Details of DPV and References
DPV NO: 396 April 2003
Species: Subterranean clover stunt virus | Acronym: SCSV
Subterranean clover stunt virus
P. W. G. Chu CSIRO Plant Industry, GPO Box 1600, Canberra, ACT, 2601, Australia
H. J. Vetten BBA, Institut f. Pflanzenvirologie, Mikrobiologie und biologische Sicherheit, Messeweg 11-12 D-38104 Braunschweig, Germany
- Main Diseases
- Geographical Distribution
- Host Range and Symptomatology
- Transmission by Vectors
- Transmission through Seed
- Transmission by Grafting
- Transmission by Dodder
- Nucleic Acid Hybridization
- Stability in Sap
- Properties of Particles
- Particle Structure
- Particle Composition
- Properties of Infective Nucleic Acid
- Molecular Structure
- Genome Properties
- Relations with Cells and Tissues
- Ecology and Control
The disease caused was first described by Grylls & Butler (1956). The causal virus was first isolated and described by Chu & Helms (1988).
- Clover stunt virus (O'Loughlin, 1958).
The virus is the type member of the genus Nanovirus, (family Nanoviridae). The virus genome consists of at least 6 distinct circular single-stranded (ss) DNA components each of about 1 kb in size with each encapsidated in a separate icosahedral particle of 17-19 nm diameter. Each DNA component appears to possess only one major gene and to contain all signals required for replication and gene expression. The virus particle contains a major protein of Mr 19,000.
The natural host range seems restricted to legumes. Like all other nanoviruses, it is transmitted only by aphids in a non-propagative circulative manner and for which a virus-encoded helper factor appears necessary.
It causes an economically important disease of pasture and grain legumes in Australia (Fig. 1, Fig. 2, Fig. 3, Fig. 4, Fig. 5) (Grylls & Butler, 1956; O'Loughlin, 1958; Harvey, 1958; Grylls, 1972; Chu et al., 1995). Subterranean clover stunt disease (SCSD) is currently estimated to cause average forage and seed losses in subterranean clover of about 31% and 35% respectively, resulting in an estimated loss of c. AUD$53.5 million to the Australian wool industry (Chu et al., 1995).
The virus causes severe reduction in growth and premature death of affected subterranean clover plants (Fig. 1, Fig. 2) with individual plant yield losses of up to 65% (Grylls & Butler, 1956; O'Loughlin, 1958; Anonymous, 1976). Infected plants are much less competitive and are more susceptible to root and crown rots, resulting in high mortality rates. As crop infections tend to be clumped and plants infected early usually die, concentration of SCSD in certain parts of a paddock can lead to bare ground or a displacement of clover by weeds. Field studies showed that when the incidence of SCSD was 13-80%, the forage yield of total plant matter was reduced by 30-76% with a corresponding reduction in yield of the clover component of 35-81%. Clover seed yields were also reduced by 58-94%. A disease incidence of 6-20% early in the growing season reduces seedling regeneration in the following season by 30-48% and increases the areas of weed and bare ground by 7-43%.
The virus also infects peas and beans and peas (Fig. 4, Fig. 5), causing a disease known as top yellows, tip yellows or leafroll (Letham, 1982). During the 1950s and 1960s, the virus was responsible for major losses of 86 to 100% in vegetable and seed crops of beans and peas throughout southern Australia annually (Smith, 1966; Grylls, 1972; Letham, 1982). Repeated crop failures and lack of resistant cultivars have resulted in reduced cropping in these regions and increased the need for spraying against the insect vector. Other potentially important susceptible or alternative hosts include annual Medicago species (O'Loughlin, 1958), lupins, lucerne, lentil, chickpea and soybean.
Severe disease epidemics in subterranean clover and various legume crops have also been attributed to dual infections of the virus with Bean yellow mosaic virus (Grylls & Peak, 1969; Grylls, 1972).
Currently restricted to Australia. In southern Australia in the late 1980s (Chu et al., 1991) the virus was prevalent in all regions with average regional incidences ranging from 13-51% in winter and 19-52% in spring.
Milk vetch dwarf virus (MDV), a closely related nanovirus, is reported only from Japan. By contrast, Faba bean necrotic yellows virus (FBNYV), a close relative of MDV, occurs in many Near Eastern, Middle Eastern, and North African countries as well as in Ethiopia and Spain.
