RNA accumulation considerably as observed with coat protein-less mutants of PAV (Mohan et aL, 1995), and BWYV (Reutenauer et aL, 1993). Inhibition of helper RNA translation is possible as satTRSV RNA mhibited translation of cherry leafiroll nepovirus (CLRV) even though CLRV cannot support satTRSV RNA replication (Ponz et aL, 1987) and there is no evidence that satTRSV RNA is translatable. Competition for components involved in cell-to-cell or long distance movement is possible in plants, but can not be the sole explanation, because satRPV RNA reduces helper virus RNA accumulation in protoplasts in which no such movement occurs. In summary, we know that 62 satRPV RNA competes with its helper for both replicase and coat protein, but further experiments, for example with coat protein-deficient mutants, will be necessary to determine the role of coat protein in reducing RPV genomic RNA levels. Finally, we must bear in mind that these experiments used only one set of environmental conditions and one host cultivar. Variation of these have been shown to affect symptom modulation by satCMV RNA (Wu et a/.,1993; BCaper et al., 1995, White et al., 1995). SatRNA symptom modulation involves a complex interaction among helper virus, satellite and host plant (Roossinck et al., 1992). The effects of satRPV RNA on different helpers, hosts and in different environmental conditions , remain to be determined. Acknowledgments We thank Cathy Chay and Bonnie Passmore for technical advise. Randy Beckett for aphid and virus maintenance, and Steward Gray for providing NY-RPV and virus-firee aphids. This work was funded by USDA National Research Initiative grant number 94373030469. Materials and Methods In vitro transcription Full-length satRPV RNA was transcribed firom permuted dimeric cDNA clone pT7Sat that was linearized with £coRI prior to transcription with T7 RNA polymerase using the T7 Megascript kit (Ambion, Austin, TX). The monomeric self-cleavage product that arose during transcription of the permuted, dimeric RNA was gel-purified, prior to inoculation (Silver et al, 1994). Plasmid pT7Sat is the same as pFL-WT (Silver et al., 1994) but has the satRPV RNA sequences flipped between the flanking EcoRI and HindOl sites relative to the promoters. Final RNA concentration was determined spectrophotometrically. '^P-Iabeled RNA probes were synthesized by in vitro transcription as described by Promega (Madison, WI) using [a ^^P]-CTP. Antisense satRPV RNA probe was transcribed with SP6 RNA polymerase from 63 pT7Sat linearized witli Hi/idin. Antisense 3'RPV probe was transcribed with SP6 RNA polymerase from plasmid pMB 102, comprising nucleotides 5026 to the end of the RPV genome, linearized with £coRI. Antisense 5'RPV probe, complementary to bases 809-1191, was synthesized by SP6 RNA polymerase transcription of Apal-linearized plasmid pML5 (Silver et al., 1994). Antisense 5'PAV probe was transcribed with T7 RNA polymerase from plasmid pPAVl-1 (Mohan et al., 1995) linearized with EcoRI. Whole plants: inoculation, virus propagation and puriHcation, and extraction of virion and total RNAs Virus isolates used were IL-PAV (from Anna Hewings, USDA/ARS University of Illinois), and NY-RPV (from Stewart Gray, USDA/ARS Comell University). No satRPV RNA was detected in either of these isolates by northern blot hybridization. Seven day-old oat plants were inoculated with BYDV by aphids (Rhopalosiphwn padi): viruliferous aphids were allowed to feed on these plants for 2-4 days and were then killed with pyrethrin msecticide (Ortho product no. 5485, San Ramon, CA). Plants were then transferred to an aphid-free growth chamber and grown with 16 hr days, at a constant temperature of 21°C. Fourteen days after aphid-inoculation, virions were purified from oat plants (Avena sativa cv. Clintland 64) and RNA extracted by the method of Waterhouse et al. (1986) as modified by Mohan et al. (1995). Total RNA was isolated two weeks after inoculation of plants by the procedure of Wadsworth et al. (1988) as modified by Dinesh-Kumar and Miller (1993). Protoplasts: inoculation, partial virus extraction, and total RNA isolation Protoplasts were isolated from Avena sativa cv. Stout suspension culture (ceU line S226 obtained from Howard Rines, USDA/ARS, University of Minnesota) as described previously (Dinesh-Kumar and Miller, 1993). Protoplasts were electroporated with 100 ng of genomic RNA from virions and 50 ng of gel-purified, monomeric satRPV RNA transcript (322 64 nt) as described by Silver et al. (1994). Total RNA was isolated 72 hours after inoculation of protoplasts by the procedure of Wadsworth et al. (1988) as modified by Dinesh-Kumar and Miller (1993). Virus was partially purified from infected protoplasts by the method of Wang et al. (1995) with the following modifications. Infected protoplasts were pelleted 72 hours after inoculation by centrifiigation 100 x ^ for 5 min. The pellet was resuspended in 100 jil of 10 mM phosphate buffer, pH 7.0, sonicated for 15 sec and centrifiiged at 7,826 x g for 5 min. The supernatant contained partially purified virions and was designated "protoplast extract". RNA analysis Denaturing agarose gel electrophoresis and northern blot hybridization was performed as in Mohan et al. (1995). Equal amounts of total RNA were loaded in each lane. This was determined by spectrophotometry and ensured by ethidium bromide staining of ribosomal RNAs prior to northern hybridization. Radioactive bands were detected using a Phosphorimager model 400 E and quantified with Imagequant 3.3 software (Molecular Dynamics, Sunnyvale, CA). Blots were stripped by boiling with O.lx SSC and 0.1% SDS for 30 min prior to re-probing with another probe. Aphid feeding on artificial membranes and inoculation of plants Aphid acquisition of virus through membranes and inoculation of plants was as described by Wang et al. (1995) with several modifications. 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A procedure for small- scale isolation of plant RNA suitable for RNA blot analysis. Analytical Biochemistry 172, 279-283. Wang, J. Y., Chay, C., Gildow, F. E., and Gray, S. M. 1995. Readthrough protein associated with virions of barley yellow dwarf luteovirus and its potential role in regulating the efficiency of aphid transmission. Virology 206,954-962. Waterhouse, P. M., Gerlach, W. L., and Miller, W. A. 1986. Serotype-specific and general luteovirus probes from cloned cDNA sequences of barley yellow dwarf virus. J. gen. Virol. 67, 1273-1281. Wen, F., and Lister, R. M. 1991. Heterologous encapsidation in mixed infections among four isolates of barley yellow dwarf virus. J. gen. Virol. 72, 2217-2223. White, J. L., Tousignant, M. E., Geletka, L. M., and Kaper, J. M. 1995. The replication of a necrogenic cucumber mosaic virus satellite is temperamre-sensitive in tomato. Arch. Virol. 140, 53-63. 71 Wu, G., Kaper, J. M., Tousignant, M. E., Masuta, C., Kuwata, S., Takanami, Y., Pena, L., and Diaz-Ruiz, J. R. 1993. Tomato necrosis and the 369 nucleotide Y satellite of cucumber virus: factors affecting satellite biological expression. J. gen. Virol. 74, 161-168. Young, M. J., Larkin, P. J., Miller, W. A., Waterhouse, P. M., and Gerlach, W. L. 1989. Infection of Triticum monococcum protoplasts with barley yellow dwarf virus. J. gen. Virol. 70,2245-2251. Young, M. J., Kelly, L., Larkin, P. J., Waterhouse, P. M., and Gerlach, W. L. 1991. Infectious in vitro transcripts from a cloned cDNA of barley yellow dwarf virus. Virology 180,372-379. 72 Inoculum: RPV+ RPV+PAV None PAV +satRPV A Probe: satRPV(-) Mmulti. h*-mono. B •gRNA Probe: 3'RPV(-) •sgRNA c Probe: & 5VAV(-) Lane: 1 2 3 4 5 6 7 8 9101112 Figure 1. Northern blot analysis of total RNA extracted from oat plants inoculated with R. padi fed on protoplast extracts infected with the indicated inoculum. Each lane represents total RNA from an individual plant. Equal amounts of RNA were loaded per lane. RNA was isolated from second leaves of the plants two weeks after inoculation, fractionated on 1% denaturing agarose gel, blotted to nylon membrane and hybridized with the indicated probes. The same blot was used in all three panels, using equal amounts of probe (5x10^ cpm/ml) per hybridization. The probe was stripped prior to hybridization with the next probe (as in Materials and Methods). Mobilities of satRPV RNA monomers (mono., 322 nt), multimers (multi.), RPV genomic (gRNA) and subgenomic (sgRNA) RNAs and PAV genomic RNA (gRNA) are indicated at right. Blots were exposed at -80°C with one intensifying screen for 3 hr (panel A), 18 hr (panel B) and 24 hr (panel C). 73 I RPV RPV No RPV+PAV RPV+PAV +satRPV Virus +satRPV Figure 2. Oat plants (Clintland 64) 8 weeks after aphid inoculation with the indicated combinations of virus and satRPV RNA. Plants per pot: panel A, 4-5; panel B, 7-8. 74 Inoculums RPV+ None RPV satRPV Probe: satRP\ multi. mono, B gRNA Probe: 5'RPV(-) - - • Lane: 1 2345678910 Figure 3. Northern blot analysis of total RNA extracted from oat plants inoculated with R. padi fed on protoplast extracts infected with the indicated inoculum. Equal amounts of RNA isolated from pooled tissue of 15 plants two weeks after inoculation were loaded on each lane. RNA was fractionated on 1% denaturing agarose gel, blotted to nylon membrane and hybridized with the indicated probes. Mobilities of satRPV RNA monomers (mono., 322 nt), multimers (multi.), and RPV genomic RNA (gRNA) are indicated at right. Equal amounts of probe were used in each hybridization. One blot was used, stripped and re-probed. Exposures were at -80°C with one intensifying screen for 1 hour (panel A) and 72 hours (panel B). 75 Inoculum: RPV+ RPV satRPV ir Probe: satRPV(-) mono. B -gRNA Probe: ki 3'RPV(-) ^ -^sgRNA M Lane: 1 2 3 4 5 6 Figure 4. Northern blot analysis of total RNA extracted from oat protoplasts 72 hours post inoculation with the indicated inoculum. Each lane represents equal amount of RNA isolated from an independent sample (-300,000 protoplasts per sample). Mobilities of satRPV RNA monomers (mono., 322 nt), multimers (multi.), and RPV genomic (gRNA) and subgenomic (sgRNA) RNAs are indicated at right. Equal amounts of probe were used in each hybridization. One blot was used, stripped and re-probed. Exposures were at -80°C with one intensifying screen for 2 hours (panel A) and 6 hours (panel B). 