Cowpea aphid-borne mosaic virus
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Means of Movement and Dispersal
- Seedborne Aspects
- Pathway Causes
- Pathway Vectors
- Plant Trade
- Vectors and Intermediate Hosts
- Impact Summary
- Risk and Impact Factors
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
Don't need the entire report?
Generate a print friendly version containing only the sections you need.Generate report
PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Cowpea aphid-borne mosaic virus
Other Scientific Names
- asparagus bean mosaic virus Snyder
- azuki bean mosaic virus
- cowpea common mosaic virus Bock & Conti
- cowpea Moroccan aphid-borne mosaic potyvirus
- cowpea mosaic virus (Ceylon) Abeygunawardena & Perera
- cowpea mosaic virus (Italy) Vidano & Conti
- cowpea vein-vanding virus Bock
- South African Passiflora virus McKern et al.
Summary of InvasivenessTop of page
Cowpea aphid-borne mosaic virus (CABMV) is a potyvirus capable of infecting many species in the family Fabaceae, and most strains also infect members of the Amaranthaceae, Chenopodiaceae, Cucurbitaceae, Laminaceae, Passifloraceae and Solanaceae. Its main hosts are cowpea and passionfruit, and along with East Asian Passiflora virus and passionfruit woodiness virus it induces passionfruit woodiness in passionfruit. CABMV occurs worldwide but is a particularly major and widespread disease of cowpea in Africa. The nature and severity of the symptoms induced by CABMV are extremely variable, and vary with host cultivars, virus strain and the time of infection. Symptoms can include various mosaics, mottling, interveinal chlorosis and vein-banding. Complete loss of cowpea crops has been reported from Nigeria, and CABMV is one of the main limiting factors to passionfruit yield in South America and Africa.
Taxonomic TreeTop of page
- Domain: Virus
- Unknown: "Positive sense ssRNA viruses"
- Unknown: "RNA viruses"
- Family: Potyviridae
- Genus: Potyvirus
- Species: Cowpea aphid-borne mosaic virus
Notes on Taxonomy and NomenclatureTop of page
Vidano and Conti (1965) were the first to report cowpea aphid-borne mosaic virus (CABMV) infecting cowpeas in Italy, and Lovisolo and Conti (1966) described its properties. Bock and Conti (1974) reviewed the properties of CABMV, and the virus was classified as a distinct member of the potyvirus group (Harrison et al., 1971; Fenner, 1976; Matthews, 1979; Hollings and Brunt, 1981). Since then the virus isolates with properties similar to CABMV have been reported from many countries of the world (Bock and Conti, 1974; Thottappilly and Rossel, 1985; Mali and Thottappilly, 1986; McKern et al., 1994). The original isolate of CAMBV reported from Italy (Lovisolo and Conti, 1966) has been lost (Dijkstra et al., 1987), and the CABMV reported in cowpea from Morocco (CABMV-Mor) (Fisher and Lockhart, 1976) represents the type isolate of the virus and is being used by many workers for comparison with other potyviruses (Taiwo et al., 1982; Dijkstra et al., 1987; Huguenot et al., 1993; Huguenot et al., 1994; McKern et al., 1994; Bashir and Hampton, 1996b). The identification and classification of CABMV was, for a long time, confused with a similar virus reported by Anderson (1955) in cowpea from the USA called blackeye cowpea mosaic (BlCMV). Both viruses have very similar properties and occur worldwide (Bos, 1970; Iwaki et al., 1975; Behncken and Maleevsky, 1977; Lima et al., 1979; Mali and Kulthe, 1980a; Pio-Ribeiro and Kuhn, 1980; Dijkstra et al. 1987). The main reason for this confusion was that neither virus was fully characterized; the CABMV genome has not yet been completely sequenced. The only individual characteristics reported at the time were the existence of serological differences, differential cowpea hosts (Taiwo and Gonsalves, 1982; Huguenot et al., 1993; Bashir and Hampton, 1996b) and different cowpea resistance genes (Provvidenti et al., 1983). Some workers regarded BlCMV and CABMV as closely related or synonymous with each other (Bock and Conti, 1974; Martyn, 1971; Taiwo et al., 1982; Taiwo and Gonsalves, 1982).