Host Range and Symptomatology
The virus is restricted to species in the family Fabaceae (Johnstone & Chu, 1996). Natural hosts include Trifolium pratense (Fig. 3), T. glomeratum, T. repens, Medicago hispida, M. lupulina, M. minima and Wisteria sinensis (Grylls & Butler, 1959). Experimentally, the virus infects a wide range of legumes after transmission with by the aphid vector, Aphis craccivora. These include species in the genera Arachis, Astralagus, Cicer, Crotalaria, Desmodium, Dolichos, Lens, Medicago, Melilotus, Phaseolus, Pisum, Onobrychis, Trifolium, Trigonella, Vicia, and Vigna. Virus isolates vary slightly in experimental host range (Johnstone & McLean, 1987).
The relative severity of disease symptoms on different hosts depends on the virus isolate rather than the host and, with mild isolates, infected plants show a partial recovery. Experimentally infected lucerne and lupins are symptomless.
- Diagnostic hosts
- Subterranean clover, cv. Mount Barker. Symptoms appear as early as 7 days but usually within 9-14 days and occasionally as late as 28 days post-inoculation, due possibly to low inoculum level. Initially, symptoms appear as mild marginal chlorosis of young leaves followed by puckering, stunting and chlorosis of new leaves and reddening of mature and old leaves (Fig. 2). The severity of symptoms depends on the virus isolate.
- Propagation species
- Pisum sativum cv. Greenfeast, is used for virus purification. Infected plants are stunted, showing leaf rolling, chlorosis of leaf margins and tips, and reduced leaf size (Fig. 5).
- Assay species
- Subterranean clover (T. subterraneum) is the main assay host and is suitable for transmission by vectors.
- Maintenance host
- Subterranean clover using A. craccivora as the vector.
- Useful non-hosts
- Useful non-hosts to distinguish the virus from some virus species in the family Luteoviridae that infect pasture and grain legumes are Beta vulgaris, Gomphrena globosa and Lactuca sativa.
Several distinct variants are reported (Grylls & Peak, 1969; Price & Blackstock, 1974; Johnstone & Mclean, 1987). Price & Blackstock (1974) described the Gippsland isolate from Eastern Victoria, the Western Victorian and the Northern Victorian isolates. The Gippsland isolate is significantly more severe than the other two isolates in all four susceptible cultivars of subterranean clover tested, and is the only isolate that infected the resistant cultivars Howard and Tallarook. The milder isolates appear to be most common in all regions in most years and while the severe isolate occurs less frequently.
Comparing distinct regional virus isolates (Chu et al., 1989) showed that most could be classified into one of three groups based on the severity of their symptoms on different hosts:
Isolates causing severe stunting that occur mainly in wetter temperate regions of the South Eastern regions of New South Wales and Victoria. These isolates induce a severe shortening of internodes and petioles, a reduction in leaf size and numbers and a consequent reduction in stem or stolon length and number. Plants affected by these isolates usually develop severe chlorosis of leaf margins, puckering in young leaves followed by reddening (especially in subterranean clover) of leaf margins, and sometimes a dark greening of leaf blades in matured leaves and subsequently premature browning and drying off of old leaves.
Isolates causing a moderate reduction in internode, petiole and stolon lengths without any obvious reduction in leaf number so that plants become bunchy in appearance. Infected plants usually develop yellowish-green cupped leaves with intermittent vein-clearing without leaf reddening but older leaves may continue to develop more severe chlorosis and subsequently dry off.
Isolates producing mild yellowing in young leaves initially but leaves later become either symptomless or show only a mild yellowing with intermittent vein-clearing but with no obvious reduction in any growth parameters. These isolates appear to be found mainly in the drier Mediterranean regions of South Eastern South Australia and Western Victoria.
Transmission by Vectors
In nature, the virus is transmitted only by aphids in a persistent manner. At least four aphid species have been reported to transmit, viz., Aphis craccivora (Grylls & Butler, 1956), A. gossypii (Grylls & Peak, 1969), Macrosiphum euphorbiae (Smith, 1966) and Myzus persicae (Grylls & Butler, 1956; O'Loughlin, 1958). However, virus isolates recovered by Grylls and Peak (1969) using aphid species other than A. craccivora, may not have been SCSV as A. craccivora was unable to transmit them. A. craccivora is by far the most efficient vector and is able to transmit the virus after short acquisition and inoculation feeding periods of 30 min each, and a latent period of less than 24 h (Grylls & Butler, 1956, 1959), although a LP50 of 8-12 h appears to be more typical for a nanovirus. This aphid species is able to transmit all known virus isolates and is the most abundant of the four species in the field (Grylls, 1972). All instars transmit but transmission is most efficient when virus is acquired in the larval stage. Aphids remain viruliferous after moulting and may remain infective for life, but the virus does not multiply in the insect vector. Transmission is therefore in a circulative manner. Aphids can transmit the virus only after feeding on intact infected tissues but not from extracts of infected leaf or from purified virus preparations. This suggests that a virus-encoded (helper) factor is required for aphid transmission as was shown for another nanovirus, Faba bean necrotic yellows virus (Franz et al., 1999).