76 CHAPTER 4. SPECmCTTY OF RPV BARLEY YELLOW DWARF VIRUS SATELLITE RNA REPLICATION IS ASSOCIATED WITH THE POLYMERASE OF SUBGROUP H LUTEOVIRUSES A paper to be submitted to the Virology Lada Rasochova, Boni K. Passmore, Bryce W. Falk, and W. Allen Miller Abstract RPV barley yellow dwarf subgroup n luteovirus (BYDV-RPV) acts as a helper for the 322 nt satellite RNA (satRPV EiNA). SatRPV RNA reduces levels of RPV helper virus RNA and attenuates RPV symptoms in oats. Beet western yellows subgroup n luteovirus (BWYV) and ST9 associated RNA (ST9a RNA), a member of subgroup I, were assayed for their ability to replicate satRPV RNA. SatRPV RNA accumulated in tobacco protoplasts inoculated with BWYV or BWYV and ST9a RNAs. In contrast, ST9a RNA alone failed to support satRPV RNA replication. Stimulation of BWYV by ST9a RNA resulted in increased levels of satRPV RNA progeny. BWYV encapsidated satRPV RNA but in a form different from that found in BYDV (RPV + PAV) particles. SatRPV RNA was transmitted to shepherd's purse by aphids only in the presence of BWYV and ST9a RNA. SatRPV RNA reduced accumulation of both BWYV helper and ST9a nonhelper RNAs in plants and slightly modulated BWYV and ST9a RNA symptoms. The results showed that satRPV RNA can replicate in both monocotyledonous and dicotyledonous hosts and suggested that the determinants of satRPV RNA replication are contained within the polymerase genes of supporting viruses rather than in structural genes or host plants. 77 Introduction Satellite RNAs (satRNAs) are subviral RNAs that are associated with their helper viruses (Mayo et ai, 1995). The main feature of satRNAs is that they do not contain sufficient information to direct their own replication and encapsidation. They depend on coinfection of a host cell with a supporting helper virus. SatRNAs lack any extensive sequence similarity to the genomic RNA of the helper virus or hosts. SatRNAs are dispensable for helper virus replication but can affect helper virus RNA accumulation and symptoms caused by helper (Collmer and Howell, 1992). SatRNAs are presumably replicated by the viral replicase that consists of helper virus encoded RNA-dependent RNA polymerase (RdRp) and host factors (Quadt et al., 1993). SatRNA of cucumber mosaic virus (CMV) can be rephcated by a partially purified CMV RdRp in vitro (Wu ef al., 1991; Hayes et al., 1992). Putative RdRp open reading firames (ORFs) were the only viral genes required for repUcation of satellite tobacco necrosis virus (Andriessen et al., 1995). Other factors that may influence the ability of any virus to replicate satRNA include the ability of satRNA to move cell-to cell, be encapsidated and spread long distance in a host plant. Therefore, the specificity of satellite replication depends on both the helper virus and host (Roossinck et al., 1992). Luteoviruses have been divided into two subgroups based on the genome organization, serological relationships, and cytopathological properties (Miller., 1995; Mayo and Ziegler- Graff, 1996). Subgroup I includes the PAV and MAV barley yellow dwarf viruses (BYDVs) and others; subgroup II includes the RPV, SGV and RMV BYDVs, beet western yellows virus (BWYV), potato leafiroll virus, and others. BYDV-RPV is a helper for satRPV RNA, a small (322 nt), linear, noncoding satRNA (Miller et al., 1991). SatRPV RNA replicates by a symmetrical rolling circle mechanism (Silver et al., 1994) with a self-cleavage at hammerhead ribozyme structure in both strands (Miller and Silver, 1991). SatRPV RNA reduces RPV helper RNA accumulation and attenuates RPV symptoms in oats. However, BYDV-PAV does not support satRPV RNA replication and the satellite has no effect on PAV RNA accumulation 78 and symptonis caused by the severe mixed infection of RPV and PAV BYDVs in oats (Rasochova and Miller, 1996). The genome of luteoviruses consists of a single-stranded, plus-sense RNA of about 5.6 kb and five or six ORFs (Miller, 1994). While the 3' halves of the luteovirus genomes encoding structural genes share resemblance between the subgroups, the 5' halves of the genomes are very different (Miller et al., 1995). 0RF2, which contains consensus motif of an RdRp in both subgroups (Koonin, 1991), is expressed as a fusion with ORFl by ribosomal frameshifting (Di et al., 1993). The polymerase genes of subgroup I viruses are more closely related to those of diantho-, carmo- and tombusviruses than they are to those of subgroup H luteoviruses. In contrast, subgroup II polymerases are similar to those of sobemovinises (Koonin and Dolja, 1993). Although similar in other properties, BYDV-RPV and BWYV have different host range and aphid vectors (Rochow and Duffiis, 1978). In general, BYDV infects monocotyledonous plants, whereas BWYV infects many families of dicotyledonous plants but under laboratory conditions is able to infect oats (Rochow and Duffiis, 1978; Duffus and Rochow, 1978). However, BYDV isolates were never transmitted to shepherd's purse, a host of BWYV (Rochow and Duffus, 1978). Myzus persicae is the major vector of BWYV, whereas Rhopalosiphum padi is the most efficient vector of BYDV-RPV. The ST9 strain of BWYV differs from other BWYV strains because it encapsidates not only genomic RNA but also a subviral 2,843 nt RNA agent with partial characteristics of satRNA designated BWYV ST9-associated RNA (ST9a RNA; Falk and Duffus, 1984; Chin et al., 1993). ST9a RNA does not code for its own coat protein but encodes putative RNA- dependent RNA polymerase (RdRp). BWYV ST9a RNA polymerase contains two regions of significant homology to the putative RdRp of subgroup I luteoviruses but no homology to BWYV (Chin et al., 1993). BWYV ST9a RNA is therefore assigned to subgroup I luteoviruses. It is capable of independent replication in tobacco protoplasts (Passmore et al., 1993) but depends on BWYV for encapsidation, movement, and aphid transmission. It 79 stimulates accumulation of BWYV genomic RNA (Passmore etal., 1993) and causes more severe symptoms in BWYV-infected shepherd's purse (jCapsella bursa-pastoris) plants (Sanger etal., 1994). In this report we analyze the interaction of subgroups I and II luteoviruses with satRPV RNA in monocotyledonous and dicotyledonous hosts. Our results show that the ability of luteoviruses to serve as effective satRPV RNA helpers is determined by the extent of polymerase gene homologies to BYDV-RPV rather than by the structural genes homologies or host plant. Materials and Methods In vitro transcription Full-length satRPV RNA was transcribed from permuted dimeric cDNA clone pT7Sat (Rasochova and Miller, 1996) that was linearized with EcoRl prior to transcripdon with bacteriophage T7 RNA polymerase using the T7 Megascript kit (Ambion, Austin, TX). The self-cleavage of full-length transcripdon product was induced by incubation in 50 mM Tris, pH 7.5, and 10 mM MgCl2 (Miller and Silver, 1991) at 37°C for one hour. The monomeric self- cleavage product that arose during the incubation was gel-purified, prior to inoculadon (Silver et al., 1994). Full-length BWYV genomic RNA was synthesized from fisiWI-linearized cDNA clone pBW 7120-7A (Passmore et al., 1993) in the presence of cap analogue m^G(5')ppp(5')G (New England Biolabs) by the action of bacteriophage T3 E^^A polymerase as described in Titus et al. (1991). ST9a RNA was transcribed by bacteriophage T7 RNA polymerase from plasmid pST9 106-8 (Passmore et al., 1993) that had been linearized with Xhol using T7 Megascript kit (Ambion, Texas, TX). Final RNA concentrations were determined spectrophotometrically. ^^p.iabeled RNA probes were synthesized by in vitro transcription as described by Titus et a/. (1991) using [a32p]-CTP. Antisense satRPV RNA 80 probe was transcribed with bacteriophage SP6 RNA polymerase from pT7Sat linearized with Hindis.. Sense satRPV RNA probe was synthesized using bacteriophage T7 RNA polymerase on £co/?I-linearized pT7Sat template. Antisense 3'BWYV probe, complementary to the 1.6 kb region at 3' end of BWYV genomic RNA, was transcribed with bacteriophage SP6 RNA polymerase from flamffl-linearized pBW7120-7A. To generate full-length complementary ST9a RNA probe, 5a/I-linearized pST9106-8 was transcribed by bacteriophage SP6 RNA polymerase. Vims propagation, purification, and extraction of virion RNA B YD virus isolates used were IL-PAV (from Anna Hewings, formerly USDA/ARS University of Illinois), and NY-RPV (from Steward Gray, USDA/ARS Cornell University). No satRPV RNA was detected in either of these isolates by northern blot hybridization. Virus propagation was as in Rasochova and Miller (1996). Virions were purified from oat plants and viral RNA extracted by the method of Waterhouse et al. (1986) as modified by Mohan et al. (1995). Electroporation of protoplasts and extraction of total RNA Tobacco {Nicotiana tabacum cv. Xanthi nc) protoplasts were isolated from rapidly growing suspension all cultures as described in Passmore etal. (1993). Protoplasts were electroporated with mock inoculum (electroporation buffer), 10 |ig of BWYV genomic RNA transcript, and/or 2 (Xg of ST9a RNA transcript, and 100 ng of gel-purified monomeric satRPV RNA transcript as described by Passmore et al. (1993). At the indicated times, 1-ml aliquots were removed, cells were collected by centrifiigation, and quick frozen m dry ice/ethanol. Pelleted cells were stored at -70°C. Total RNA was isolated using TRI REAGENT™ kit and recommended RNA extraction procedure (Molecular Research Center, Inc., Cincinnati, OH). 