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The first symptoms to develop when Brassica seedlings are inoculated with TuMV are chlorotic spots on inoculated leaves, mottling followed by systemic vein clearing, mosaic and/or necrosis, leaf distortion and often stunting. In Cheiranthus cheiri, Matthiola incana, Zinnia, Tropaeolum, Petunia and Anemone, flower-breaks occur. Infected plants are often stunted. Some Brassica cultivars develop progressive necrosis of leaves, petioles and stem with some virus isolates, leading to plant death, particularly in B. napus (Walsh and Tomlinson, 1985). A number of biotic and abiotic factors affect symptom expression, particularly temperature and virus isolate (pathotype/'strain').
The symptoms (particularly in interactions between isolates and cultivars and plant species where necrotic symptoms are not induced) can become progressively less pronounced when the plants become infected as they age, to the point of not being easily visible in late infections. The effect of temperature varies considerably for different host and virus isolate interactions. Pound and Walker (1945) noted that mottle symptoms of cabbage plants infected by TuMV were more pronounced and virus titre was higher in plants grown at 28°C than those grown at 16°C. Some resistance genes to TuMV are known to be temperature-sensitive with resistance breaking down at temperatures greater than 30°C (Walsh, 1989). In other studies on other species (Armoracia rusticana and Cichorium intybus), lower temperatures of 15-16°C have been shown to induce more severe symptoms than higher temperatures of 28°C (Pound, 1948; Provvidenti et al., 1979). Symptoms in Brassicaceae can be confused with those caused by Cauliflower mosaic virus (CaMV) (Tomlinson, 1970).
Seed pods of infected Brassicae can be reduced in number and size, some are malformed and seedless, and the size of individual seeds and seed yield are reduced (Walsh and Tomlinson, 1985). Seed viability is affected (Walsh and Tomlinson, 1985). The cytoplasm of infected epidermal, mesophyll and phloem cells contain cylindrical cytoplasmic inclusions consisting of pinwheels, bundles, scrolls and laminated aggregates (Edwardson, 1974; Edwardson and Christie, 1986).
Infection by TuMV may stress plants and allow secondary infection by destructive pathogens. In field situations, mixed infections of TuMV and other viruses are often encountered. In China, mixed infections often occur with Cucumber mosaic virus and a tobamovirus (Chiu and Chang, 1982). In Europe, TuMV often occurs with CaMV (Jenner and Walsh, 1996), Broccoli necrotic yellows virus (Walsh and Tomlinson, 1985) and Beet western yellows virus (Hardwick et al., 1994). The symptoms are usually more severe where plants are simultaneously infected with other plant viruses (Pound and Walker, 1945; Sano and Kojima, 1989).
Control of TuMV is quite difficult due to the very wide host range of the virus, the ineffectiveness of insecticides in controlling the spread of non-persistently transmitted viruses, and the lack of immune crop cultivars.
The use of resistant cultivars would be the simplest, cheapest, most environmentally friendly and possibly the most effective means of control.
Extreme forms of 'race'-specific resistance to the virus have been identified in swede (Tomlinson and Ward, 1978; 1982; Shattuck and Stobbs, 1987; Doucet et al., 1990; Shattuck, 1992) and rape (Walsh and Tomlinson, 1985; Walsh, 1989; Stobbs et al., 1989), forms of B. napus and in B. rapa (Chen, 1980; Provvidenti, 1980; Niu et al., 1983; Green and Deng, 1985; Yoon et al., 1993; Suh et al., 1995). Some of these sources of resistance may be immunity where the virus fails to replicate (Provvidenti, 1980; Tomlinson and Ward, 1982; Shattuck and Stobbs, 1987; Walsh, 1989). However, no such major sources of resistance have been found in B. oleracea types (Walkey, 1982; Pink et al., 1986; Pink and Walkey, 1988; Walkey and Pink, 1988). Where known, the B. napus resistance genes are in the A genome of B. napus, suggesting that there are no genes for extreme forms of resistance to TuMV in the C genome of B. oleracea.
Some more recently identified sources of resistance to TuMV in B. rapa appear to have resistance to a broad spectrum of pathotypes (Suh et al., 1995; J A Walsh, Horticulture Research International, Wellesbourne, UK, 1996). For one of these resistances, no isolate of TuMV so far tested has been found to overcome the resistance. The resistance is not immunity, replication is detected in inoculated leaves, but the viruses do not appear to spread systemically indicating that the resistance is effective against long-distance movement of the virus. One Brassica resistance gene (TuRB01) has been mapped (J A Walsh, Horticulture Research International, Wellesbourne, UK, 1997) and others are currently being mapped (J A Walsh, Horticulture Research International, Wellesbourne, UK, 1996). This will facilitate rapid incorporation of resistance to TuMV into desirable genetic backgrounds via marker-assisted breeding. Inter-species transfer from B. rapa to B. oleracea is also being attempted using these techniques.