Lima et al. (1979) supported the contention that the CABMV isolate described from Morocco and a strain of BlCMV isolated from cowpea in Florida, USA (BlCMV-Flor) were serologically related but differed in other properties. They concluded that BlCMV-Flor and CABMV-Mor were distinct potyviruses. The confusion about the taxonomic status of CABMV was further resolved by Tiawo and co-workers (Taiwo and Gonsalves, 1982; Taiwo et al., 1982), who proved on the basis of biological, physiochemical and serological properties, that some isolates of CABMV such as CABMV-Kenya and CABMV-Nigeria were strains of BlCMV and were very closely related to BlCMV-Flor and BlCMV-New York, while other isolates such as CABMV-Mor and CABMV-Cyprus were different and appeared to be strains of a distinct cowpea aphid-borne mosaic potyvirus. These two groups of viruses were clearly differentiated on the basis of their reactions on selected cowpea genotypes (Taiwo et al., 1982). These results clearly demonstrated that BlCMV represented by BlCMV-Flor isolate and CABMV represented by CABMV-Mor isolate were distinct potyviruses.
However, the confusion on the taxonomic status of the two viruses (BlCMV and CABMV) arose again when Dijkstra et al. (1987) proposed that distinctions between CABMV and BlCMV were unreliable, the term CABMV should be dropped in favour of BlCMV, and that all CABMV isolates should be recognized as strains of BlCMV, as the latter was described earlier (Anderson, 1955) than the former (Lovisolo and Conti, 1966). They concluded that the biological and serological properties of the isolates of these two groups were sufficiently similar to be considered as strains of the same virus. They also argued that original isolates of both CABMV and BlCMV had been lost and were no longer available for direct comparisons, and workers have had to rely on the original description of the viruses to compare their isolates. This issue was further complicated by misidentification of some isolates of both viruses in the past, because the reported CABMV isolates from Kenya, Nigeria and Tanzania (Patel and Kuwite, 1982) were recognized as BlCMV (Taiwo et al., 1982; Taiwo and Gonsalves, 1982).
Previous studies of the comparisons of CABMV and BlCMV were largely based on a few isolates of each virus. Bashir (1992) compared 99 native potyviral isolates derived from cowpea seeds of germplasm accessions and naturally infected field-grown plants, and concluded on the basis of serological and biological properties of these isolates that CABMV and BlCMV were indeed two distinct potyviruses, and not the strains of the same virus. The works of McKern et al. (1993), Huguenot et al. (1993, 1994) and Bashir and Hampton (1996b), further supported the contention that CABMV and BlCMV are two distinct potyviruses.
In another attempt to resolve the taxonomic status of CABMV, McKern et al. (1994) on the basis of high-performance liquid chromatography (HPLC) of tryptic peptides (Shukla et al., 1991) concluded that CABMV-Mor is a potyvirus distinct from bean common mosaic virus (BCMV) (which includes BlCMV, and peanut stripe virus, PStV). Although the distinction between CABMV and BlCMV had already been established (Huguenot et al., 1993; McKern et al., 1994; Bashir and Hampton, 1996b), coat-protein properties such as molecular weight and HPLC peptide profiles are additional criteria by which to discriminate between BlCMV and CABMV (Huguenot et al., 1994; McKern et al., 1994). Partial sequence comparison between CABMV-Mor and BCMNV-Nl3 (McKern et al., 1994) established that the two viruses are distinct. The use of immunoblotting and HPLC in the analysis of capsid proteins of several legume-infecting potyviruses allowed their clear distinction into two groups. The first, with 35 kDa coat protein and 65% peptide profile similarity includes all BCMV strains (BCMV, BCMV-BlC, BCMV-PSt and BCMV-Az). These viruses are so closely related that they are now considered strains of the same virus (Shukla et al., 1991; Mink et al., 1994; Fauquet and Martelli, 1995). The second group, exhibiting a 32 kDa coat protein and consisting of CABMV serotypes, appears to be very heterogeneous in HPLC analysis (37% similarity).