81 Oat (Avena sativa cv. Stout) protoplasts were isolated from cell suspension culture (cell line S226 obtained from Howard Rines, (USDA/ARS, University of Minnesota) as described in Dinesh-Kumar and Miller (1993). Protoplasts were electroporated with either mock inoculum (electroporation buffer) or 100 ng of viral RNA isolated from the mixture of RPV and PAV BYDVs and 50 ng of gel-purified monomeric satRPV RNA transcript as described in Rasochova and Miller (1996). At indicated times, 5-ml aliquots were removed, cells were collected by centrifiigation, quick frozen in liquid nitrogen, and stored at -80°C. Total RNA was isolated by the procedure of Wads worth et al. (1988) as modified by Dinesh-Kumar and Miller (1993). Encapsidadon assay Encapsidation assay was performed as described in Reutenauer etal. (1993). Protoplasts were pelleted 72 hours after inoculation, homogenized in 200 (il of PIPES buffer and incubated for 30 minutes at 37°C. Following incubation in cell lysates, nuclease-resistant RNA was isolated by extraction with 200 pi of TE-equilibrated phenol and mixture of phenol and chloroform (24:1). After centrifiigation, the aqueous phase was removed, and nucleic acids were precipitated by addition of 0.1 vol. of 3M sodium acetate and ethanol to 70%. The precipitated nucleic acids were collected by centrifiigation, washed once with 70% ethanol, dried and resuspended in 20 ml of diethylpyrocarbonate-treated water. SatRPV RNA protected from cellular nucleases was considered to be encapsidated in coat protein of helper virus. As a control, 100 ng of gel-purified satRPV RNA monomeric transcript was added to uninoculated, lysed protoplasts, prior to 30 minutes incubation at 37°C. Encapsidated RNA was analyzed by northern blot hybridization. 82 Purification of virions from tobacco protoplasts Virions, used in aphid transmission experiments, were extracted from protoplasts 5 days after electropcraticn. The protoplasts were collected by centrifiigation and homogenized in 5 vol. of 100 mM potassium phosphate buffer, 10 mM glycine, pH 7.0 in 3-ml glass homogenizator as described in Sanger et al. (1994). The homogenate was centrifliged at 7650 g for 10 minutes. The supernatant was removed and centrifuged at 175,000 g for 1.5 hours. The pelleted virions were resuspended in 300 ml of 100 mM potassium phosphate buffer, 10 mM glycine, pH 7.0, and 10% sucrose prior to aphid feeding. Aphid feeding on artificial membranes, inoculation of plants and isolation of total leaf RNA Aphid acquisition of virus was through Parafilm™ membranes as described by Falk et al. (1979). Three plants were used per treatment. After feeding for 24 hours, aphids {Myzus persicae) were transferred to shepherd's purse plants {Capsella bursa-pastoris) at the four-leaf stage for 72 hours transmission access period. Aphids were then killed and plants grown in a greenhouse. Subsequent routine transmission of virus from plant to plant by aphids was as described by Falk et al. (1989). Plants were observed for symptoms, photographed, and assayed for viral RNA accumulation by northem blot hybridization. Total RNA was isolated from leaves of individual plants or pooled sample of several plants 21 days after aphid- inoculadon by the method of Wadsworth etal. (1988) as modified by Dinesh-Kumar and MUler(1993). RNA analysis Denaturing 1% agarose gel electrophoresis and northem blot hybridization was as performed in Rasochova and Miller (1996). Equal amounts (approx. 5 |ig) of total RNA were loaded in each lane. This was determined by spectrophotometry and ensured by ethidium 83 bromide staining of ribosomal RNA prior to northern blot hybridization. Denaturing 5% polyacrylamide/TM urea (19:1) and 6% polyacrylamide/TM urea (24:1) electrophoresis, electroblotting, and northern blot hybridization was as described by Passmore and Bruening (1993). 5x10^ or 1x10^ cpm/ml of radioactive probe of hybridization buffer were used per hybridization experiment. Blots that were used in subsequent hybridizations were stripped by boiling in O.lx SSC and 0.1% SDS prior to re-probing with another probe as described in Rasochova and Miller (1996). Results Subgroup n BWYV supports replication of satRPV RNA in tobacco cells To test the specificity of satRPV RNA association with subgroup II luteoviruses we analyzed the ability of BWYV to support satRPV RNA replication since the putative RdRp of BWYV shares an extensive homology with that of BYDV-RPV (Mayo and Ziegler-Graff, 1996; Fig. 1). Tobacco protoplasts were co-inoculated with infectious transcripts of monomeric satRPV EiNA and BWYV genomic RNA in the presence or absence of ST9a RNA. The satRPV RNA progeny was detected in cells inoculated with BWYV with or without ST9a RNA and the (+) strand satRPV RNA accumulated to higher levels than the (-) strand (Fig. 2, compare panel A and B, lanes 5-10). Bands representing monomeric linear and circular forms and linear dimers that are formed during rolling circle replication were identified in both strands of satRPV RNA by comparing their migration pattern on acrylamide gels with different cross- linking and by comparing them to corresponding forms of satRNA of tobacco ringspot virus (satTRS V RNA; data not shown). SatTRS V RNA is similar to satRPV RNA in length (satTRSV RNA - 359 nt, satRPV RNA - 322 nt) and mode of replication (synmietrical rolling circle). Bands of higher molecular weight detected in satRPV RNA progeny represent higher linear multimers and possibly circles. SatRPV RNA accumulated to much higher levels in the presence of BWYV ST9a RNA in the inoculum (Fig. 2, panels A and B, compare lanes 5-7 to 84 8-10). No signal was detected in mock-inoculated protoplasts indicating the absence of nonspecific hybridization. We conclude that satRPV RNA is supported by BWYV (subgroup n luteovirus) and can replicate in dicotyledonous tobacco cells. ST9a RNA is not required for satRPV RNA replication but its presence in the inoculum resulted in the increased accumulation of satRPV RNA progeny. Subgroup I ST9a RNA failed to support satRPV RNA replication in tobacco protoplasts The enhanced accumulation of satRPV RNA in the presence of ST9a RNA may be due to the stimulation of BWYV genomic RNA replication by ST9a RNA (Passmore et al., 1993; Sanger et aL, 1994) or due to the ability of ST9a RNA itself to replicate the satellite. The ability of ST9a RNA to serve as satRPV RNA helper was tested in tobacco protoplasts. A different gel system was used to resolve the replication products (denaturing 1% agarose). This system does not allow resolution of linear and circular forms of satRPV RNA but is sufficient for detection of satRPV RNA progeny. Monomeric satRPV RNA was used as inoculum and thus the detection of dimers and higher multimers would be considered the evidence of replication. No replication of satRPV RNA was detected at 24 and 48 hours post co-inoculation with ST9a RNA transcript (Fig. 3, panel A, lanes 7-9). As expected, ST9a RNA replicated to high levels in the absence of BWYV genomic RNA (Fig. 3, panel B, lanes 4-9). The replication of ST9a RNA was independent of presence of satRPV RNA in the inoculum (Fig. 3, panel B, compare lanes 4-6 to 7-9). Residual satRPV RNA (Fig. 3, panel A, lane 7) and ST9a RNA (Fig. 3, panel B, lanes 4 and 7) from the inoculum could be detected at 0 hours post inoculation. There was no cross-hybridization between ST9a RNA and satRPV RNA (Fig. 3, panel A, lanes 4-5) and no nonspecific hybridization to tobacco RNA (Fig. 3, panels A and B, lanes 1-3). We conclude that ST9a RNA (subgroup I luteovirus) does not serve as a helper for satRPV RNA replication. Therefore, the enhanced accumulation of 85 satRPV RNA in cells inoculated with satRPV RNA, BWYV genomic and ST9a RNAs is an indirect result of increased BWYV genomic RNA replication caused by the presence of ST9a RNA. SatRPV RNA is encapsidated in mostly circular form in BWYV capsids but as a linear form in BYDV particles SatRNAs depend on helper virus not only for replication but also for encapsidation. We tested the ability of BWYV to encapsidate satRPV RNA by comparing total and nuclease resistant (encapsidated) RNA from tobacco protoplasts inoculated with satRPV RNA monomer and BWYV genomic RNA with or without BWYV ST9a RNA transcripts. Encapsidated RNA is resistant to degradation by cellular nucleases but RNA that is not encapsidated is readily degraded upon cell lysis in the absence of phenol. First we determined how well nonencapsidated satRPV RNA transcript resists the lysed cell treatment. 100 ng of satRPV RNA monomer was added to the noninoculated, lysed tobacco cells and incubated for 30 minutes at 37°C prior to RNA extraction and northern blot hybridization. Treated satRPV RNA transcript was completely degraded and undetectable (Fig. 4, panel A, lane 2). SatRPV RNA encapsidated in BWYV particles was clearly resistant to lysate treatment (Fig. 4, panel A, lanes 4 and 6). A circular, monomeric, (+) stranded satRPV RNA was the predominant encapsidated form as indicated by its lysate resistance. No (-) stranded form of satRPV RNA was encapsidated (Fig. 4, lanes 4 and 6, compare panels A and B). ST9a RNA greatly enhanced levels of encapsidated satRPV RNA (Fig. 4, panel A, lane 4). In contrast, very low, almost undetectable levels of encapsidated satRPV RNA accumulated in the absence of ST9a RNA in the inoculum (Fig. 4, panel A, lane 6). In contrast to BWYV, BYDV was reported previously to encapsidate satRPV RNA predominantly as a linear monomer in the virion preparations from oat plants infected with BYDV and satRPV RNA (Miller et al., 1991). To compare encapsidated forms of satRPV 86 RNA extracted from plants and protoplasts we analyzed total and nuclease-resistant satRPV RNA from oat protoplasts infected with BYDV (RPV + PAV) virion RNA and satRPV RNA monomeric transcript. In the total RNA preparation, monomeric linear and circular forms of satRPV RNA were detected in both strands (Fig. 5, lane 1). Large quantities of (+) stranded linear monomers were encapsidated (Fig. 5, panel A, lane 2) and no encapsidated (-) strand was detected (Fig. 5, panel B, lane 2). The bands of molecular weight lower than a Unear monomer resistant to degradation by cellular nucleases were present in both strands. These bands were also detected in the virion preparation from plants (Miller et al., 1991) but their origin is unknown. Very weak