Details of DPV and References

DPV NO: 65 October 1971

Family: Bromoviridae

Genus: Cucumovirus

Species: Peanut stunt virus | Acronym: PSV

This virus is now regarded as a distinct strain of peanut stunt virus

Robinia mosaic virus

K. Schmelzer Institut für Phytopathologie Aschersleben der Deutschen Akademie der Landwirtschaftswissenschaften zu Berlin, Germany

Contents

Introduction

Described by Atanasoff (1935) and Schmelzer (1967a).

Selected synonyms

Black locust true mosaic virus (Rev. appl. Mycol. 46: 1346)

Echtes Robinienmosaik-Virus (Rev. appl. Mycol. 46: 1346)

A virus with isometric particles c. 40 nm in diameter with a wide host range. Transmissible by inoculation of sap, by aphids in the non-persistent manner and also by some Cuscuta species. The virus occurs commonly in south-eastern Europe and rarely in central Europe.

Main Diseases

Causes mosaic and leaf deformation of Robinia pseudo-acacia (Fig. 1).

Geographical Distribution

South-eastern and central Europe (Schmelzer, 1968).

Host Range and Symptomatology

Species of Robinia, especially R. pseudo-acacia, seem to be the only natural hosts, but about 75 plant species in 15 dicotyledonous families have been infected experimentally by sap inoculation (Schmelzer, 1967a), including the following:

Diagnostic species

Chenopodium quinoa. Chlorotic local lesions; systemic mosaic and leaf distortion. Some strains always infect systemically, others only in winter.

C. murale. Chlorotic dots and necrotic pin-point spots; not systemic.

Lycopersicon esculentum (tomato). Systemic mosaic and leaf curling.

Petunia hybrida. Necrotic local lesions (Fig. 5); most isolates not systemic.

Phaseolus vulgaris (French bean). Chlorotic or necrotic local lesions. All tested isolates infect at least some varieties systemically (Fig. 4), giving mosaic, necrosis, and deformation of tip leaves.

Vigna sinensis (cowpea). Chlorotic or necrotic local lesions; systemic mosaic (Fig. 3).

Propagation species

Nicotiana glutinosa (Fig. 6) is suitable both for maintaining cultures and as a source of virus for purification.

N. megalosiphon is also good for maintaining the virus.

Useful amounts of the virus may be obtained from infected N. tabacum except in summer.

Assay species

Chenopodium quinoa is a good local lesion host; C. foetidum has been used in aphid transmission tests.

Strains

Not well studied. Isolates differ in virulence and host range. For example, strain Ro 25, isolated in Hungary (Schmelzer, 1967a), induces chlorotic local lesions as well as mosaic and leaf deformation in Chenopodium quinoa throughout the year, whereas less virulent isolates become systemic in this species only during the winter.

Transmission by Vectors

The aphids Myzus persicae and Aphis craccivora transmit the virus in the non-persistent manner. The transmission rate is rather low. The aphids acquire the virus within 10 min and there is no latent period. Aphis craccivora seems to be the most important vector in nature (Schmelzer & Milicic 1965).

Transmission through Seed

Not found.

Transmission by Dodder

Possible with Cuscuta californica, C. campestris and C. subinclusa. The virus infects the dodder (Schmelzer, 1967a).

Serology

The virus is moderately antigenic. Antisera with titres of 1/128 are readily obtained. The virus reacts well with specific antiserum in agar gel double diffusion tests, which may be done with extracts made by grinding infected Chenopodium quinoa leaves with an equal amount of 0.067 M phosphate buffer (pH 7) or 0.85% NaCl solution. Only one band of precipitate is formed (Schmelzer, 1967a).

Relationships

There seems to be no great serological difference between isolates that differ in their host plant reactions or between isolates obtained from different countries. There is complete cross-protection between different isolates of the virus.

Stability in Sap

Using Chenopodium quinoa as virus source and test plant, the thermal inactivation temperature lies between 66 and 72°C, the dilution end-point above 10^-4, and infectivity is retained at 20-22°C for 5-10 days.

Purification

Homogenize infected tissue with an equal amount of 0.067 M phosphate buffer (pH 7.0) containing 0.3% (w/v) sodium sulphite and 0.3% (w/v) ascorbic acid. Shake the resulting extract thoroughly with twice its volume of ether and, after removing the ether and the coagulated plant material, with half a volume of carbon tetrachloride. Centrifuge at low speed and retain the aqueous phase. Concentrate the virus by one cycle of high and low speed centrifugation. Resuspend in neutral 0.067 M phosphate buffer.

Properties of Particles

Not investigated.

Particle Structure

Particles are isometric, c. 40 nm in diameter (Fig. 2).

Particle Composition

Unknown.

Relations with Cells and Tissues

All tissues seem susceptible with the possible exception of the meristematic regions.

Notes

Cucumber mosaic virus has properties similar to those of robinia mosaic virus, but seems to have smaller particles. Numerous plant species react in almost the same way to both viruses, but, unlike cucumber mosaic virus, robinia mosaic virus usually infects Petunia hybrida only locally and Vigna sinensis, Arachis hypogaea and most cultivars of Phaseolus vulgaris systemically. French bean cultivars that are not infected systemically show large necrotic local lesions, in contrast to the pinpoint lesions induced by cucumber mosaic virus.

Alfalfa mosaic virus also has a host range similar to that of robinia mosaic virus, but common strains infect Phaseolus vulgaris and Vigna sinensis only in inoculated leaves and all strains induce necrotic local or severe necrotic systemic reactions in tomato, which robinia mosaic virus never does. Also the bacilliform particles of alfalfa mosaic virus are unlike those of robinia mosaic virus.

Besides robinia mosaic virus, tomato black ring (both potato bouquet and beet ringspot serotypes) and strawberry latent ringspot viruses cause mosaic symptoms in Robinia (Schmelzer, 1967b). However, these viruses infect Chenopodium murale systemically, whereas robinia mosaic virus is confined to the inoculated leaves, and neither is transmitted by aphids or by the species of dodder that transmit robinia mosaic virus.

Figures

References list for DPV: Robinia mosaic virus (65)

1. Atanasoff, Phytopath. Z. 8: 197, 1935.
2. Schmelzer, Phytopath. Z. 58: 59, 1967a.
3. Schmelzer, A XVII. Növényvédelmi tudományos Értekezlet 1: 139, 1967b.
4. Schmelzer, Arch. Forstw. 17: 621, 1968.
5. Schmelzer & Milicic, Acta bot. croat. 24: 189, 1965.