CABMV is also capable of infecting passionfruit, inducing woodiness of the fruit (Nascimento et al., 2006). Woodiness can also be caused by two other potyviruses, Passionfruit woodiness virus (PWV) (Wylie and Jones, 2011) and East Asian Passiflora virus (EAPV) (Iwai et al., 2006). This has led to another taxonomical confusion, with the initial identification of a CABMV isolate from passionfruit being regarded as a new virus, South African Passiflora virus (SAPV) (Brand et al., 1993). SAPV has now been correctly renamed CABMV-SAP (McKern et al., 1994).
The availability of complete genome sequences (Fang et al., 1995; Mlotshwa et al., 2002; Barros et al., 2011) finally settled the taxonomy, establishing CABMV as a distinct species of the genus potyvirus in the family Potyviridae, whereas BlCMV is regarded as a strain of BCMV. CABMV is a closely-related but distinct virus within the BCMV subgroup of potyviruses, which also includes PWV and EAPV (Adams et al., 2012).
DescriptionTop of page CABMV has flexuous filamentous particles 727-765 nm in length and 11 nm wide (Lovisolo and Conti, 1966; Bock, 1973; Kaiser and Mossahebi, 1975; Fischer and Lockhart, 1976; Behncken and Malveesky, 1977) with model length 725-750 nm (Mali et al., 1988; Bashir and Hampton, 1995a). CABMV is one of the potyviruses the particles of which are unaffected by Mg ions (Edwardson and Christie, 1986).
CABMV isolates differ somewhat in stability (in vitro properties) but in cowpea sap the thermal inactivation point (TIP) lies between 57 and 60°C, the dilution end-point (DEP) between 0.001 and 0.0001 and longevity in vitro (LIV) is retained at 20°C for 1-3 days. Frozen infected leaves retain infectivity for at least 7 weeks (Bock, 1973; Lockhart and Conti, 1966). Different workers have reported a variation in properties of CABMV isolates in vitro from different parts of the world. The properties of an isolate of CABMV from Iran were reported as DEP-infection at 0.0001, but none at 0.00001; TIP-infection after heating sap for 10 min at 55°C, none at 60°C; an LIV-infection at 20°C of 7 days, but none at 8 days (Kaiser and Mossahebi, 1975). Ladipo (1976) reported for the Nigerian isolate of CABMV, a DEP of between 0.0001 and 0.00001; a TIP of 50-55°C; and an LIV at 28-30°C of between 8 and 23 h. The CABMV isolate from India exhibited a DEP of 0.001-0.0001; a TIP between 55 and 60°C; and an LIV of 24-36 h (Mali et al., 1988).
Sedimentation coefficients at zero concentration (s°<(sub)20,w>) of 150S have been reported for unaggregated particles of CABMV (Bock and Conti, 1974). Taiwo et al. (1982) reported sedimentation coefficients for CABMV RNA of 40.6 ± 0.7S (CABMV-Mor isolate) and of 40 ± 0.5S (CABMV-Cyprus isolate).
The molecular mass of CABMV coat-protein subunits has been reported as 29,000 kDa, 31,000 kDa and 34,000 kDa (Taiwo et al., 1982). Dijkstra et al. (1987) reported the molecular masses of the protein subunits of five isolates of BlCMV including CABMV-Mor as being within the range 28,000-34,000 kDa.
Electrophoresis and immunoblotting techniques have been used to distinguish capsid protein and to establish relationships with other potyviruses. HPLC was used to compare trypsin digests of the coat proteins in crude extracts of cowpea infected with CABMV isolate. The Mr values of coat proteins, specifically detected by the antibodies were estimated to be 32,000 for CABMV isolates from Nigeria, Cameroon, Mozambique and Morocco (Huguenot et al., 1994). A variation has been reported in the HPLC peptide profiles of tryptic digests of coat proteins of CABMV isolates. Profile similarities among CABMV isolates were estimated and only 37% similarity was observed (Huguenot et al., 1994).