cross-hybridization between (-) strand satRPV RNA and the antisense satRPV RNA probe could be detected (Fig. 5, panel A, lane 3) as reported previously (Silver et al., 1994) but did not affect the interpretation of results. We conclude that satRPV RNA was encapsidated predominately as a linear monomer in oat protoplasts infected with satRPV RNA and BYDV (PAV + V). This is in agreement with the original preparation of virion RNA from oat plants. SatRPV RNA could be encapsidated by BWYV capsids in tobacco protoplasts but in a form of circles rather than linear molecules. The amounts of encapsidated satellite correlated with the accumulation of satRPV RNA progeny in BWYV vs BWYV + ST9a RNA inoculated cells. Aphid transmission of satRPV RNA from protoplasts to shepherd's purse plants required both BWYV and ST9a RNA Aphid transmission of satRPV RNA to plants requires that both helper and satRNA are encapsidated (Rasochova and Miller, 1996). Because BWYV was able to encapsidate satRPV RNA we attempted to transmit satRPV RNA to shepherd's purse {Capsella bursa-pastoris) plants by feeding aphids {Myzus persicae) on virus extracted from tobacco protoplasts. Protoplasts were inoculated with satRPV RNA monomer and BWYV genomic RNA transcript in the presence or absence of ST9a RNA. Virus particles were purified by differential 87 centrifugatioa and aphid feeding performed as described in Materials and Methods. Monomeric and multimeric forms of satRPV RNA accumulated in plants three weeks after inoculation with aphids that had fed on extract from protoplasts infected with satRPV RNA, BWYV, and ST9a RNAs (Fig. 6, lanes 3-6). In contrast, no replication of satRPV RNA was detected in plants inoculated with aphids that had acquired virus from protoplasts infected with satRPV RNA and BWYV in the absence of ST9a RNA (Fig. 6, lanes 7-10). In this inoculum combination the concentration of encapsidated satRPV RNA was probably too low to facilitate satellite transmission (Fig. 4, panel A, lane 6). No satRPV RNA signal was detected in uninoculated (Fig. 6, lane 1) or mock inoculated plants (Fig. 6, lane 2). We conclude that sufficient titer of encapsidated satRPV RNA was critical for successful transmission of satRPV RNA from protoplasts to plants by aphids. Once transmitted, satRPV RNA was capable of replication and movement in BWYV and ST9a RNA infected dicotyledonous shepherd's purse plants. Effect of satRPV RNA on accumulation of BWYV and ST9a RNA and disease symptoms in shepherd's purse plants Although no transmission of satRPV RNA was detected in the presence of BWYV helper only, we were able to analyze the effect of satRNA on BWYV helper and ST9a nonhelper RNA accumulation in shepherd's purse plants infected with BWYV, ST9a RNA, and satRPV RNA. Plants infected with BWYV only accumulated low levels of BWYV genomic RNA (gRNA; Fig. 7, panel C, lanes 4-6). The accumulation of BWYV gRNA, subgenomic RNA 1 (sgRNA 1), and especially subgenomic RNA 2 (sgRNA 2) was enhanced dramatically by the presence of abundant ST9a RNA (Fig. 7, panel C, lanes 7-9). Large quantities of sgRNA of ST9a RNA were also detected (Fig. 7, panel D, lanes 7-9). The detection of BWYV sgRNA 2 (== 700 nt) and sgRNA of ST9a RNA (= 4(X) nt) that have no apparent coding capabilities was reported previously in infected plants (Falk et ai, 1989 ; 88 Passmore et al., 1993). As expected, satRPV RNA replicated in plants co-infected with BWYV and ST9a RNA as indicated by the abundant monomeric and multimeric forms (Fig. 7, panel B, lanes 10-12). The accumulation of helper BWYV genomic and sgRNAs was reduced in the presence of satRPV RNA (Fig. 7, panel C, compare lanes 10-12 to 7-9). The levels of the full-length ST9a RNA were also significantiy lowered but the accumulation of sgRNA of ST9a RNA remained unchanged by the presence of satRPV RNA (Fig. 7, panel D, compare lanes 10-12 to 7-9). No signals were detected in mock inoculated plants (Fig. 7, panels B, C and D, lanes 1-3). Shepherd's purse plants infected with BWYV only developed very mild or no symptoms (Fig. 8, plants C and D). Plants infected with BWYV and ST9a RNA with or without satRPV RNA exhibited yellowing and stunting typical of BWYV and ST9a RNA infection (Fig. 8, plant E and F). Compared to plants infected with BWYV and ST9a RNA, those which also contained satRPV RNA showed less yellowing and slightly more vigorous growth (Fig. 8, compare plants E and F). However, these differences were not as obvious as the differences detected between plants infected with BWYV and those infected with BWYV and ST9a RNA (Fig. 7, compare plants C and D, E, and F). Our observations indicate that although satRPV RNA reduced the accumulation of both helper and nonhelper RNAs in shepherd's purse plants, it attenuated the symptoms induced by BWYV and ST9a RNA only slightly. Discussion Specificity of satRPV RNA - helper virus polymerase interaction In this work, we have used infectious transcripts of subgroup I and II luteoviruses to determine helper virus specificity of satRPV RNA replication in divergent monocotyledonous and dicotyledonous hosts. We were able to correlate the differential satRPV RNA activation by subgroup I and n luteoviruses with the features of their polymerase genes. We had shown 89 previously that subgroup H luteovirus, BYDV-RPV, supports satRPV RNA replication in oat protoplasts and plants but subgroup I BYDV-PAV does not (Silver et al, 1994; Rasochova and Miller, 1996). Here, we demonstrated that subgroup H BWYV, which shares an extensive sequence homology with BYDV-RPV in the putative polymerase genes (59%, Mayo and Ziegler-Graff, 1996), is also able to support replication of satRPV RNA in tobacco protoplasts but is less efficient at replicating the satellite compared to BYDV-RPV (Rasochova and Miller, 1996). It is conceivable that successfiil replication of the satRNA depends upon the efficiency of the specific interactions between the satRNA and the replicase. SatRPV RNA is probably best adapted to the BYDV-RPV replicase. Thus, the less efficient amplification of satRPV RNA by BWYV may be caused by weaker interactions between the satRPV RNA and other sobemo-Iike replicases. BWYV may also lack parts of the genome which may stimulate satRPV RNA multiplication. In contrast, satRPV RNA did not replicate in cells co-infected with ST9a RNA that shares extensive polymerase homology with BYDV-PAV and other members of the carmo-Iike supergroup (Tombusviridae; Chin et al., 1993) but not with the sobemo-like polymerases of subgroup n luteovinises (Miller et al, 1995). The failure of BYDV-PAV and ST9a RNA to replicate satRPV RNA most probably resulted from the inability of the carmo-Iike replicases to recognize satRPV RNA replication origin. The incompatibility between satRPV RNA and subgroup I luteovinises was not due to an inefficient movement of satRPV RNA as reported for other satellites (Moriones et al., 1992; Moroines et al., 1994) because the potential helper viruses were assayed in protoplasts. Although ST9a RNA was unable to support satRPV RNA, the accumulation of satellite increased dramatically in the presence of ST9a RNA. The levels of BWYV gRNA, which serves as the replicase mRNA, were reported to be ten times higher in infections which also contained ST9a RNA (Sanger et al., 1994). Therefore, the stimulation of satRPV RNA replication can be explained by more abundant supply of BWYV 90 replicase in the presence of ST9a RNA than may be available in cells infected by the BWYV alone The efficiency of satRPV RNA replication may be, to some extent, controlled by the ability of helper viruses to support satRPV RNA encapsidation. We have shown that satRPV EWA can be encapsidated by BWYV capsids in form of circular monomer. In contrast, B YDV encapsidates linear forms of satRPV RNA (Miller et al., 1991 and this work) Differential encapsidation has not been reported for any other satRNA. The reasons for it are unknown and may include both helper virus (BYDV vs BWYV) and host plant (oat vs tobacco) factors. In contrast to BYDV, BWYV alone encapsidated very low levels of satRPV RNA which resulted in lack of aphid transmission of satRPV RNA from protoplasts to shepherd's purse plants. However, when ST9a RNA was included in the inoculum, large quantities of encapsidated satRPV RNA were detected and the satRPV RNA was successfully transmitted to plants. The increase in satRPV RNA encapsidation can be explained by more abundant supply of BWYV coat protein in the presence of ST9a RNA (Passmore et al., 1993). Effect of satRPV RNA on symptoms and viral RNAs accumulation in plants Although satRPV RNA greatly reduced both helper BWYV and nonhelper BWYV ST9a RNAs accumulation, it attenuated symptoms induced by BWYV and ST9a RNA in shepherd's purse plants only slighdy. Similar failure of satRPV RNA to significantly ameliorate symptoms was also observed in oats infected with RPV and PAV BYDVs (Rasochova and Miller, 1996). It is likely that the effect of satRNA was silenced in mixed infections by the presence of more severe ST9a RNA and/or by the synergistic interactions between BWYV and ST9a RNA (Sanger et al., 1994). The suppressive effect of satRPV RNA on BWYV gRNA accumulation is similar to that reported for BYDV-RPV but the effect on the nonhelper ST9a RNA differs from that observed for B YDV-PAV (Rasochova and Miller, 1996). In contrast to BYDV-PAV, the yield of ST9a RNA was reduced significantly in the 91 presence of satRPV RNA. Because ST9a RNA does not support satRPV EINA replication and replicates independently of BWYV (Passmore et al., 1993), the competition of satRPV RNA with ST9a RNA for replication is not likely to be the cause. However, because ST9a RNA does not encode its own coat protein and depends on BWYV for encapsidation (Passmore et al., 1993), it is conceivable that this relatively higher titer RNA must compete with satRPV RNA and BWYV gRNA for available BWYV capsids to ensure its spread by aphid transmission. The observed reduction in the accumulation of encapsidated fiiU-length ST9a RNA but not subgenomic RNA, that is known not to be encapsidated, is consistent with this hypothesis. Thus, in our system satRPV RNA acts as parasite of not only the helper virus but also has negative, indirect effect on non-supporting but helper virus-associated RNA. Host plant requirements of satRPV RNA Several reports indicate that satRNA replication depends not only on the helper virus but also on the host plant (Roossinck et al., 1992). BYDV and BWYV have different host ranges (Martin and D'Arcy, 1995). Although BWYV isolates were reported to occasionally infect oat plants, BYDV isolates were never transmitted to shepherd's purse (Rochow and Duffus, 1978). In addition, BYDV-RPV did not replicate in tobacco protoplasts (data not shown). We demonstrated that satRPV RNA replicates effectively in both tobacco protoplasts and shepherd's purse plants. This indicates that the requirements for the replication of helper virus and satRNA may not be the same and that satRNA may interact with a suitable helper in divergent monocotyledonous and dicotyledonous hosts. Only two other satellites, satRNA of luceme transient streak sobemovinis (satLTSV RNA) and satRNA of bamboo mosaic virus, were reported to replicate in both monocotyledonous and dicotyledonous plants (Sehgal et al, 1993; Lin and Hsu, 1994). SatRPV and satLTSV RNAs are the only known satellites that replicate in hosts which their original helper virus does not infect (Sehgal et al., 1993). 