Details of DPV and References
DPV NO: 389 January 2002
Species: Potato mop-top virus | Acronym: PMTV
This is a revised version of DPV 138
Potato mop-top virus
B. D. Harrison Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK
B. Reavy Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK
- 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
Described by Calvert & Harrison (1966) and Harrison & Jones (1970).
A sap-transmissible virus with a tripartite RNA genome and fragile straight tubular particles that occur in low concentration in tissue extracts. The host range is narrow. Found mainly in South America and north-western Europe. Vector is the soil-borne plasmodiophoromycete Spongospora subterranea.
In tuber-bearing potatoes (Solanum tuberosum and other Solanum spp.), causes a wide range of symptoms, which depend greatly on the cultivar (Calvert, 1968) and the environmental conditions. The three commonest shoot symptoms are yellow blotching or mottling (Fig. 1), particularly of the lower leaves, chlorotic V-shaped markings in the leaflets (Fig. 2) and extreme stunting of the shoots, known as `mop-top' (Fig. 3). Shoot symptoms develop best in cool conditions (5-15°C). In the year of infection from soil, the tubers of some cultivars, such as Arran Pilot and Saturna, develop internal brown arcs (`spraing') seen as brown rings on the tuber surface (Fig. 4). The arcs develop at the boundary of the virus-infected tissue in response to a specific change of temperature before or after harvest but do not prevent the virus spreading through the tuber (Harrison & Jones, 1971a). The tubers of other cultivars may develop superficial raised rings without internal symptoms or may show little evidence of infection (Calvert, 1968). In the year following infection from soil, the tubers of some cultivars may be symptomless, or cracked and distorted (Fig. 5), or they may develop internal brown arcs centred on the stolon. When infected tubers are planted, the virus is usually passed to only half, or fewer, of the resulting plants.
Mainly the Andean region of South America (Salazar & Jones, 1975), Scandinavia and the British Isles but also other parts of Western and Central Europe, and other countries where potatoes are grown, such as Japan. In Scotland, the prevalence of infection increases with the annual rainfall (Cooper & Harrison, 1973).
Host Range and Symptomatology
Transmitted by inoculation of sap to 26 species in the Solanaceae or Chenopodiaceae and to Tetragonia expansa; species in 11 other families were not infected (Harrison & Jones, 1970). The virus is also transmissible by grafting (Reavy et al., 1995). Symptoms are greatly affected by the environmental conditions (Harrison & Jones, 1971b).Diagnostic species
Chenopodium amaranticolor. Concentric fine necrotic ringspot lesions develop in inoculated leaves after a week or more at 13-16°C (Fig. 6). A single lesion may spread to cover half a leaf. Not systemic.
Nicotiana debneyi. Inoculated leaves develop necrotic spots, or necrotic or chlorotic ringspots. The first systemically infected leaves show chlorotic or necrotic `thistle-leaf' line patterns (Fig. 7).
N. tabacum (tobacco) cv. Xanthi-nc or Samsun NN. Inoculated leaves develop necrotic or chlorotic ringspots below 20°C, but usually no symptom at higher temperatures. Lesion type can be altered by changing the environmental conditions (Harrison & Jones, 1971b). Systemic infection occurs predominantly in the winter months, causing necrotic or chlorotic `thistle-leaf' line patterns, which appear first in a leaf immediately above an inoculated leaf and later in leaves nearer the shoot tip (Fig. 8).
N. benthamiana and N. debneyi can be used for maintaining cultures. Their systemically infected leaves are the best sources of virus.
N. tabacum cv. Xanthi-nc or Samsun NN is best for isolates that produce necrotic ringspot lesions, and C. amaranticolor for other isolates.
N. debneyi seedlings (systemic symptoms) are useful for testing transmission by vectors (Fig. 7).
The type strain (isolate T; Harrison & Jones, 1970) produces necrotic lesions in many Nicotiana spp. and is more virulent than most other isolates. Several isolates differing in virulence are reported (Harrison & Jones, 1970). The T isolate differs from other recently collected field isolates, such as PMTV-S from Scotland and PMTV-Sw from Sweden, in not being transmissible by the fungal vector (Arif et al., 1995) and in containing a shorter RNA 3 (Reavy et al., 1998; Sandgren et al., 2001). There is little serological variation between isolates.