CABMV is one of the potyviruses for which the complete nucleotide sequence has not yet been determined. The 3'-terminal 1221 nucleotide of a Zimbabwe isolate of CABMV has been sequenced. The sequence comprises an open-reading frame (ORF) of 990 nucleotides and a 3' non-coding region of 231 nucleotides followed by a poly-A tail. The ORF has high similarity to coat protein (CP) of potyviruses. A potential CP Q/S cleavage site was identified, yielding a CP of 30.5 kDa containing amino acids (Sithole-Niang et al., 1996).
DistributionTop of page
CABMV occurs in cowpea wherever the crop is grown. The virus occurs worldwide; however, CABMV is a major and widespread disease of cowpea, particularly in African countries. The virus is widespread through sub-Saharan Africa (Bock, 1973; Ladipo, 1976; Thottappilly and Rossel, 1985; Burke et al., 1986), north to the Mediterranean basin (Lovisolo and Conti, 1966; Fisher and Lockhart, 1976; Taiwo et al., 1982), eastward to Turkey, Iran, Iraq and the Indian subcontinent including Afghanistan, Bangladesh and Pakistan (Kaiser and Mossahebi, 1975; Mali and Kulthe, 1980b; Bashir and Hampton, 1993), Indonesia, China and Taiwan (Iwaki, 1979; Tsuchizaki et al., 1984; Thottappilly and Rossel, 1985; Chang et al., 1991) and thence to Australia (Behncken and Maleevsky, 1977), Brazil (Lin and Rios, 1985) and the USA (Kline and Anderson, 1997).
The virus also occurs in Europe (Lovisolo and Conti, 1966; Behncken and Maleevsky, 1977; Lima et. al., 1981; Lin et. al., 1981; Dijkstra et al., 1987). A record of CABMV in the Netherlands (Bos, 1970; CABI/EPPO, 2010) published in previous versions of the Compendium refers to an isolate obtained from Italy and is therefore invalid. CABMV is endemic in many African countries such as Botswana, Egypt, Morocco, Mozambique, Nigeria, Senegal, Tanzania, Togo, Uganda and Zambia (Bock and Conti, 1974; Mali and Thottapilly, 1986; Thottappilly and Rossel, 1992; Ndiaye et al., 1993; Thottappilly et al., 1995; Sithole-Niang et al., 1996).
In South America, the virus has been reported in Brazil in peanuts (Pio-Ribeiro et al., 2000), passionfruit (Nascimento et al., 2004; 2006) and beach bean (Canavalia rosea) (Kitajima et al., 2008). Many reports of ‘passionfruit woodiness virus’ from the 1970s and 1980s probably refer to CABMV, since these two viruses were also misidentified until complete genome sequences became available.
A record of CABMV in Japan (Hino, 1960; Tsuchizaki et al., 1970, 1971; CABI/EPPO, 2010) was included in previous versions of the Compendium on the understanding that CABMV was a synonym of Blackeye cowpea mosaic virus (BlCMV). These viruses are now recognized as separate species and Tsuchizaki et al. (1984) concluded that the virus reported by Tsuchizaki et al. (1970) was BlCMV and not CABMV. CABMV has not been reported in Japan (Ministry of Agriculture, Forestry and Fisheries (MAFF), Japan, personal communication, 2013).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Afghanistan||Present||Bashir and Hampton, 1996a; CABI/EPPO, 2010|
|China||Present||Yu, 1946; CABI/EPPO, 2010|
|Gaza||Present||Taiwo et al., 1982|
|India||Present||Mali and Kulthe, 1980a; Mali et al., 1988; Mali et al., 1989; CABI/EPPO, 2010|
|-Delhi||Absent, intercepted only||CABI/EPPO, 2010|
|-Uttar Pradesh||Present||CABI/EPPO, 2010|
|Indonesia||Present||Iwaki et al., 1975; CABI/EPPO, 2010|
|Iran||Present||Kaiser et al., 1968; Kaiser and Mossahebi, 1975; CABI/EPPO, 2010|
|Iraq||Present||Felga et al., 1981; CABI/EPPO, 2010|