92 We have shown that at least two subgroup n luteoviruses (BYDV-RPV and BWYV) are able to facilitate satRPV RNA replication in divergent hosts. What particular feature(s) of these viruses allows them to replicate satRPV RNA is not known. Our results suggest that RdRp genes that are required for BWYV and BYDV-PAV RNA replication in protoplasts (Reutenauer et ai, 1993; Mohan et al., 1995) may be sufficient. The helper virus range of satRPV RNA may not be limited to subgroup n luteoviruses. Sobemoviruses, that show significant sequence similarities to subgroup H luteoviruses in their polymerase genes (Koonin, 1991; Koonin and Dolja, 1993) and support many small circular rolling circle satRNAs (Francki, 1985; Roossinck et al., 1992), may also be able to replicate satRPV RNA. The ability of sobemoviruses to support satRPV RNA and the role of RdRp of subgroup II luteoviruses in satRPV RNA replication will be investigated. Acknowledgments The authors thank Jamal Buzayan for technical advice and Randy Beckett for virus preparation. This work was fiinded by USDA National Research Initiative grant number 94373030469. References Andriessen, M., Meulewaeter, F., and Comelissen, M. (1995). Expression of tobacco necrosis virus open reading frames 1 and 2 is sufficient for the rephcation of satellite tobacco necrosis virus. Virology 212, 222-224. Chin, L.-S., Foster, J. L., and Falk, B. W. (1993). The beet western yellows virus ST9- associated RNA shares structural and nucleotide sequence homology with carmo-like viruses. Virology 192, 473-482. Collmer, C. W., and Howell, S. H. (1992). Role of satelUte RNA in the expression of symptoms caused by plant viruses. Annu. Rev. Phytopathol. 30,419-442. 93 D'Arcy, C. J., and Burnett, P. A., Eds. (1995). "Barley Yellow Dwarf: 40 Years of Progress." APS Press, St. Paul, MN. Di, R., Dinesh-Kumar, S. P., and Miller, W. A. (1993). Translational frameshifting by barley yellow dwarf vims RNA (PAV serotype) in Escherichia coli and eukaryotic cell-free extracts. Mol. Plant-Microbe Interact. 6,444-452. Dinesh-Kumar, S. P., and Miller, W. A. (1993). Control of start codon choice on a plant viral RNA encoding overlapping genes. Plant Cell 5, 679-692. Falk, B. W., Duffus, J. E., and Morris, T. J. (1979). Transmission, host range, and serological properties of the viruses that cause lettuce speckles disease. Phytopathology 69, 612-617. Falk, B. W., and Duffus, J. E. (1984). Identification of small single- and double-stranded RNAs associated with severe symptoms in beet western yellows virus-infected Capsella bursa-pastoris. Phytopathology 1^, 1224-1229. Falk, B. W., Chin, L.-S., and Duffus, J. E. (J1989). Complementary DNA cloning and hybridization analysis of beet western yellows luteovirus RNAs. J. Gen. Virol. 70, 1301- 1309. Francki, R. I. B. (1985). Plant virus satellites. Ann. Rev. Microbiol. 39, 151-174. Hayes, R. J., Tousch, D., Jacquemond, M., Pereira, V. C., Buck, K. W., and Tepfer, M. (1992). Complete replication of a satellite RNA in vitro by a purified RNA-dependent RNA polymerase. /. Gen. Virol. 73, 1596-1600. Koonin, E. V. (1991). The phylogeny of RNA-dependent RNA polymerases of positive- strand RNA viruses. J. Gen. Virol. 72, 2197-2206. Koonin, E. V., and Dolja, V. V. (1993). Evolution and taxonomy of positive strand RNA viruses: ImpUcations of comparative analysis of amino acid sequences. Crit. Rev. Biochem. Mol. Biol. 28, 375-430. 94 Lin, N.-S., and Hsu, Y.-H. (1994). A satellite EINA associated with bamboo mosaic potexvirus. Viro/ogy 202, 707-714. Martin, R. R., Keese, P. K., Young, M. J., Waterhouse, P. M., and Gerlach, W. L. (1990). Evolution and molecular biology of luteoviruses. Ann. Rev. Phytopathol. 28,341-363. Martin, R. R., and D'Arcy, C. J. (1995). Taxonomy of barley yellow dwarf viruses. In "Barley Yellow Dwarf: 40 Years of Progress" (C. J. D'Arcy and P. Burnett, Eds.), pp. 203-216. APS Press, St. Paul. Mayo, M. A., and Ziegler-Graff, V. (1996). Recent developments in luteovirus molecular biology. Adv. Virus Res. 46,413-460. Miller, W. A., Hercus, T., Waterhouse, P. M., and Gerlach W. L. (1991). A satellite RNA of barley yellow dwarf virus contains a novel hammerhead structure in the self-cleavage domain. Virology 183,711-720. Miller, W. A., and Silver, S. L. (1991). Alternative tertiary structure attenuates self-cleavage of the ribozyme in the satellite RNA of barley yellow dwarf virus. Nucleic Acids Res. 19, 513-520. Miller, W. A. (1994). Luteoviruses. In "Encyclopedia of Virology" (R. G. Webster and A. Granoff, Eds.), pp. 792-798. Academic Press, London. Miller, W. A., Dinesh-Kumar, S. P., and Paul, C. P. (1995). Luteovuois gene expression. Crit. Rev. Plant Sci. 14, 179-211. Mohan, B. R., Dinesh-Kumar, S. P., and Miller, W. A. (1995). Genes and c/j-acting sequences involved in replication of barley yellow dwarf virus-PAV RNA. Virology 212, 186-195. Moriones, E., Diaz, I., Rodriguez-Cerezo, E., Fraile, A., and Garcia-Arenal, F. (1992). Differential interactions among strains of tomato aspermy virus and satellite RNAs of cucumber mosaic virus. Virology 186,475-480. 95 Moriones, E., Diaz, L, Femandez-Cuartero, B., Fraile, A., Burgyan, J., and Garcia-Arenal, F. (1994). Mapping helper virus functions for cucumber mosaic virus satellite EINA with pseudorecombinants derived from cucumber mosaic and tomato aspermy viruses. Virology 205,574-577. Passmore, B. K., and Bruening, G. (1993). Similar structure and reactivity of satellite tobacco ringspot virus RNA obtained from infected tissue and by in vitro transcription. Virology 197, 108-115. Passmore, B. K., Sanger, M., Chin, L.-S., Falk, B. W., and Bruening, G. (1993). Beet western yellows virus-associated RNA: an independently replicating RNA that stimulates virus accumulation. Proc. Natl. Acad. Sci. USA 90, 10168-10172. Quadt, R., Kao, C., Browning, K., Herhberger, R., and Ahlquist, P. (1993). Characterization of a host protein associated with brome mosaic virus RNA-dependent RNA polymerase. Proc. Natl. Acad. Sci. USA 90, 1498-1502. Rasochova, L., and Miller, W. A. (1996). Satellite RNA of barley yellow dwarf-RPV virus reduces accumulation of RPV helper virus RNA and attenuates RPV symptoms in oats. Mol. Plant-Microbe Interact. 9, 646-650. Reutenauer, A., Ziegler-Graff, V., Lot, H., Scheidecker, D., Guilley, H., Richards, K., and Jonard, G. (1993). Identification of beet westem yellows luteovirus genes implicated in viral replication and particle morphogenesis. Virology 195, 692-699. Rochow, W. F. and Duffiis, J. E. (1978). Relationships between barley yellow dwarf and beet westem yellows viruses. Phytopathology 68,51-58. Roossinck, M. J., Sleat, D,. and Palukaitis, P. (1992). Satellite RNAs of plant viruses: structures and biological effects. Microbiol. Rev. 52, 265-279. 96 Sanger, M., Passmore, B., Falk, B. W., Bruening, G., Ding, B., and Lucas, W. J. (1994). Symptom severity of beet western yellows virus strain ST9 is conferred by the ST9- associated RNA and is not associated with virus release from the phloem. Virology 200, 48-55. Sehgal, O. P., Sinha, R. C., Gellatly, D. L., Ivanov, L, and AbouHaidar, M. G. (1993). Replication and encapsidation of the viroid-like satellite RNA of lucerne transient streak virus are supported in divergent hosts by cockfoot mottle virus and turnip rosette virus. J. Gen. Virol. 74, 785-788. Silver, S. L., Rasochova, L., Dinesh-Kumar, S. P., and Miller W. A. (1994). Replication of barley yellow dwarf virus satellite RNA transcripts in oat protoplasts. Virology 198, 331- 335. Titus, D. E., Ed. (1991). "Protocols and Application Guide," pp. 58-64. Promega, Madison, WI. Wadsworth, G. J., Redinbaugh, M. G., and Scandalios, J. G. (1988). A procedure for small- scale isolation of plant RNA suitable for RNA blot analysis. Anal. Biochem. 172, 279- 283. Waterhouse, P. M., Gerlach, W. L., and Miller, W. A. (1986). Serotype-specific and general luteovirus probes from cloned cDNA sequences of barley yellow dwarf virus. J. Gen. Virol. 67, 1273-1281. Wu G.-S., Kaper, J. M., and Jaspars, E. M. J. (1991). Replication of cucumber mosaic virus satellite RNA in vitro by an RNA-dependent RNA polymerase from virus-infected tobacco. FEBSUtt. 292, 2\3-l\6. Andriessen, M., Meulewaeter, F., and Comelissen, M. (1995). Expression of tobacco necrosis virus open reading frames 1 and 2 is sufficient for the replication of satellite tobacco necrosis virus. Virology 212, 222-224. 97 Chin, L.-S., Foster, J. L., and Falk, B. W. (1993). The beet western yellows virus ST9- associated RNA shares structural and nucleotide sequence homology with carmo-like viruses. Virology 192,473-482. CoUmer, C. W., and Howell, S. H. (1992). Role of satellite RNA in the expression of symptoms caused by plant viruses. Anna. Rev. Phytopathol. 