Experimentally, the virus is also transmitted by grafting (O'Loughlin, 1958) but not by mechanical inoculation of plant sap and or by three species of leafhoppers (Grylls & Butler, 1959).
Transmission through Seed
The virus was not transmitted through seed of infected French bean, subterranean clover, pea or soybean (O'Loughlin, 1958; Smith, 1966; Grylls & Butler, 1959; Chu et al., 1995).
Transmission by Dodder
The virus is moderately immunogenic. Antisera produced to the virus in rabbits reach titres of 1/128 in agar gel double diffusion tests. Due to the small size of the particle and the low concentration of the virus in infected plants, the only suitable diagnostic serological test is ELISA and tissue blot immunoassay. ELISA detects all the known virus isolates and is about 5 times more sensitive than nucleic acid hybridization, and detects virus in crude sap extracts at dilutedions of over 1/625 for the homologous isolate, but with a dilution end point of 1/25 for some heterologous (usually very mild) isolates. ELISA was unable to detect the virus in viruliferous aphids (Chu et al., 1995).
ISEM, Western blotting, dot-blotting and agar gel double diffusion tests are suitable only for detecting virus in purified preparations but and are less sensitive than ELISA. No monoclonal antibodies to the virus have been produced (Chu et al., 1995)
All known virus isolates are related serologically and cross-hybridise with each other. Phenotypically distinct isolates may differ in symptom severity, host range, serological reactivity and the electrophoretic mobility of their DNA components in polyacrylamide gel electrophoresis. New South Wales isolates A, F and AA infect all 54 lines of subterranean clover tested, but a Tasmanian isolate SCS6 was unable to infect 14 of these lines. Only some isolates infect lupin. Cross-protection experiments performed with subterranean clover indicated that a mild virus isolate can protect plants against severe symptoms from infection by a severe isolate (Chu et al., 1995).
Sequence analysis of PCR products of DNA-S from different virus isolates found significant nucleotide sequence variations between them in both coding and non-coding regions. The non-coding region of DNA-S is the most variable, differing by up to 34% between isolates whereas coding regions differed by up to 15% resulting in up to 10% amino acid changes (Boevink et al., 1995). This may reflect the variable serological reactivity of the isolates. There was no association between sequence variation in DNA-S and specific differences in biological properties and in the geographic origins of the isolates.
PCR amplification of the DNA from different isolates using a conserved set of primers to the common region, identified a polymorphism in the size of some minor bands in addition to a constant major band of c. 1 kb for every isolates, suggesting the presence of different sized subgenomic DNAs in these isolates. Virus isolates may also be characterised by the presence or absence of specific satellite-like DNA components. However, the biological variation of virus isolates could not be related to a specific size or intensity of the subgenomic DNAs and/or the presence or absence of a specific satellite-like DNA (Chu et al., 1995).
These results indicate the very variable nature of the virus and it may exist as a broad continuum of closely related isolates. The large number of genomic components suggests the huge potential for the virus to change through genetic reassortment. The complete sequence analysis of the genomic DNAs of several isolates may determine the full extent of variation and the taxonomic relationships amongst them.
Based upon the analysis of the available sequence information, 6 to 8 distinct circular viral ssDNA components are thought to be integral parts of the nanovirus genome (Timchenko et al., 1999, 2000). In the 6 genomic DNAs described from SCSV, the virus shares an overall nucleotide sequence identity of about 55% with the three other known nanovirus species (Faba bean necrotic yellows virus [FBNYV], Milk vetch dwarf virus [MDV] and Banana bunchy top virus [BBTV]) and takes an intermediate position between BBTV and the two very closely related nanovirus species FBNYV and MDV (Katul et al., 1998; Randles et al., 2000). The most highly conserved gene product of nanoviruses is the master replication initiator (M-Rep) protein, which, in SCSV, is encoded by DNA-R (formerly C8; Timchenko et al., 2000). The M-Rep protein of the virus shares amino acid sequence identities of about 83% and 55% with those of FBNYV (and MDV) and BBTV, respectively (Timchenko et al., 2000). Further significant levels of amino acid sequence identity among protein homologs encoded by all known nanoviruses (> 45%) have been observed only for the putative nuclear shuttle protein encoded by DNA-N (formerly DNA C4 in SCSV). In all the other gene products, including the coat protein (CP), the virus shares significant levels of amino acid sequence identity with those of FBNYV and MDV (> 44%) but not with those of BBTV (< 25%) (Katul et al., 1998; Randles et al., 2000).