|Japan||Absent, invalid record||Hino, 1960; Tsuchizaki et al., 1970; Tsuchizaki and Hibino, 1971; Tsuchizaki et al., 1984; CABI/EPPO, 2010|
|Malaysia||Present||Brunt et al., 1990; Dahal and Albrechtsen, 1996; CABI/EPPO, 2010|
|Nepal||Present||Dahal and Albrechtsen, 1996; CABI/EPPO, 2010|
|Pakistan||Present||Bashir and Hampton, 1996c; Bashir, 1992; Bashir and Hampton, 1993; CABI/EPPO, 2010|
|Philippines||Present||Beningo and Favali-Hedayat, 1977; CABI/EPPO, 2010|
|Saudi Arabia||Present||Damiri et al., 2013|
|Singapore||Present||Dahal and Albrechtsen, 1996; CABI/EPPO, 2010|
|Sri Lanka||Present||McKern et al., 1994; Dahal and Albrechtsen, 1996; CABI/EPPO, 2010|
|Taiwan||Present||Chang et al., 1991; CABI/EPPO, 2010|
|Thailand||Present||Brunt et al., 1990; Dahal and Albrechtsen, 1996; CABI/EPPO, 2010|
|Turkey||Present||Yilmaz and Ozaslan, 1989; Fidan and Yorganci, 1990; CABI/EPPO, 2010|
|Botswana||Widespread||Bashir and Hampton, 1996b; Burke et al., 1986; Bashir, 1992; CABI/EPPO, 2010|
|Burkina Faso||Widespread||Hampton et al., 1992; CABI/EPPO, 2010|
|Cameroon||Widespread||Thottappilly et al., 1995; CABI/EPPO, 2010|
|Egypt||Widespread||Phatak, 1974; Mazyad et al., 1981; CABI/EPPO, 2010|
|Ghana||Widespread||Thottappilly et al., 1995; CABI/EPPO, 2010|
|Guinea||Widespread||Thottappilly et al., 1995; CABI/EPPO, 2010|
|Kenya||Absent, invalid record||Bock, 1973; CABI/EPPO, 2010|
|Mozambique||Widespread||Hampton et al., 1992; CABI/EPPO, 2010|
|Nigeria||Widespread||Raheja and Leleji, 1974; Ladipo, 1976; Rossel, 1977; Gumedzoe, 1985; CABI/EPPO, 2010|
|Senegal||Widespread||Bashir and Hampton, 1996a; CABI/EPPO, 2010|
|Sierra Leone||Widespread||Huguenot et al., 1996; CABI/EPPO, 2010|
|South Africa||Widespread||Huguenot et al., 1996; CABI/EPPO, 2010|
|Tanzania||Widespread||Patel and Kuwite, 1982; CABI/EPPO, 2010|
|Togo||Widespread||Gumedzoe et al., 1989; Gumedzoe et al., 1990; Thottappilly et al., 1995; CABI/EPPO, 2010|
|Zimbabwe||Present||Thottappilly et al., 1995; CABI/EPPO, 2010|
|USA||Present||Bashir and Hampton, 1996a; Taiwo et al., 1982; Kline and Anderson, 1997; Pappu et al., 1997; CABI/EPPO, 2010|
|-Florida||Present||Taiwo et al., 1982|
|-Georgia||Present||Pappu et al., 1997; CABI/EPPO, 2010|
|-Texas||Present||Kline and Anderson, 1997; CABI/EPPO, 2010|
|Brazil||Widespread||Pio-Ribeiro and Kuhn, 1980; Lima et al., 1981; Nascimento et al., 2004; Nascimento et al., 2006; CABI/EPPO, 2010|
|-Bahia||Widespread||Nascimento et al., 2006|
|-Espirito Santo||Widespread||Nascimento et al., 2006|
|-Minas Gerais||Widespread||Nascimento et al., 2006|
|-Paraiba||Widespread||Nascimento et al., 2004; Nascimento et al., 2006; CABI/EPPO, 2010|
|-Pernambuco||Widespread||Nascimento et al., 2004; Nascimento et al., 2006; CABI/EPPO, 2010|
|-Sao Paulo||Widespread||Nascimento et al., 2006; CABI/EPPO, 2010|
|-Sergipe||Widespread||Nascimento et al., 2004; Nascimento et al., 2006; CABI/EPPO, 2010|
|Germany||Present||Brandes, 1994; CABI/EPPO, 2010|
|Hungary||Present||Bashir and Hampton, 1996a; CABI/EPPO, 2010|
|Italy||Present||Lovisolo and Conti, 1966; CABI/EPPO, 2010|
|Netherlands||Absent, invalid record||Bos, 1970; CABI/EPPO, 2010|
|Australia||Present||Behncken and Maleevsky, 1977; CABI/EPPO, 2010|
|Papua New Guinea||Present||Thottappilly et al., 1995; CABI/EPPO, 2010|
Risk of IntroductionTop of page
The risk criteria may be summarized as follows: economic importance, high; distribution, widespread; seedborne incidence, low in naturally infected seed; seed treatments, none; overall risk, high.