Sources of resistance to the virus have also been identified in non-Brassica species. The dominant Tu gene in lettuce confers resistance to infection by TuMV and this gene has been mapped (Robbins et al., 1994). Most accessions of chicory (Cichorium intybus) tested by Provvidenti et al. (1979) were found to be resistant to TuMV and were proposed as a valuable source of resistance for endive and escarole (Cichorium endivia), all accessions of which tested were susceptible to the virus. Resistance to one strain of the virus has been reported in Impatiens balsamina (Provvidenti, 1982). Other sources of resistance have been reviewed by Shattuck (1992).
Non-persistently transmitted viruses are difficult to control with insecticides. Most insecticides do not kill the insects rapidly enough to prevent virus introduction during brief feeding probes. There are examples of their use increasing the incidence of virus in crops (Rice et al., 1983; Roberts et al., 1993) by causing increased probing activity and movement of aphids. Additionally, widespread use of broad-spectrum insecticides is known to reduce populations of beneficial insects including aphid predators and parasitoids and has led to aphids with insecticide resistance. The use of selective insecticides is likely to reduce secondary spread of TuMV in crops following primary infection. There are reports of insecticidal sprays proving ineffective in the control of the virus (Evans and MacNeil, 1983; Niu et al., 1983). As new insecticides are produced and aphid-repellent substances developed, they will have to be evaluated on a case by case basis for their ability to control TuMV.
Oil sprays, although expensive, have been shown to reduce aphid transmission of potyviruses (Bradley et al., 1966; Simons, 1982). Effectiveness seems to vary depending on the type of oil used, the choice of emulsifier, the type of spray nozzle, spray pressure, crop coverage, plant density and spray timing. Some oils are phytotoxic and reduce plant yield. Such treatments have to be repeated frequently to protect newly developing shoots. In greenhouse experiments, Walkey and Dance (1979) showed that a 1% mineral oil spray reduced the transmission of TuMV by Myzus persicae and Brevicoryne brassicae to Indian mustard and was not phytotoxic. However, in field experiments, weekly sprays at the same rate failed to protect cabbage plants from virus infection and appeared to be phytotoxic. In Canada, weekly applications of 1-2% Superior oil to rutabaga (Brassica napus var. napobrassica) production fields reduced, but did not eliminate TuMV infection (Shattuck, 1992). Lowery et al. (1990) found in field experiments that weekly application of cypermethrin with 1% mineral oil TuMV infection of rutabaga from 92 to 37%.
Removal of TuMV-infected plant debris and the eradication of infected plants around fields can help to reduce virus inoculum and hence spread. Scheduling plantings to avoid peak aphid migration periods and allowing susceptible plants to put on as much growth as possible before periods of high risk of virus infection have been practised when no other options have been available. In Canada, the late rutabaga crop is usually planted prior to mid-June and production practices are optimized in order to promote the rapid development of plants before the peak aphid activity in July and August. The effectiveness of this approach is dependent upon keeping the susceptible plants free of TuMV infection during the critical early stages of development when the virus has the most severe effect on plants.
The isolation of crops may be an option in some circumstances. The isolation of nursery beds where the highly susceptible transplants are raised from TuMV infection sources should be a priority, and is practised in some regions to ensure virus-free seedlings. It has been recommended that the complete removal of infected volunteer plants along with dispersal and isolation of field planting of statice (Limonium perezii) will reduce TuMV infection, even in years when the aphid populations are high (Laird and Dickinson, 1972). Aphids can spread viruses over long distances, and minimum isolation distances are usually difficult to determine because they depend on many interacting factors, including the abundance and nature of inoculum sources, aphid population species (vector efficiency, retention time etc.) and density, aphid activity, wind direction and speed, alternative hosts between isolation fields etc. For TuMV, an isolation distance of 3 km may not be sufficient to ensure adequate protection (Laird and Dickinson, 1972).
In the future, it is likely that effective sustainable control measures will depend on the successful integration and management of a number of different control measures. Integrated approaches have been adopted in China for Chinese cabbage (Chin and Chang, 1982) and are reported to be widely accepted by growers. Generally, they consist of postponing sowing by a few days (in northern China), elimination of visiting aphids with systemic aphicides before the 7-leaf stage (plants are less susceptible after this stage) and growing darker green cultivars which are more tolerant to infection than pale green cultivars. The application of some metabolites of Actinomyces is only effective when applied shortly before or after infection. Glazed metallic strips in nurseries or constructing networks of metallic glazed plastic ribbons over transplants being raised in nurseries has also been shown to be an effective component (Chin and Chang, 1982).