Transmission by Vectors
The only vector known is the plasmodiophoromycete, Spongospora subterranea f.sp. subterranea (Fig. 9), the causal agent of potato powdery scab disease. The virus is carried, apparently internally, in the resting spores (cystosori), in which it persists for at least 2 years; transmission to roots is by zoospores released by virus-carrying cystosori (Jones & Harrison, 1969). Virus-free S. subterranea zoospores fail to acquire transmissible virus when exposed to a virus suspension but a single-cystosoral isolate of S. subterranea acquired the virus when cultured on infected N. debneyi (Arif et al., 1995). Viruliferous zoospores released from zoosporangia in root cells following this acquisition could apparently transmit the virus to test plants. Field soil is best tested for virus carrying vectors by air-drying the soil at 20°C for 2 weeks, then moistening it and planting N. debneyi bait seedlings (Jones & Harrison, 1969).
The virus seems moderately immunogenic but difficulty may be experienced with most isolates in obtaining enough purified virus particles for production of polyclonal antiserum. This can be useful for immunosorbent electron microscopy (Roberts & Harrison, 1979). Monoclonal antibodies are valuable aids to virus detection, identification and assay in plant shoots and roots, potato tubers and S. subterranea zoospores by triple antibody sandwich ELISA (Torrance et al., 1993; Arif et al., 1994, 1995), and for locating epitopes in the virus coat protein by peptide scanning and in virus particles by immuno-electron microscopy (Pereira et al., 1994). Polyclonal antiserum to the coat protein readthrough protein can be useful for immuno-electron microscopy (Cowan et al., 1997).
A very distant serological relationship to Tobacco mosaic virus (genus Tobamovirus) (Kassanis et al., 1972) and a less distant relationship to Soil-borne wheat mosaic virus (SBWMV; genus Furovirus) (Randles et al., 1976) are reported. PMTV and SBWMV have similar-shaped particles and are transmitted in a similar way by different plasmodiophoromycete vectors (Brakke, 1971).
Sequence comparisons have shown little variation in the coat proteins of PMTV isolates from the Andes, Scandinavia and Scotland (Mayo et al., 1996; Reavy et al., 1997). Sequence analysis of RNA 3 molecules of the T isolate and a recent Scottish field isolate (S) shows that the readthrough protein gene of the T isolate has a 543 nucleotide deletion compared to the S isolate (Reavy et al., 1998). The readthrough protein genes of three Scandinavian isolates are 109 nucleotides longer than that of PMTV-S, due to an insertion of nucleotides at the 3' end of the gene (Sandgren et al., 2001).
PMTV has a similar tripartite genome organisation to that of three other pomoviruses, Beet soil-borne virus (BSBV) (Koenig et al., 1997), Beet virus Q (BVQ) (Koenig et al., 1998) and Broad bean necrosis virus (Lu et al., 1998). However, RNA 3 of PMTV is equivalent to RNA 2 of the other viruses, and their RNA 3 is equivalent to PMTV RNA 2. The deduced amino acid sequences of the coat proteins of PMTV and BSBV are 52% identical but PMTV is not serologically related to BSBV or BVQ (Koenig et al., 1997; R. Koenig and D-E. Lesemann, unpublished data).
Stability in Sap
In tobacco sap, the thermal inactivation point (10 min) is 75 to 80°C and dilution end-point from 10-2 to 10-4. In sap at 20°C, the virus loses most of its infectivity in 1 day but retains a little for 10 weeks.