30,419-442. D'Arcy, C. J., and Burnett, P. A., Eds. (1995). "Barley Yellow Dwarf: 40 Years of Progress." APS Press, St. Paul, MN. Di, R., Dinesh-Kumar, S. P., and Miller, W. A. (1993). Translational frameshifting by barley yellow dwarf virus RNA (PAV serotype) in Escherichia coli and eukaryotic cell-free extracts. MoL Plant-Microbe Interact. 6,444-452. Dinesh-Kumar, S. P., and Miller, W. A. (1993). Control of start codon choice on a plant viral RNA encoding overlapping genes. Plant Cell 5, 679-692. Falk, B. W., Duffiis, J. E., and Morris, T. J. (1979). Transmission, host range, and serological properties of the viruses that cause lettuce speckles disease. Phytopathology 69, 612-617. Falk, B. W., and Duffus, J. E. (1984). Identification of small single- and double-stranded RNAs associated with severe symptoms in beet western yellows virus-infected Capsella bursa-pastoris. Phytopathology 1A, 1224-1229. Falk, B. W., Chin, L.-S., and Duffus, J. E. (J 1989). Complementary DNA cloning and hybridization analysis of beet westem yeUows luteovirus RNAs. /. Gen. Virol. 70, 1301- 1309. Francki, R. I. B. (1985). Plant virus satellites. Ann. Rev. Microbiol. 39, 151-174. Hayes, R. J., Tousch, D., Jacquemond, M., Pereira, V. C., Buck, K. W., and Tepfer, M. (1992). Complete replication of a satellite RNA in vitro by a purified RNA-dependent RNA polymerase. J. Gen. Virol. 73, 1596-1600. 98 Koonin, E. V. (1991). The phytogeny of RNA-dependent RNA polymerases of positive- strand RNA viruses. J. Gen. Virol. 72,2197-2206. Koonin, E. V., and Dolja, V. V. (1993). Evolution and taxonomy of positive strand RNA viruses: Implications of comparative analysis of amino acid sequences. Crit. Rev. Biochem. Mol. Biol. 28, 375-430. Lin, N.-S., and Hsu, Y.-H. (1994). A satellite RNA associated wi± bamboo mosaic potexvirus. Virology Martin, R. R., Keese, P. K., Young, M. J., Waterhouse, P. M., and Gerlach, W. L. (1990). Evolution and molecular biology of luteoviruses. Ann. Rev. Phytopathol. 28, 341-363. Martin, R. R., and D'Arcy, C.J. (1995). Taxonomy of barley yellow dwarf viruses. In "Barley Yellow Dwarf: 40 Years of Progress" (C. J. D'Arcy and P. Burnett, Eds.), pp. 203-216. APS Press, St. Paul. Mayo, M. A., and Ziegler-Graff, V. (1996). Recent developments in luteovirus molecular biology. Adv. Virus Res. 46,413-460. Miller, W. A., Hercus, T., Waterhouse, P. M., and Gerlach W. L. (1991). A satellite RNA of barley yellow dwarf virus contains a novel hammerhead structure in the self-cleavage domain. Virology 183,711-720. Miller, W. A., and Silver, S. L. (1991). Alternative tertiary structure attenuates self-cieavage of the ribozyme in the satellite RNA of barley yellow dwarf virus. Nucleic Acids Res. 19, 513-520. Miller, W. A. (1994). Luteoviruses. In "Encyclopedia of Virology" (R. G. Webster and A. Granoff, Eds.), pp. 792-798. Academic Press, London. Miller, W. A., Dinesh-Kumar, S. P., and Paul, C. P. (1995). Luteovirus gene expression. Crit. Rev. Plant Sci. 14, 179-211. 99 Mohan, B. R., Dinesh-Kuraar, S. P., and Miller, W. A. (1995). Genes and cij-acting sequences involved in replication of barley yellow dwarf virus-PAV RNA. Virology 212, 186-195. Moriones, E., Diaz, I., Rodriguez-Cerezo, E., Fraile, A., and Garcia-Arenal, F. (1992). Differential interactions among strains of tomato aspermy virus and satellite RNAs of cucumber mosaic virus. Virology 186,475-480. Moriones, E., Diaz, I., Femandez-Cuartero, B., Fraile, A., Burgyan, J., and Garcia-Arenal, F. (1994). Mapping helper virus functions for cucumber mosaic virus satellite RNA with pseudorecombinants derived from cucumber mosaic and tomato aspermy viruses. Virology 205,574-577. Passmore, B. K., and Bruening, G. (1993). Similar structure and reactivity of satellite tobacco ringspot virus RNA obtained from infected tissue and by in vitro transcription. Virology 191, 108-115. Passmore, B. K., Sanger, M., Chin, L.-S., Falk, B. W., and Bruening, G. (1993). Beet western yellows virus-associated RNA: an independently replicating RNA that stimulates virus accumulation. Proc. Natl. Acad. Sci. USA 90, 10168-10172. Quadt, R., Kao, C., Browning, K., Herhberger, R., and Ahlquist, P. (1993). Characterization of a host protein associated with brome mosaic virus RNA-dependent RNA polymerase. Proc. Natl. Acad. Sci. USA 90, 1498-1502. Rasochova, L., and Miller, W. A. (1996). Satellite RNA of barley yellow dwarf-RPV virus reduces accumulation of RPV helper virus RNA and attenuates RPV symptoms in oats. Mol. Plant-Microbe Interact. 9, 646-650. Reutenauer, A., Ziegler-Graff, V., Lot, H., Scheidecker, D., Guilley, H., Richards, K., and Jonard, G. (1993). Identification of beet western yellows luteovirus genes implicated in viral replication and particle morphogenesis. Virology 195,692-699. 100 Rochow, W. F. and Duffus, J. E. (1978). Relationships between barley yellow dwarf and beet western yellows viruses. Phytopathology 68,51-58. Roossinck, M. J., Sleat, D,. and Palukaitis, P. (1992). Satellite RNAs of plant viruses: structures and biological effects. Microbiol Rev. 52,265-279. Sanger, M., Passmore, B., Faik, B. W., Bruening, G., Ding, B., and Lucas, W. J. (1994). Symptom severity of beet western yellows virus strain ST9 is conferred by the ST9- associated RNA and is not associated with virus release from the phloem. Virology 200, 48-55. Sehgal, O. P., Sinha, R. C., Gellatly, D. L., Ivanov, I., and AbouHaidar, M. G. (1993). Replication and encapsidation of the viroid-like satellite RNA of lucerne transient streak virus are supported in divergent hosts by cockfoot mottle virus and turnip rosette virus. J. Gen. Virol. 74, 785-788. Silver, S. L., Rasochova, L., Dinesh-Kumar, S. P., and Miller W. A. (1994). Replication of barley yellow dwarf virus satellite RNA transcripts in oat protoplasts. Virology 198,331- 335. Titus, D. E., Ed. (1991). "Protocols and Application Guide," pp. 58-64. Promega, Madison, WI. Wadsworth, G. J., Redinbaugh, M. G., and Scandalios, J. G. (1988). A procedure for small- scale isolation of plant RNA suitable for RNA blot analysis. Anal. Biochem. 172, 279- 283. Waterhouse, P. M., Gerlach, W. L., and Miller, W. A. (1986). Serotype-specific and general luteovirus probes from cloned cDNA sequences of barley yellow dwarf virus. J. Gen. Virol. 67, 1273-1281. Wu G.-S., Kaper, J. M., and Jaspars, E. M. J. (1991). Replication of cucumber mosaic virus satellite RNA in vitro by an RNA-dependent RNA polymerase from virus-infected tobacco. FEBSLett. 292,213-216. 101 Subgroup I BYDV-PAV 22K 39^KgS2POLiBBa •• 50-S8K «.7K I 1 S»«OK 17-22K m gRNA tgRNAI •gRNA 2 •gRNA 3 ST9a RNA 23K 56K 3.4K I 1 Baamaa sj ' gRNA Subgroup 11 BYDV-RPV BWYV 66-71K 22K I 1 I I 0 I n SO^K 28-29K 67-72K 17-19K •gRNA Figure 1. Diagram of the genome organization of the luteovirus subgroups. Boxes indicate ORFs, numbered as in Martin et al. (1990), with coding capacity shown in kilodaltons (K). Black-shaded ORFs are conserved between subgroups. CP indicates coat protein gene, POL indicates putative polymerase gene. Cross-hatched POL ORFs have homology to Carmovirales. Stripped POL ORFs are homologous to sobemovirales. Unshaded ORFs have no significant similarity to ORFs of any other virus, with the exception of protease motif in ORFl of subgroup n. Known positions of subgenomic RNAs are shown below the genomes. 102 Inoculum: A Probe: CM satRPV(-) LM2 LM Hours: 0 48 72 0 48 72 0 48 72 B CM Probe: LM2 satRPV(+) LM Lane: 1 23456789 10 Figure 2. Northern blot hybridization analysis of total RNA from N. tabacum protoplasts. Protoplasts were inoculated with mock inoculum (lanes 2-4), BWYV RNA + ST9a RNA + satRPV RNA monomer (lanes 5-7), and BWYV RNA + satRPV RNA monomer (lanes 8-10), as indicated above the lanes. Lane 1 contains 10 ng of (+) strand satRPV RNA linear monomer as a standard. SatRPV RNA monomer was electrophoretically purified prior to electroporation. Protoplasts were collected at 0,48, and 72 hours after inoculation, as indicated below the panel A. Equal amounts of total RNA, verified by ethidium bromide staining of ribosomal RNAs, were loaded in each lane. RNA was fractionated on two 6% polyacrylamide/7M urea gels, electroblotted to nylon membrane and hybridized with antisense satRPV RNA probe (panel A) and sense satRPV RNA probe (panel B), as indicated on the left. Equal amounts of probe (1 x 10^ cpm/ml) were used in each hybridization Exposures were at - 70°C with one intensifying screen for 2 hours (panel A) and 3 hours (panel B). Autoradiography revealed bands for linear monomeric (LM, 322 nt), circular monomeric (CM, 322 nt), and linear dimeric (LM2, 644 nt) forms of satRPV RNA, as indicated on the right. 103 Inoculum: ST9a Mock ST9a +satRPV Probe: satRPV(-) LM smsiSiiaasB Hours: 0 24 48 0 24 48 0 2448 Probe: aRNA ST9a(-) Lane: 123456789 Figure 3. Northern blot hybridization of total RNA from N. tabacum protoplasts. Protoplasts were electroporated with mock inoculum (lanes 1-3), ST9a RNA transcript (lanes 4-6), and ST9a RNA + satRPV RNA monomer (lanes 7-9), as indicated above the lanes. Sat RPV RNA monomer was electrophoretically purified from in vitro transcription and self- cleavage reactions prior to electroporation. Protoplasts were collected at 0,24, and 48 hours after electroporation, as indicated bellow the panel A. Equal amounts of total RNA, verified by ethidium bromide staining of ribosomal RNAs, were loaded in each lane. RNA was fractionated on a denaturing 1% agarose gel, blotted to nylon membrane and hybridized to antisense satRPV RNA probe (panel A) and antisense ST9a RNA probe (panel B), as indicated on the left. One blot was used, stripped and re-probed with the second probe. Equal amounts of probe (5 x 10^ cpm/ml) were used in each hybridization. The autoradiographic exposures were at -80°C with one intensifying screen for 24 hours (panel A) and 1 hour (panel B). Mobility of ST9a RNA (aRNA; 2,843 nt) and expected mobility of linear monomeric satRPV RNA (LM, 322 nt) are indicated on the right. 