Antisera to the virus and to FBNYV cross-react weakly with each other in Western immuno-blots and immuno-electron microscopy, but not at all in DAS-ELISA (Katul et al., 1993; Chu et al., 1995). Whereas the majority of 19 monoclonal antibodies to FBNYV reacted with MDV, only one of them reacted with the SCSV isolates F and I, indicating that the serological relationship between FBNYV and MDV is close but remote between SCSV and FBNYV (Katul et al., 1993; Franz et al., 1996). This is confirmed by amino acid sequence identities of 53.5 to 55% for the coat proteins of SCSV and FBNYV (Katul et al., 1998), and of 83% for those of FBNYV and MDV (Sano et al., 1998).
Stability in Sap
Particles remain intact after more than 16 h at 37°C but aphid transmissibility is lost as soon as the virus is released from the intact tissue of host plants such as subterranean clover (Chu & Helms, 1988).
Virions are best purified from pea plants inoculated by A. craccivora with a moderately severe isolate. Leaves and stems materials from plants showing conspicuous symptoms (Fig. 5) 6-8 weeks after inoculation are pulverised in liquid nitrogen and homogenised in 0.1 M sodium citrate buffer, pH 6.0, containing 0.1% 2-mercaptoethanol. The extract is incubated at 28°C for 16 h in 3% (v/v) Celluclast and 0.5% Driselase as described by Chu & Helms (1988). After clarification with chloroform, virus particles are precipitated with 8% polyethylene glycol mol. wt 6000 and 0.3 M NaCl and resuspended in 0.1 M phosphate buffer, pH 7.4, containing 10 mM EDTA and 1% (v/v) Triton X-100. The crude virus preparations are purified further by multiple cycles of high (200,000g, 2 h) and low speed centrifugation and 10-40% (w/v) sucrose density gradient centrifugations at 200,000g for 70-90 min, in a Beckman SW50 rotor (Chu and Helms, 1988). Partially purified virus is resuspended in 10 mM phosphate buffer, pH 7.4, containing 0.01 mM EDTA. The virions sediment as a single, sometimes broad, band in sucrose density gradients and are similar in size to phytoferritin. The virus nature of the particles is verified by biochemical analyses, such as nucleic acid electrophoresis, ELISA, and electron microscopy of the sucrose gradient fractions containing the partially purified virus (Chu & Helms, 1988). The yield and purity of virus particles depends on both on the virus isolate and the propagation host used, ranging from 0.5 to 5 mg/kg tissue. For mild isolates, virus yield is lower and two cycles of sucrose density gradient centrifugation followed by isopycnic centrifugation in caesium sulphate may be necessary to obtain homogeneous preparations. Phytoferritin is a major contaminant of virion preparations.
Properties of Particles
Virus particles are stable in 2% ammonium molybdate and in caesium sulphate but are less stable in caesium chloride. They sediment as a single component in sucrose and caesium sulphate density gradients.
Buoyant density: 1.24 g.cm-3 in caesium sulphate and 1.34 g.cm-3 in caesium chloride.
Particle weight: Approximately 1.6 million daltons.
Extinction coefficient: 3.6 at A260 nm (1 mg/ml, 1 cm light path) (corrected for light-scattering).
A260/A280: 1.35 (corrected for light-scattering) (Chu & Helms, 1988)
Virus particles are isometric, 17-19 nm in diameter and presumably of an icosahedral T = 1 symmetry containing 60 subunits (Fig. 6).
Purified virus particle preparations contain up to eight circular ssDNA components (identified previously as C1-C8), each of about 1 kb in size and thought to be encapsidated individually in particles (Fig. 7A and Fig. 8). However, only six of these appear to be integral components of the virus genome (Boevink et al., 1995; Timchenko et al., 2000). By analogy with Faba bean necrotic yellows virus (Timchenko et al., 1999) and Banana bunchy top virus (Horser et al., 2001), the two Rep-encoding DNAs, C2 and C6, are regarded as satellite-like DNAs.
DNA prepared from purified particles also contains linearised forms (Fig. 7B) of the circular DNA components that probably result from degradation during virus purification or DNA extraction. The DNA is about 23% of the particle weight. In addition to the two satellite-like DNAs, subsidiary nucleic acids are likely to include primers required for dsDNA synthesis upon infection (Hafner et al., 1997a).
The virus particle is presumably comprised of 60 chemical subunits of a single protein species (Chu & Helms, 1988) that in SDS-PAGE migrates as a 19- kDa band but has a Mol. Wt of approximately 18.6K as deduced from the major ORF on DNA-S (formerly DNA-C5) (Boevink et al., 1995). This protein is about 77% of the particle weight.