CABMV isolates from passionfruit could be disseminated by cuttings and other types of vegetative propagules. There are some reports on the contamination of cowpea germplasm by CABMV maintained in Georgia, USA (Hampton, 1983; Gillaspie et al., 1995; Bashir and Hampton, 1996a). Similar observations have been reported from developing countries (Mali et al., 1983; Hampton et al., 1992; Bashir and Hampton, 1993; Bashir and Hampton, 1996a). In the case of germplasm-borne viruses there is always a risk that the virus in the germplasm may be transmitted together with the desired genes in populations of breeding progenies, and later spread by vector. It is important to evaluate the cowpea germplasm maintained in each country for seedborne infection and production of virus-free seed to supply to breeders. Strict quarantine measures should be adopted to minimize the risks of virus introduction into new areas, regions and countries. Because CABMV is widely distributed in most African countries, the imported seed from these countries must be passed through quarantine to avoid introduction of CABMV (Bashir, 1995a).
Hosts/Species AffectedTop of page
Cowpea (Vigna unguiculata (L.) Walp.) and passionfruit (Passiflora edulis) are the main hosts of CABMV. However, CABMV has a very large host range (Bock, 1973; Edwardson and Christie, 1986; Chang and Kuo, 1988), including non-cultivated species that could act as natural reservoirs (Kitajima et al., 2010). CABMV infects many species in the family Fabaceae, and most strains also infect members of the Amaranthaceae, Chenopodiaceae, Cucurbitaceae, Laminaceae, Passifloraceae and Solanaceae (Lovisolo and Conti, 1966; Bos, 1970; Bock, 1973).
The host range of the original CABMV isolate included 19 species of the families Amaranthaceae, Chenopodiaceae, Cucurbitaceae, Laminaceae, Leguminosae and Solanaceae (Lovisolo and Conti, 1966). The host range of the CABMV isolate reported from Iran included 15 species in six families (Kaiser and Mossahebi, 1975). The East African strains of CABMV infected 19 species of Leguminosae and 12 species of non-legumes (Bock, 1973). The Moroccan isolate of CABMV (CABMV-Mor) which represents a type isolate of the CABMV, induced systemic symptoms in only five of 27 species inoculated (Fischer and Lockhart, 1976). The Indian isolate of CABMV induced systemic infection in only three legume species (Canavlia ensiformis; Phaseolus lunatus cv. Susses Wonder; and Vigna unguiculata cv. Early Rashorn) (Mali et al., 1988). The CABMV isolate from Tanzania (CABMV-Tanz) induced systemic infection in Vigna aconitifolia, Vigna radiata and Vigna mungo. The following species proved to be symptomless carriers: Glycine max, Cajanus cajan, Cicer arietinum, Lablab purpureus, Vigna subterranea and Lens culinaris. The CABMV isolate from Egypt systemically infected Cucumis sativus cv. Ealady, Lablab purpureus, Phaseolus vulgaris cvs Contender, Giza 3 and Seminol, Trigonella foenum-graecum cv. Giza, Vigna cylindrica, Vigna unguiculata cvs Azmerly, blackeye, Cream No. 7, Petriaat, Pusa Phalguni and Primusa, and Glycine max cvs Clark and Hill (Mazyad et al., 1981).