Meristem tissue culture techniques combined with chemotherapy and thermotherapy have been used to generate TuMV-free clonal material of cauliflower, horseradish, rhubarb and watercress (Walkey, 1980).
It has been demonstrated recently that transformation of plants with portions of viral genomes frequently gives rise to lines of plants that are resistant to the virus from which the sequence was derived. This phenomenon has been termed 'pathogen-derived resistance'. The nature of resistance obtained is variable and can be either protein- or RNA-mediated (sense and anti-sense). The type and degree of resistance obtained is influenced, at least in part, by the way and position the transgene is inserted into the plant chromosome. Examples of protein-mediated (Stark and Beachy, 1989; Ling et al., 1991; Farinelli and Malnoë, 1993) and RNA-mediated resistance (Lindbo and Dougherty 1992; van den Heuvel et al., 1994) have been demonstrated for a number of potyviruses. Other potential transgenic forms of resistance to potyviruses include the expression of antibodies to potyviruses in plants and the insertion of cloned plant resistance genes. Risks that may be associated with transgenic resistance include escape of transgenes to wild plant species, heterologous encapsidation (Farinelli et al., 1992) and recombination of viral RNA (Falk and Bruening, 1994).
Transgenic resistance to TuMV has been demonstrated in Brassica napus (Lehman et al., 1996). Plants containing, one, two and multiple copies of the coat protein coding region of the viral genome were produced and challenged with high and low doses of two different isolates of TuMV which were different from the isolate (UK 1, pathotype 1) from which the sequence used for transformation was taken (Lehman et al., 1996). Some of the transgenic lines were resistant to one or both of the virus isolates. Interestingly, the B. napus line that was transformed possessed the resistance gene TuRB01, conferring resistance to pathotype 1 isolates of the virus, so the two isolates used to test the resistance belonged to pathotypes 3 and 4. These transgenic lines are undergoing further evaluation.
TuMV does not have quarantine status due to its worldwide distribution. However, movement of plant material infected by the virus, particularly between countries, should be avoided to prevent transfer of new strains to areas where previously they were not present. Restricting or ideally avoiding the introduction of the virus to new areas where it has not been known to occur would also be advisable and beneficial.
Testing of vegetatively propagated crops and plants such as Brassica spp., rhubarb and watercress would be advisable at regular intervals, to ensure stock is virus-free.
In a recent survey of plant virologists in 28 countries and regions, TuMV was ranked as the second most important virus infecting field-grown vegetables (Tomlinson, 1987). It was considered to be the most important virus in northern China and in Taiwan, the second most important in the UK, Greece, Japan and Sweden, the third most important in Eastern China and Ireland and the fourth most important in New South Wales (Australia), Denmark, Poland and Thailand.
Of the brassicaceous crops sampled around Beijing, China, 69% were infected with TuMV (Feng et al., 1990). Considerable production losses in rape and other cultivated Brassicaceae were recorded as long ago as 1940 (Ling and Young, 1940). Similarly, it is widespread in Taiwan in commercial Brassica crops (Yoon et al., 1993). In the UK, TuMV has caused serious losses in swede (Tomlinson and Ward, 1978), white cabbage (Walkey and Webb, 1978; Walkey and Neely, 1980), Brussels sprouts (Tomlinson and Ward, 1981), cauliflower (Tomlinson and Shepherd, 1978) and rape (Hardwick et al., 1994). In Canada, the virus has caused severe production losses in rutabaga (Brassica napus var. napobrassica) (Shattuck and Stobbs, 1987).
Epidemics have occurred periodically in many regions of the world causing serious losses in statice (Limonium perezii) (Niblett et al., 1969) and lettuce in California (Zink and Duffus, 1970), Matthiola incana in Argentina (Pontis, 1973), endive and escarole (Cichorium endiva) in New Jersey and New York (Citir and Varney, 1974b; Provvidenti et al., 1979), collards (B. oleracea) in Georgia and South Carolina (Khan and Demski, 1982), brassicaceous crops in New Zealand (Palmer, 1983), M. incana in the Middle East (Bahar et al., 1985), turnips in Alabama (Wilson and Stevens, 1986) and horseradish in Sweden (Shattuck, 1992).
In addition to reducing yield, TuMV affects quality. In the UK, infected crops of cauliflower have been downgraded and sent for freezing rather than fresh market, representing a considerable economic loss. In Canada, changes in glucosinolate levels have been demonstrated in rutabaga due to TuMV infection and it was suggested that such changes may affect flavour (Stobbs et al., 1991). The virus can also delay crop maturity, e.g. delaying harvest date in cauliflowers by 31 days (Pink and Walkey, 1988). In the UK, complete crop failure has occurred due to TuMV in which whole fields of swede have been ploughed in.