Torrance et al. (1993). Triturate leaves in 0.025M sodium acetate, pH 5.5, 0.05% (v/v) thioglycerol. Strain through muslin and centrifuge at 10,000g for 20 min. Resuspend pellet in half the original volume with 0.05M Tris HCl, pH 8.7, 0.15% (w/v) urea by stirring for 2 hr at 4°C then clarify by adding an equal volume of chloroform. Centrifuge at 10,000g for 15 min and filter aqueous phase through glass wool before centrifuging at 76,000g for 160 min through a 55% (w/v) sucrose cushion. Pellets are resuspended in 0.05M Tris HCl, pH 8.7, 0.3% (w/v) urea and centrifuged at 8,000g for 5 min. Virus is pelleted from the supernatant fluid twice more through sucrose, resuspending each time in 0.1M Tris HCl, pH 8.7. Virus can be further purified in CsCl gradients.
Properties of Particles
Sedimentation coefficients (s20, w) of components in partially purified preparations were 126, 171 and 239 S. The 239 S component may consist of dimers of 171 S particles (Kassanis et al., 1972).
Particles are straight, helically constructed, with a pitch of 2.4-2.5 nm and a hollow core. Particle width is 18-20 nm and length usually 100-150 or 250-300 nm without well defined peaks in the distribution because the particles easily break. The helix is loosely coiled at one end of many particles in sap (Fig. 10) (Harrison & Jones, 1970; Kassanis et al., 1972). Coat protein subunits probably have a tertiary structure similar to that of Tobacco mosaic virus (Pereira et al., 1994). The coat protein readthrough protein is found at one end of some of the virus particles (Cowan et al., 1997).
Nucleic acid: Three linear single-stranded RNA species of 6043 nt (RNA 1; Savenkov et al., 1999), 2962 nt (RNA 2; Scott et al., 1994), and 2315 nt (RNA 3 of the T isolate; Kashiwazaki et al., 1995). RNA 3 of the of the S isolate is at least 543 nt larger than that of the T isolate (Reavy et al., 1998) and RNA 3 of a Swedish isolate (PMTV-Sw) is 3134 nt (Sandgren et al., 2001).
Protein: The major coat protein has a predicted Mr of 19,720. PMTV particles also contain a small amount of a readthrough protein produced by suppression of the coat protein termination codon (Cowan et al., 1997). The readthrough protein has a Mr of 66,900 in the T isolate (Kashiwazaki et al., 1995), 87,000 in the S isolate (Reavy et al., 1998) and 91,000 in the Sw isolate (Sandgren et al., 2001).
The genome organisation is shown in Fig 11. RNA 1 encodes a 148K replicase protein containing methyltransferase and helicase domains, and a 206K replicase readthrough protein which also contains an RNA-dependent RNA polymerase domain within the readthrough portion. RNA 2 encodes triple gene block proteins, the largest (51K) of which contains a helicase domain, and a cysteine-rich 8K protein of unknown function. RNA 3 encodes the coat protein (CP) and a CP-readthrough protein (RT). All three genomic RNA species appear to have tRNA like structures at the 3'end (Kashiwazaki et al., 1995; Savenkov et al., 1999). The RNA 2 molecule potentially forms a stem-loop structure at the 5' end (Scott et al., 1994).
RNA 1 and RNA 2 can be detected in extracts from inoculated plants that contain a CP transgene. Such extracts produce a symptomless infection that lacks RNA 3 when inoculated onto non-transgenic plants, indicating that the mixture of RNA 1 and RNA 2 is infectious (McGeachy & Barker, 2000).
Nucleotide sequence accession numbers are: PMTV-Sw RNA 1, AJ238607 (Savenkov et al., 1999); PMTV RNA 2, D30753 (Scott et al., 1994); PMTV-Sw RNA 2, AJ277556; PMTV-T RNA 3, D16193 (Kashiwazaki et al., 1995); PMTV-S RNA 3, AJ224991 (Reavy et al., 1998) and PMTV-Sw RNA 3, AJ243719 (Sandgren et al., 2001).
Relations with Cells and Tissues
The virus particles aggregate in sheaves in the cytoplasm of tobacco cells (Fig. 12; White et al., 1972). Tubular structures are reported in cells in potato leaves (Fraser, 1976). The pattern of systemic invasion of test plants suggests that the virus moves from cell to cell in parenchyma tissue and not through sieve tubes. Cell-to-cell movement probably is controlled by the proteins encoded by the triple gene block in RNA 2. The virus is unevenly distributed in potato shoots, roots and tubers.