The viral genome consists of at least 6, structurally similar circular ssDNA species, each of c. 1 kb (Boevink et al., 1995; Timchenko et al., 2000). Each of these species contains one major ORF transcribed unidirectionally in the viral sense, and a non-coding region in which there are inverted repeat sequences potentially forming a stem-loop structure and encompassing the origin of replication. Each DNA species contains both a typical TATA box and a polyadenylation signal flanking the ORF, indicating the start and end of transcription, respectively (Boevink et al., 1995). The stem-loop sequence has been shown to be involved in nanovirus DNA replication (Hafner et al., 1997b; Timchenko et al., 1999).
By analogy with other nanoviruses (Timchenko et al., 1999; Vetten and Katul, 2001), the virus genome consists of at least the following ssDNA species and encodes the following proteins (Boevink et al., 1995; Timchenko et al., 2000) (Fig. 8):
- DNA-R (formerly C8, of 1005 nts) encoding the master replication initiator (M-Rep) protein of 33.2 kDa (Accession no. AJ290434);
- DNA-M (C1, of 1001 nts) encoding a putative movement protein of 12.7 kDa (Accession no. U16730);
- DNA-C (C3, of 991 nts) encoding "Clink", a putative cell- cycle link protein of 19.1 kDa (Accession. no. U16732);
- DNA-S (C5, of 998 nts) encoding the coat protein (18.7 kDa) (Accession no. U16734; Chu et al., 1993a);
- DNA-N (C4, of 1002 nts) encoding a putative nuclear shuttle protein of 17.7 kDa (Accession no. U16733);
- DNA-U1 (C7, of 988 nts) encoding a 16.9- kDa protein of unknown functions (Accession no. U16736).
The sequences in the stem and loop are totally conserved between DNA-S, - N, and -U1 and vary only slightly between DNA-M, -C, and -R. They are all clearly distinct from the stem sequences of the satellite-like DNAs C2 and C6.
The degree of sequence conservation in the non-coding regions varies with the different DNA components. The non-coding regions of DNA-C and -S are most similar, sharing 258 conserved nucleotides, DNA-C, -N, -S, and -U1 share 170 identical nucleotides and DNA- C, -M, -N, -S, and -U1 share 152 nt.
The biochemical events involved in SCSV replication have not been determined. Available information for other nanoviruses suggests that replication is similar to that of geminiviruses in being completely dependent on the host cell's DNA replication enzymes, and occurs in the nucleus through double-stranded DNA intermediates by a rolling circle replication mechanism (Laufs et al., 1995; Bisaro, 1996; Hanley-Bowdoin et al., 2000). Upon inoculation, the nuclear localisation signals in the N-terminus of the coat protein ensure that the virus is translocated to the nucleus. Upon decapsidation of viral ssDNA, one of the first biosynthetic events is the synthesis of viral dsDNA (Chu et al., 1993a) with the aid of host DNA polymerase. As the virus DNAs have the ability to self-prime during dsDNA synthesis (Chu & Helms, 1988), it is likely that pre-existing primers are used for dsDNA replicative form (RF) synthesis, as has been shown for mastreviruses (Donson et al., 1984, 1987) and BBTV (Hafner et al., 1997a). From these dsDNAs (RF) forms, host RNA polymerase would then transcribe mRNAs encoding the M-Rep and other proteins required for virus replication. Viral DNA replication is initiated by the M-Rep protein that interacts with common sequence signals on all the genomic DNAs (Timchenko et al., 1999, 2000; Horser et al., 2001). Replication of the DNAs is by cellular enzymes, facilitated and enhanced by the action of Clink, a nanovirus-encoded cell cycle modulator protein (Aronson et al., 2000).
DNA-R encodes the only known replication initiator (Rep) protein essential for the replication of the multipartite genome (Timchenko et al., 2000), and this, together with further evidence for other nanoviruses (Timchenko et al., 1999, 2000; Horser et al., 2001), suggests strongly that the two other Rep-encoding DNAs (C2 and C6) frequently, but not always, associated with SCSV infection, are satellite-like DNAs. By analogy with the satellite rep DNAs of Faba bean necrotic yellows virus and Banana bunchy top virus (BBTV), the SCSV C2- and C6-encoded Rep proteins seem to be capable of initiating replication of their cognate DNA. Unlike the M-Rep protein, however, they cannot catalyse the replication of the other genomic DNA components that encode "non-Rep" proteins. In addition, these satellite-like DNAs are very diverse and do not share any sequence homology in their non-coding regions with each other or with those of the other non-Rep components. In contrast, the M-Rep-encoding DNAs of the nanoviruses are not only phylogenetically distinct from the satellite-like DNAs, but they are also the only rep DNAs that share with the "non-rep" DNAs of a given nanovirus the highly conserved non-coding sequences and in close proximity to, and encompassing the stem loop. (Timchenko et al., 1999, 2000; Horser et al., 2001).