104 Probe: satRPV(-) i'*-LU Lane: 1 2 Probe: satRPV(+) RNA preparation: 4, \ %•-'Q J %\ Figure 4. Northern hybridization of total (lanes 3 and 5) and encapsidated (Virion, lanes 4 and 6) satRPV RNA extracted from tobacco protoplasts 72 hours post inoculation with BWYV RNA + ST9a RNA + satRPV RNA monomer (lanes 3-4) and BWYV RNA+ satRPV RNA (lanes 5-6), as indicated above the lanes. Encapsidated RNA was extracted from lysed tobacco cells. Lane 1 contains 10 ng of monomeric satRPV RNA (LM, 322 nt) as a marker. Lane 2 represents 100 ng of satRPV RNA monomer added to uninoculated lysed tobacco cells and subjected to 30 min incubation in 37°C prior to RNA extraction. RNA was fractionated on two 6% polyacryamide/7M urea gels, electroblotted to nylon membrane and hybridized with antisense (panel A) and sense satRPV RNA probe (panel B), as indicated on the left. Equal amounts of probe (1 x 10^ cpm/ml) were used in each hybridization. Exposures were at -70°C with one intensifying screen for 2 hours (panel A) and 3 hours (panel B). Autoradiography revealed bands for linear monomer (LM, 322 nt) and circular monomeric form (CM, 322 nt) of satRPV RNA, as indicated on the right. 105 A Probe: satRPV(-) Lane; 1 2 3 B Probe: satRPV(+) Figure 5. Northern blot analyses satRPV RNA encapsidated in B YDV capsids in oat protoplasts. Total (lane 1) and encapsidated (Virion, lane 2) RNAs were extracted 72 hours post inoculation with BYDV (RPV + PAV) + satRPV RNAs. Encapsidated RNA was protected from cellular nucleases after 30 minutes incubation of lysed cells at 37°C prior to RNA extraction. Lanes 3 [LM (-)] and 4 [LM (+)] contain 30 ng of gel-purified, monomeric (-) and (+) strand satRPV RNA transcript (322 nt). RNA was fractionated on a 5% polyacrylamide/TM urea gel, electroblotted to nylon membrane and hybridized with antisense satRPV RNA probe (panel A) and sense satRPV RNA probe (panel B), as indicated on the left. The same blot was used in both hybridizations, stripped and re-probed with the second probe. Equal amounts of probe (5 x 10^ cpm/ml) were used in each hybridization reaction. Exposures were at -80°C with one intensifying screen for 30 min (panel A) and 1 hour (panel B). Positions of linear monomer (LM, 322 nt) and circular monomeric form (CM, 322) of satRPV RNA are indicated on the right. 106 Probe: satRPV(-) M3 Lane: 1 23456789 10 Figure 6. Northern blot hybridization oftotal RNA from shepherd's purse plants. Plants were aphid-inoculated with mock inoculum Gane 2), BWYV + ST9a RNA + satRPV RNA (lanes 3-6), and BWYV + satRPV RNA (lanes 7-10), as indicated above the lanes. Lane 1 contains total RNA isolated from uninoculated plant. RNA was isolated from leaves of each individual plant three weeks after inoculation. Equal amounts of RNA were loaded in each lane, verified by ethidium bromide staining of ribosomal RNAs. RNA was fractionated on denaturing 1% agarose gel, blotted to nylon membrane and hybridized with the antisense satRPV RNA probe (5 x 10^ cpm/ml), as indicated on the left. The blot was exposed at -80°C with one intensifying screen for 2 hours. Autoradiography revealed bands for satRPV RNA monomer (M, 322 nt), dimer (M2, 644 nt), trimer (M3, 966 nt), and higher multimers (Mx), as indicated on the right. 107 '—II—ir^nn— Probe: satRPV(-) Probe: f: gRNA 3BWr/f-)* sgRNAI sgRNA2 Probe: ST9a(-) aRNA sgRNA Lane: i 2 3 4 5 6 7 8 9101112 Figure 7. Analyses of total RNA from shepherd's purse. Plants were aphid-inoculated with mock (lanes 1-3), BWYV (lanes 4-6), BWYV + ST9a RNA (lanes 7-9), and BWYV + ST9a RNA + satRPV RNA (lanes 10-12). RNA was isolated in triplicates three weeks after inoculation. Equal amounts of RNA were loaded, verified by ethidium bromide staining of ribosomal RNAs (Fig. 7, panel A). RNA was fractionated on one denaturing 1% agarose gel, blotted, and hybridized with 5 x 10^ cpm/ml of antisense satRPV RNA (panel B), 3'BWYV RNA (panel C), and ST9a RNA probes (panel D). The exposures at -80°C with one intensifying screen for 1 hour (panel B), 6 hours (panel C), and 4 hours (panel D) revealed bands for satRPV RNA monomer (M^, 322 nt), dimer (M2, 644 nt), trimer (M3,966 nt), BWYV gRNA (5,641 nt), BWYV sgRNAl (« 2,600 nt), BWYV sgRNA 2 (= 700 nt), ST9a RNA (aRNA, 2843 nt), and ST9a sgRNA (sgRNA, == 400 nt). 108 Figure 8. Biological assay of satRPV RNA effect oa the severity of BWYV and ST9a RNA symptoms in shepherd's purse plants. Plants A, C, and E were inoculated by Myzus persicae aphids that had acquired the virus from tobacco protoplasts. Plants D and F were inoculated by aphids that had acquired the virus from previously inoculated plants. Plant B was inoculated with nonvirulent aphids. Inoculum: plant A - mock, plant C - BWYV + satRPV RNA, plant D - BWYV, plant E - BWYV + ST9a RNA + satRPV RNA, plant F - BWYV + ST9a RNA. Northern blot hybridization analyses of the total RNA from each individual plant revealed that plant D was infected with BWYV only. The photograph was taken four weeks after inoculation. One plant with typical symptom expression was selected from each group of inoculated plants. 109 CHAPTERS. GENERAL CONCLUSIONS Conclusions The ultimate goal of the research on satRPV RNA is to completely understand how satellite, helper virus and host plant components interact and how a virus which normally replicates only itself can replicate heterologous satRNA with such a high efficiency but ignore all other RNAs present in a host cell. SatRNAs are one of the smallest known molecules capable of replication. Most of them do not have or regulate genes required for their replication. Thus, satRNAs can be used as model molecules to study minimal sequences and structures that are required for multiplication of genetic material as well as to study evolutionary processes. The better understanding of this complex three-way interaction between the satellite, helper virus and host can help us leam how viruses themselves replicate. In addition, satRNAs often alter disease symptoms induced by their helper viruses. Thus, better understanding of how satellites affect replication of helper viruses and modulate symptoms may lead to the development of new control strategies for viral disease which could benefit agriculture in general. SatRPV RNA is the only known satellite of luteoviruses. It is a linear, ssRNA molecule of 322 nt, with no significant sequence homology to its helper, BYDV-RPV genomic RNA. SatRPV RNA sequence must contain all the information that allows it to act as an enzyme, be specifically replicated by its helper virus, be encapsidated and modulate disease symptoms induced by the helper virus. The goal of my research was to answer some basic biological questions regarding satRPV RNA helper virus specificity, host range and symptom expression. I showed that: (i) satRPV RNA seems not to be mechanically transmissible; (ii) satRPV RNA can be transmitted to plants by aphids that were allowed to feed on partially purified virus extract from protoplasts; 110 (iii) the concentration of encapsidated helper virus and satRNA has to be above a threshold level to facilitate aphid transmission of both the helper virus and satellite to plants; (iv) satRPV RNA attenuates symptoms induced by BYDV-RPV in oats; (v) satRPV RNA has no effect on disease symptoms caused by the severe mixed infection of BYDV-RPV and PAV; (vi) satRPV RNA significantly decreases accumulation of B YDV-RPV helper virus RNA in both protoplasts and plants but has no effect on accumulation of nonhelper BYDV-PAV RNA; (vii) BWYV, member of subgroup II luteoviruses supports satRPV RNA replication, encapsidation and movement in dicotyledonous protoplasts and plants; (viii) BYDV-PAV and ST9a RNA, subgroup I luteoviruses, do not support satRPV RNA replication; (ix) satRPV RNA is encapsidated predominantly as a linear monomer in B YDV particles but as a circle in BWYV virions; (x) satRPV RNA can be transmitted to shepherd's purse plants but only in the presence of BWYV and ST9a RNA; (xi) satRPV RNA reduces accumulation of both BWYV helper and ST9a nonhelper RNAs in plants; (xii) satRPV RNA slightly ameUorates severe symptoms induced by the mixed infection of BWYV and ST9a RNA in shepherd's plants. The results suggest that possible mechanisms of symptom attenuation by satRPV RNA involve competition between the helper virus and its satellite for replication and encapsidation rather than inducing the host defense response or inhibition of cell-to-cell and long distance movement within the plant host. Based on the differences and/or similarities between subgroup I (BYDV-PAV, ST9a RNA) and EI (BYDV-RPV, BWYV) luteoviruses it seems Ill likely that the determinants of satRPV RNA replication are contained within the polymerase genes of supporting viruses rather than in structural genes or host plans. The results indicate that the host plant requirements for replication of satRPV RNA and its helper may not be the same and that satRPV RNAs can multiply in divergent monocotyledonous and dicotyledonous hosts co-infected with a suitable helper. Recoiimiendation for Future Research The basic questions that remain to be answered include: (i) What components must be provided in trans for replication of satRPV RNA; (ii) What must the satellite itself provide to ensure its replication? The future research should be oriented towards identifying the minimal or optimal genes and sequences in the genomes of supporting viruses necessary for satRPV RNA multiplication in protoplasts and in the whole plant. This can be accomplished by large deletions or point mutagenesis of infectious helper virus clones, and by testing mutated helper viruses for their ability to replicate satRPV RNA in protoplasts or by transient expression of helper virus genes from expression plasmids in protoplasts co-inoculated with satRPV RNA transcripts. It is likely that the putative RdRp proteins encoded by ORFl and 2 of BYDV-RPV and BWYV are the only virus-encoded proteins essential for satRPV RNA replication (Reutenauer et al., 1993; Andriessen et al., 1995; Mohan et al., 1996). However, it