Ecology and Control
In Scotland, potato seems to be the only important host and the virus survives between potato crops in soil-borne resting spores of S. subterranea (Jones & Harrison, 1972). However, in S. America, a range of crop and weed species in the Solanaceae and Chenopodiaceae (as well as potato species) are likely to be natural hosts of the virus and S. subterranea (Jones, 1988). The virus is spread from field to field in virus-containing resting spores carried by potato tubers or in contaminated soil. Because the virus is not passed to all the progeny tubers of an infected potato plant, it is gradually self-eliminating in potato stocks grown on virus-free soil; elimination is accelerated by removing all symptom-bearing plants (Cooper et al., 1976).
In field plots, S. subterranea infection and PMTV transmission are greatly decreased by treating soil with sulphur to bring the pH below 5.0, or with ZnO at 1400 kg/ha, but these treatments are not economically viable for large-scale application and neither eliminates the virus- carrying vectors (Cooper et al., 1976).
Transgenic N. benthamiana plants expressing the viral coat protein gene are highly resistant or immune to manually-, graft- or vector-inoculated virus (Reavy et al., 1995). This resistance was effective against Scottish and Scandinavian PMTV isolates (Reavy et al., 1997). Similarly, coat protein-transgenic potato plants were highly resistant or immune to vector-inoculated virus in screenhouse tests (Barker et al., 1998).
Symptoms induced by PMTV in potato tubers resemble those caused by Tobacco rattle virus (TRV; genus Tobravirus), although most potato cultivars react somewhat differently to the two viruses. Also, the spraing symptoms caused by TRV in potato tubers involve the production of corky tissue whereas those induced by PMTV do not. The two viruses have different effects on Chenopodium amaranticolor, and PMTV seems not to infect Phaseolus vulgaris. The soil-inhabiting vectors of TRV are nematodes (Trichodoridae), which are killed by air-drying infested soil.
PMTV can be distinguished from other viruses occurring in potato by its particle size and shape, and by the reaction of C. amaranticolor, Nicotiana debneyi, N. tabacum and P. vulgaris. It also can be distinguished from other pomoviruses by the reactions of these four species. RT-PCR is the most sensitive and reliable method for detecting and identifying the virus (Arif et al., 1994) but non-uniform distribution of virus in foliage can lead to sampling problems.
Yellow mottling in leaf of naturally infected potato cv. Red Craig's Royal. (Photograph: Scottish Crop Research Institute)
Chlorotic V-shaped marking in leaf of naturally infected potato cv. Arran Pilot. (Photograph: Scottish Crop Research Institute)
Healthy (left) and naturally mop-top affected (right) plants of potato cv. Alpha. (Photograph: Scottish Crop Research Institute)
External (above) and internal (below) symptoms of spraing in naturally infected tubers of potato cv. Arran Pilot. (Photograph: Scottish Crop Research Institute)
Cracked and distorted naturally infected tubers of potato cv. Alpha produced in second year of infection. (Photograph: Scottish Crop Research Institute)
Lesions consisting of concentric necrotic etched rings in inoculated leaf of Chenopodium amaranticolor. (Photograph: Scottish Crop Research Institute)
Systemically infected Nicotiana debneyi bait seedlings grown in infective soil. (Photograph: Scottish Crop Research Institute)
Systemically infected plant of Nicotiana tabacum cv. Xanthi-nc. (Photograph: Scottish Crop Research Institute)
Spongospora subterranea zoosporangium in root hair of potato cv. Arran Pilot. (Photograph: Scottish Crop Research Institute)
Virus particles 20 nm wide in sap, showing loose helical structure at one end. (Photograph: Scottish Crop Research Institute)
Genome organisation of PMTV. Boxes indicate open reading frames in the three RNA species. The sizes of proteins encoded by RNA 1 and RNA 2 are shown. The size shown for the readthrough (RT) protein encoded by RNA 1 is that of the two ORFs combined and the opal UGA codon is indicated. MT is the methyltransferase domain, Hel are helicase domains, Pol is the polymerase domain. * is the tRNA-like structure at the 3' end of the RNA molecules. The amber UAG codon that terminates the coat protein (CP) ORF in RNA 3 is indicated. Triple-gene block proteins encoded by RNA 2 are shown in green. The readthrough domain of RNA 3 is essentially that of the T isolate with additional sequences found in both the S and Sw isolates shown by horizontal lines. The extra sequence found near the 3' end of RNA 3 of the Sw isolate is shown by vertical lines.