Relations with Cells and Tissues
The virus is detected by ELISA in all plant parts and A. craccivora can transmit the virus from all organs of infected plants, including roots. The concentration of the virus particles is highest in young shoots and roots. No cytopathological effects and no accumulations (inclusions) of virus-like particles have been observed in SCSV-infected plants (Francki et al., 1983), or for any other nanovirus. No conclusive work has been done to determine the in planta or subcellular localisation of the virus and its pathogenesis but the luteovirus-like symptomatology, including phloem necrosis, the need to use cellulases for virus purification and the fact that nanoviruses are not transmitted by mechanical means, suggest that virus replication and virus particles are restricted to phloem tissues. This is supported by the observation that the activity of the promoter sequences present on each genomic DNA species is predominantly confined to vascular tissues (Fig. 9) (Surin et al., 1998).
The virus is able to replicate in isolated pea and subterranean clover protoplasts (Chu et al., 1993b) and de novo synthesis of virus coat protein and DNA was detected by ELISA and nucleic acid hybridization using strand-specific probes, respectively. Nucleic acid hybridization showed that both complementary- and virion-sense DNAs accumulated in protoplasts to a maximum between days 3-7 and at day 7, respectively. The amount of virus coat protein decreased from day 0 to day 3 post-inoculation but increased thereafter over several days to reach a maximum at day 10. The kinetics of virus synthesis in protoplasts is similar to that observed for a geminivirus synthesis in protoplasts (Townsend et al., 1986; Woolston et al., 1989).
Ecology and Control
The virus and its aphid vector are endemic in most regions of southern Australia and are probably maintained all year round in a variety of perennial and annual legume crops and weeds growing over extensive areas. Infection occurs from late autumn to early spring in subterranean clover pastures, which is when pasture productivity is the chief factor limiting stock number and production per head (Grylls & Butler, 1959). High virus incidence in autumn is associated with large numbers of vectors during the summer on a variety of legume plants (Grylls & Butler, 1959; Grylls, 1972). A. craccivora transmits the virus over short distances from crop to crop (Grylls, 1972) and over long distances by migration (Johnson, 1957; Gutierrez et al., 1971). Climatic conditions have a major influence on colonization, establishment and dispersal of vector aphids (Gutierrez et al., 1974). Vector populations are thought to breed on various legume crops in the cooler, wetter coastal and highland areas of southern Australia during late spring and summer and, during autumn and winter, migrate to milder areas as far as northern New South Wales and southern Queensland where subterranean clover, annual Medicago species and other legume crops begin germinating. Annual aphid migration corresponds to annual subterranean clover germination and establishment in various parts of southern Australia. The aphid continues to breed on grain legume hosts, especially broad beans, until early winter (Grylls, 1972). A. craccivora multiplies rapidly under favourable conditions, sometimes reaching 11,000 aphids per plant and serious virus epidemics follow.
ELISA, PCR and nucleic acid hybridization are used for virus diagnosis. No cross-reactions occur with geminiviruses, luteoviruses and other legume- infecting viruses. ELISA detects the virus 4-5 days before the appearance of symptoms, which occurred at about 10 days after inoculation, whereas the nucleic acid hybridisation assay detects the virus 2-3 days before the onset of symptoms. PCR was is the most sensitive assay, detecting the virus 6-7 days before symptoms appeared.
There are no effective natural sources of virus resistance in either French bean (Smith, 1966) or subterranean clover (Chu et al., 1995). Attempts to breed subterranean clover lines with resistance to all SCSD isolates (Grylls & Peak, 1960) have been unsuccessful. Control measures include increasing seeding rates from 3.4-10 kg/ha to compensate for loss from virus infection soon after germination. The large number of different virus isolates, the extensive genetic variation among them, and the potential capacity for genetic recombination between the isolates makes successful classical breeding for virus resistance unlikely. Genetic engineering of resistance using virus-derived genes may provide a better hope of success.
The causal agent of subterranean clover stunt was thought previously to be a luteovirus (Matthews, 1982; Rochow & Duffus, 1981; Ashby & Johnstone, 1985) because of its transmission properties, yellowing symptoms and presumed confinement of the virus to phloem tissues. This confusion may have been reinforced further by the frequent natural association of the virus with members of the family Luteoviridae, such as Soybean dwarf virus (SDV) in subterranean clover.