is possible that requirements for helper virus and satellite replication are not the same. Once the minimal genes/sequences are identified, random, point or small deletion mutants can be constructed to determine the specific structures, nucleotides or amino acids in the helper virus that confer the specificity of satRPV RNA replication. It would be interesting to identify helper virus mutants that lose the ability to replicate satRPV RNA without losing the ability to replicate its own genomic RNA or mutants with increased or reduced ability to support satRPV RNA. 112 It is not known if satellites move from cell-to-cell or long distance as naked molecules without assistance from their helper viruses, in association with movement proteins of their helper viruses, or in an encapsidated form. To investigate if any movement functions normally provided by the helper virus are necessary for satRPV RNA infectivity, transgenic plants expressing the optimal helper virus replication- (but not movement-) related genes could be inoculated with satRPV RNA transcripts. This system would eliminate the need for aphid transmission and overcome the phloem limitation of luteoviruses, as every plant cell would be expressing genes required for satellite replication. If replication, cell-to-cell movement and a long distance spread of satRPV RNA is observed, it would suggest that satellites, similarly to viroids, can move in the absence of helper virus movement and/or capsid proteins. If cell-to- cell movement is observed but no systemic infection occurs, it would indicate that satRPV RNA moves from cell-to-cell without the need for supporting virus gene products but the long distance movement requires satellite encapsidation and/or some movement functions provided by the helper. If no movement is observed in plants but satRPV RNA can replicate effectively in protoplasts isolated from transgenic plants expressing replication-related proteins, it would suggest that some movement fiinctions supplied by the helper virus are missing and/or encapsidation is required for satRPV RNA infectivity at the whole plant level. The reasons for differential encapsidation observed for satRPV RNA in oats co-infected with BYDV (RPV + PAV) helper and in tobacco co-infected with BWYV helper are unknown. The possible causes include those determined by differences in BYDV and BWYV helper virus coat proteins or those determined by plant hosts. Because BYDV-RPV does not replicate in tobacco protoplasts, it is not possible to test these possibilities directly. On the other hand, BYDV-PAV can replicate in tobacco (unpublished observation) but does not support repUcation of satRPV RNA (Silver et al., 1994). Based on the similarities between coat protein sequences of BYDV-RPV and B YDV-PAV (46%; Mayo and Ziegler-Graff, 1996), B YDV-PAV and BWYV (44%; Mayo and Ziegler-Graff, 1996), or BYDV-RPV and BWYV (63%; Mayo and 113 Ziegler-Graff, 1996) BYDV-PAV itself may support satRPV RNA encapsidation. Because linear forms of encapsidated satRPV RNA were detected in BYDV-RPV as well as in BYDV (RPV+PAV) particles, BYDV-PAV most probably encapsidates linear satRPV RNA molecules rather than circles. Therefore, it could be possible to inoculate tobacco protoplasts with satRPV RNA, BWYV mutant that is unable to express coat protein but can replicate satRPV RNA, and BYDV PAV that cannot replicate satRPV RNA but provides coat protein for satRPV RNA encapsidation. The form of satRPV RNA encapsidated in BYDV-PAV capsids could be assayed by northem blot hybridization of nuclease-resistant RNA. The detection of linear encapsidated forms would suggest that BWYV coat protein rather than some unknown host plant factor is involved in or promotes circularization of satRPV RNA and vice versa. On the other hand, if no encapsidated satRPV RNA is detected it would indicate that BYDV-PAV coat protein is unable to encapsidate satRPV RNA. Furthermore, this would suggest that specific signals exist within the satRPV RNA sequence that direct satRPV RNA encapsidation and that satRNAs are not encapsidated simply because of their high titer in infected cells. The replicase homology hypothesis can be investigated further by testing the ability of viruses with more or less distantly related polymerases to support satRPV RNA replication. Potato leaf roll virus (PLRV) is another subgroup n luteovirus likely to replicate satRPV RNA because its polymerase gene shows 58% similarity to P2 of BYDV-RPV and 60% similarity to P2 of BWYV (Mayo and Ziegler-Graff, 1996). Moreover, the range of helper viruses that support satRPV RNA replication may extend beyond subgroup H luteoviruses. Sobemoviruses may serve as satRPV RNA helpers because their polymerases are related to those of subgroup II luteoviruses (Koonin and Dolja, 1993) and some of them are known to support rolling circle satRNAs (Roossinck et al., 1992). Southern bean mosaic sobemovirus and cocksfoot mosaic sobemovirus (CfMV) were shown to replicate satLTSV RNA in both dicotyledonous and monocotyledonous hosts. In addition, the polymerase gene of CfMV, which is, in contrast to other sobemoviruses, expressed by -1 ribosomal frameshifting, shows 114 about 55% amino acid sequence homology to polymerases of subgroup n luteoviruses, BYDV-RPV and BWYV (Makinen et al., 1995). These viruses can be tested for their ability to replicate satRPV RNA in protoplasts of divergent hosts, including members of family Poaceae, Leguminosae, Chenopodiaceae, Solanaceae and Brassicaceae. TRSV nepovirus could be also tested as a potential satRPV RNA helper. Its polymerase gene is more distantly related to subgroup n luteoviruses but still falls into the same supergroup identified by Koonin and Dolja (1993). Moreover, TRSV supports multiplication of satTRSV RNA, that replicates by the same rolling circle mechanism as satRPV RNA (Bruening et al., 1991). If some of these viruses support satRPV RNA in protoplasts, their ability to support satellite replication, cell-to-cell and long distance movement on the whole plant level can be tested and the effect of satRPV RNA on disease symptoms monitored. The transmission to whole plants would require simply rubbing the partially purified virus from protoplasts on plant surface with abrasive, as sobemoviruses and nepoviruses are mechanically transmissible. Another goal includes identification of sequences and/or structures within satRPV RNA that are necessary for satellite infectivity. These include sequences and/or structures involved in self-cleavage, ligation, replication (replication origin), encapsidation (origin of assembly), and possibly cell-to-cell movement. To map these sequences specific deletion and site-directed mutagenesis as well as random mutagenesis and evolutionary approaches can be used and mutant satRPV RNA transcripts can be tested in protoplasts co-inoculated with helper virus and/or in transgenic plants expressing helper virus replication-related genes identified previously. Once the replication origin of satRPV RNA is identified, it could be used as a promoter to drive the expression of useful genes, such as those conferring resistance to plant viruses, in transgenic plants. The RNA-templated transcription of such genes would be switched on upon the presence of attacking virus that is able to recognize satRPV RNA-derived promoter sequence. Such approach could have a great potential because of the following: 115 (i) it would eliminate the danger of mutations changing the protective satellite to a form that exacerbate symptoms with the same helper virus and host, or chance of variable effect of satellite on symptoms depending on the host and helper virus combination when the satellite sequence itself is used as a resistance gene in transgenic plants. (ii) Although satRNAs have usually relatively narrow range of helper viruses, my results indicate that viruses with polymerases similar to those of subgroup n luteoviruses are able to recognize satRPV RNA promoter sequences and replicate the satellite in divergent monocotyledonous and dicotyledonous hosts. BYDV and BWYV luteoviruses are economically important pathogens, worldwide. Traditional breeding for resistance to those viruses has met with little success and the approaches that are used to control these virases are limited to spraying large quantities of insecticides to reduce the number of aphid vectors. Another subgroup n luteovirus, PLRV, is the most important viral pathogen of potatoes which renders the infected tubers unusable. If sobemoviruses or TRSV nepovirus are found to be able to replicate satRPV RNA, the potential use of satRPV RNA-derived promoters for control of viral diseases could be expanded to these viruses as well. The results of my research suggest that satRPV RNA is best adapted to BYDV-RPV. This seems to be a result of evolutionary processes that led to the selection of an optimal satRPV RNA sequence with respect to the efficiency of the replication by its helper virus. BWYV can replicate satRPV RNA but is less efficient than BYDV-RPV. It would be interesting to determine if satRPV RNA can adapt to BWYV as well as it did to BYDV-RPV and evolve into more efficient template in the presence of BWYV. This could be accomplished by repeated passaging of the satellite in tobacco protoplasts co-infected with BWYV to allow enough mutations to accumulate during satellite replication for evolution of highly efficient satRPV RNA template. The rate of mutation could be increased by inoculating randomly mutagenized satRPV RNA transcripts (Cadwell and Joyce, 1992) and by repeating cycles of in 116 vitro replication without selection (PGR) and in vivo replication with selection for efficient replication in tobacco protoplasts. The use of protoplasts would eliminate the selection for encapsidation and movement and allow rapid testing of many satRPV RNA mutants as each cell is an isolated replication unit The progeny representing the most efficient satRPV RNA templates would be amplified and sequenced. 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