References list for DPV: Potato mop-top virus (389)
- Arif, Torrance & Reavy, Potato Research 37: 373, 1994.
- Arif, Torrance & Reavy, Annals of Applied Biology 126: 493, 1995.
- Barker, Reavy & McGeachy, European Journal of Plant Pathology 104: 737, 1998.
- Brakke, CMI/AAB Descriptions of Plant Viruses 77, 1971.
- Calvert, Record of Agricultural Research of the Ministry of Agriculture of Northern Ireland 17: 31, 1968.
- Calvert & Harrison, Plant Pathology 15: 134, 1966.
- Cooper & Harrison, Plant Pathology 22: 73, 1973.
- Cooper, Jones & Harrison, Annals of Applied Biology 83: 215, 1976.
- Cowan, Torrance & Reavy, Journal of General Virology 78: 1779, 1997.
- Fraser, Protoplasma 90: 15, 1976.
- Harrison & Jones, Annals of Applied Biology 65: 393, 1970.
- Harrison & Jones, Annals of Applied Biology 68: 281, 1971a.
- Harrison & Jones, Annals of Applied Biology 67: 377, 1971b.
- Jones, in Developments in Applied Biology 2. Viruses with Fungal Vectors, p. 255, eds J.I. Cooper & M.J.C. Asher, Wellesbourne, UK: Association of Applied Biologists, 1988.
- Jones & Harrison, Annals of Applied Biology 63: 1, 1969.
- Jones & Harrison, Annals of Applied Biology 71: 47, 1972.
- Kassanis, Woods & White, Journal of General Virology 14: 123, 1972.
- Kashiwazaki, Scott, Reavy & Harrison, Virology 206: 701, 1995.
- Koenig, Commandeur, Loss, Beier, Kaufmann & Lesemann, Journal of General Virology 78: 469, 1997.
- Koenig, Pleij, Beier & Commandeur, Journal of General Virology 79: 2027, 1998.
- Lu, Yamamoto, Tanaka, Hibi & Namba, Archives of Virology 143: 1335, 1998.
- McGeachy & Barker, Molecular Plant-Microbe Interactions 13: 125, 2000.
- Mayo, Torrance, Cowan, Jolly, Macintosh, Orrega, Barrera & Salazar, Archives of Virology 141: 1115, 1996.
- Pereira, Torrance, Roberts & Harrison, Virology 203: 277, 1994.
- Randles, Harrison & Roberts, Annals of Applied Biology 84: 193, 1976.
- Reavy, Arif, Kashiwazaki, Webster & Barker, Molecular Plant-Microbe Interactions 8: 286, 1995.
- Reavy, Sandgren, Barker, Heino & Oxelfelt, European Journal of Plant Pathology 103: 829, 1997.
- Reavy, Arif, Cowan & Torrance, Journal of General Virology 79: 2343, 1998.
- Roberts & Harrison, Annals of Applied Biology 93: 289, 1979.
- Salazar & Jones, American Potato Journal 52: 143, 1975.
- Sandgren, Savenkov & Valkonen, Archives of Virology 146: 467, 2001.
- Savenkov, Sandgren & Valkonen, Journal of General Virology 80: 2779, 1999.
- Scott, Kashiwazaki, Reavy & Harrison, Journal of General Virology 75: 3562, 1994.
- Torrance, Cowan & Pereira, Annals of Applied Biology 122: 311, 1993.
- White, Kassanis & James, Journal of General Virology 15: 175, 1972.