Field identification of the disease is difficult due to the large variation of symptoms displayed, its similarity to other diseases and disorders, and mixed infections with other viruses. The virus can be confused with infections by luteoviruses such as Bean leaf roll virus and SDV, and in subterranean clover, with those of Subterranean clover mottle virus. Although characteristic symptoms as described above are often seen in subterranean clover pastures, usually the symptoms of the disease vary even within a field. Well spaced plants infected by a severe virus isolate early in the season usually develop a rosetted or stunted and bunchy appearance while those infected by a severe isolate late in spring or in a sheltered position develop a spindly appearance with marked chlorosis and puckering of young leaves. Plants infected by a mild isolate may develop mild yellowing or reddening or appear symptomless.
There are both biological similarities and differences between SCSV, Faba bean necrotic yellows virus and Milk vetch dwarf virus. These three legume-infecting nanoviruses are serologically related, share many common natural and experimental host plants and exhibit similar symptoms on many of these common host species (Chu et al., 1995; Katul et al., 1993; Franz et al., 1997). They also have a common aphid vector, A. craccivora. All the three viruses are highly variable (Chu et al., 1995; Katul et al., 1993, 1999; Franz et al., 1998) in terms of symptoms, other aphid vectors and host plant species as well as serological reactivity among isolates (Franz et al., 1996, 1997, 1998). Differentiation between these viruses is currently based firstly on their different geographical location, then serological analysis (e.g., DAS-ELISA using polyclonal antibodies), and finally confirmed by by sequence analysis of individual genomic DNAs (e.g., DNA-S). Coat protein amino acid sequence differences of > 15% are regarded as a species discriminator.
Effect of virus infection on stunting, reduced foliage and root growth in subterranean clover production.
Moderately severe virus infection on subterranean clover causing, from left to right, reddening of mature leaves, marginal chlorosis on young leaves and general chlorosis and significantly reduced leaf size.
Effects of different virus isolates on the growth of broad bean plants. From left, uninfected control, very severe chlorosis with no growth (isolate A), severe stunting and chlorosis (isolate F), moderate stunting but with significantly reduced leaf size (isolate AA), mild stunting and normal growth form (isolate DD).
Effects of different virus isolates on the growth of pea plants. From left, uninfected control, moderate stunting but with prominent leafrolling and chlorosis and significantly reduced leaf size (isolate F), very severe stunting and chlorosis with no growth (isolate A).
Electron micrograph of virus particles in a purified preparation stained with 2% ammonium molybdate. Bar = 50 nm.
Electron micrographs of viral genomic DNA showing circular (A) and linear (B) molecules. Line tracings of the linear molecules (B) are included to indicate the ends of these molecules.
Diagram illustrating the putative genomic organisation of the virus and depicting the structure of the identified viral DNA components. Each DNA circle contains its designated name (former name in parentheses) and size, and the name/function and size of the encoded protein is given below each circle. Stars refer to the positions of the origin of replication of the DNAs containing the stem-loop structure. Arrows refer to the location and size of the ORFs and the direction of transcription.
References list for DPV: Subterranean clover stunt virus (396)
- Anonymous., CSIRO, Division of Entomology Report, Canberra, 1976. 5 pp.
- Aronson, Meyer, Gyorgyey, Katul, Vetten, Gronenborn & Timchenko, J. Virology 74: 2967, 2000.
- Ashby & Johnstone, Australasian Plant Pathology 14: 2, 1985.
- Bisaro, DNA Replication in Eukaryotic Cells, Cold Spring Harbor Laboratory Press, New York, p. 833, 1996.
- Boevink, Chu & Keese, Virology 207: 354, 1995.
- Chu & Helms, Virology, 167: 38, 1988.
- Chu, Waterhouse & Helms, 7th Australasian Plant Pathology Society Conference, Brisbane, p. 78, 1989.
- Chu, Helms, Waterhouse & Gerlach, 8th Australasian Plant Pathology Society Conference, Sydney, p. 78, 1991.
- Chu, Keese, Qiu, Waterhouse & Gerlach, Virus Research 27: 161, 1993a.
- Chu, Qui, Li & Larkin, Virus Research, 27: 173, 1993b.
- Chu, Boevink, Surin, Larkin, Keese & Waterhouse, Pathogenesis and Host Specificity in Plant Diseases. Vol III, Viruses and Viroids, ed. R.P.Singh, U.S. Singh & K. Kohmoto, Pergamon, Oxford, p. 311, 1995.
- Donson, Morris-Krsinich, Mullineaux, Boulton & Davies, EMBO Journal 3: 3069, 1984.
- Donson, Accotto, Boulton, Mullineaux & Davies, Virology 161: 160, 1987.
- Francki, Randles, Hatta, Davies, Chu & McLean, Plant Pathology 32: 47, 1983.
- Franz, Makkouk, Katul & Vetten, Annals of Applied Biology 128: 255, 1996.
- Franz, Makkouk & Vetten, Phytopathologia Mediterranea 36: 94, 1997.
- Franz, Makkouk & Vetten, Journal of Phytopathology 146: 347, 1998.
- Franz, van der Wilk, Verbeek, Dullemans & van den Heuvel, Virology 262: 210, 1999.
- Grylls, Australian Journal of Experimental Agriculture and Animal Husbandry 12: 668, 1972.
- Grylls & Butler, Journal of Australian Institute of Agricultural Science 22: 73, 1956.
- Grylls & Butler, Australian Journal of Agricultural Research 10: 145, 1959.
- Grylls & Peak, Australian Journal of Agricultural Research 11: 723, 1960.
- Grylls & Peak, Australian Journal of Agricultural Research 20: 37, 1969.
- Gutierrez, Morgan & Havenstein, Journal of Applied Ecology 8: 699, 1971.
- Gutierrez, Nix, Havenstein & Moore, Journal of Applied Ecology 11: 21, 1974.
- Hafner, Harding & Dale, Journal of General Virology 78: 479, 1997a.
- Hafner, Stafford, Wolter, Harding & Dale, Journal of General Virology 78: 1795, 1997b.
- Hanley-Bowdoin, Settlage, Orozco, Nagar & Robertson, Critical Reviews in Biochemistry and Molecular Biology 35: 105, 2000.
- Harvey, Journal of Agriculture Western Australia 7: 634, 1958.
- Horser, Karan, Harding & Dale, Archives of Virology 146: 71, 2001.
- Johnson, Proceedings of the Linnean Society of New South Wales 82: 191, 1957.
- Johnstone & Chu, Viruses of Plants, Descriptions and Lists from the VIDE Database, ed. A.A. Brunt, K. Crabtree, M.J. Dallwitz, A.J.Gibbs & L.Watson, CAB International, Wallingford, p. 1199, 1996.
- Johnstone & McLean, Annals Applied Biology 110: 421, 1987.
- Katul, Vetten , Maiss, Makkouk, Lesemann & Casper, Annals of Applied Biology 123: 629, 1993.
- Katul, Timchenko, Gronenborn & Vetten, Journal of General Virology 79: 3101, 1998.
- Katul, Timchenko, Gronenborn & Vetten, Proceedings of the 15th Meeting of the International Working Group on Legume Viruses, 1999, ed. R. Jones, Fremantle, p.31, 1999.
- Laufs, Jupin, David, Schumacher, Heyraud-Nitschke & Gronenborn, Biochimie 77: 765, 1995.
- Letham, Agfact H8. AB.8, 1st ed., Agfacts, Department of Agriculture, New South Wales, 1982. 2 pp.
- Matthews, Classification and Nomenclature of Viruses, Karger, Basel, 1982.
- O'Loughlin, Journal of Agriculture Victoria 56: 385, 1958.
- Price & Blackstock, Australasian Plant Pathology Society Newsletter 3: 1974.
- Randles et al., Virus Taxonomy. Seventh Report of the International Committee on Taxonomy of Viruses, ed. M.H.V. van Regenmortel et al., Academic Press, San Diego, p.303, 2000.
- Rochow & Duffus, Handbook of Plant Virus Infections: Comparative Diagnosis, ed. E. Kurstak, Elsevier/North Holland Biomedical Press, Amsterdam, p.147, 1981.
- Sano, Wada, Hashimoto, Matsumoto & Kojima, Journal of General Virology 79: 3111, 1998.
- Smith, Australian Journal of Agricultural Research 17: 875, 1966.
- Surin, Boevink, Keese, Chu, Larkin, Llewellyn, Khan, Ellacott & Waterhouse, Agricultural Biotechnology: Laboratory, Field and Market, p.121, ed. P.J. Larkin, Under the Counter Publishing, Canberra, 1998.
- Timchenko, de Kouchkovsky, Katul, David, Vetten & Gronenborn, Journal of Virology 73: 10173, 1999.
- Timchenko, Katul, Sano, de Kouchkovsky, Vetten & Gronenborn, Virology 274: 189, 2000.
- Townsend, Watts & Stanley, Nucleic Acids Research 14: 253, 1986.
- Vetten & Katul, Encyclopedia of Plant Pathology, eds. O.C. Maloy & T.D. Murray, John Wiley & Sons, Inc., New York, p. 670, 2001.
- Woolston, Reynolds, Stacey & Mullineaux, Nucleic Acids Research 17: 6029, 1989.