Edwardson and Christie (1986) have listed that CABMV infected 82 species in 46 genera of 13 families, including 53 species in 28 genera of the Leguminosae. The 13 families are: Amaranthaceae, Aizcaceae, Chenopodiaceae, Cucurbitaceae, Hydrophyllaceae, Labiatae, Iridaceae, Leguminosae, Scrophulariaceae, Polygonaceae, Solanaceae and Pedaliaceae. It has been observed that isolates from different parts of the world varied in their host range. The reason for such variation could be attributed to the existence of the virus in several strains which differ in their ability to infect different host plants (Bock, 1973).
CABMV isolates from Brazil obtained from passionfruit also infect Nicotiana benthamiana, N. clevelandii, common bean and cowpea. The isolates induced chlorotic local lesions in Chenopodium amaranticolor and C. quinoa (Nascimento et al., 2006).
Host Plants and Other Plants AffectedTop of page
|Arachis hypogaea (groundnut)||Fabaceae||Other|
|Glycine max (soyabean)||Fabaceae||Other|
|Pachyrhizus erosus (yam bean)||Fabaceae||Other|
|Passiflora edulis (passionfruit)||Passifloraceae||Main|
|Phaseolus vulgaris (common bean)||Fabaceae||Main|
|Pisum sativum (pea)||Fabaceae||Other|
|Sesamum indicum (sesame)||Pedaliaceae||Main|
|Vigna radiata (mung bean)||Fabaceae||Other|
|Vigna unguiculata (cowpea)||Fabaceae||Main|
|Vigna unguiculata subsp. unguiculata||Fabaceae||Main|
|Voandzeia subterranea (bambara groundnut)||Fabaceae||Main|
Growth StagesTop of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage
SymptomsTop of page
The nature and severity of the symptoms induced by CABMV vary with host cultivars, virus strain and the time of infection (Rossel and Thottappilly, 1985). Natural infection of cowpea causes various mosaics, mottling, interveinal chlorosis and vein-banding (Bock and Conti, 1974). The European (type) strain causes a severe distorting mosaic in cowpea (Lovisolo and Conti, 1966); the African (neotype) strain induces irregular angular broken mosaic; the African mild strain induces a very mild mottle with little or no effect on plant growth; the African vein-banding strain induces a broad dark-green vein-banding (Bock, 1973). The CABMV variant from Australia caused variable mosaic symptoms on cowpea (Mali and Thottappilly, 1986). In the inoculated plants of Vigna cylindrica and Vigna unguiculata, the vein-banding strain of CABMV reported from Nigeria induced vein prominence which was followed on the next trifoliate leaf at least by the characteristic vein-banding symptoms; the subsequent trifoliate leaves either developed vein-banding or mosaic. Systemic mosaic symptoms were induced in Vigna unguiculata cvs New Era, Arlington and Crimson. Varying degrees of susceptibility were exhibited by Vigna unguiculata cv. Ife Brown. Some infected plants were killed; those surviving were severely infected with necrosis of stem and leaves; some developed mild mosaic, while others developed no symptoms and the virus could not then be recovered. In Phaseolus vulgaris cv. LB64003 the virus induced mild to severe systemic mosaic that was accompanied by general stunting. In P. vulgaris cv. H65008, the virus induced local necrotic lesions on inoculated leaves, followed by systemic flecky mosaic, leaf drooping and the eventual death of the plants. Systemic chlorotic mottle was induced in Cajanus cajan.
CABMV symptoms observed on cowpea under field conditions were extremely variable. Factors such as genetic variability of cowpea cultivars grown and stage of growth at the time of infection, influence the type and severity of symptoms produced. Apart from the characteristic vein-banding symptoms that distinguish CABMV from other virus symptoms, conspicuous mild mosaic and mosaic mottle are sometimes observed on plants infected by CABMV under field conditions, depending upon the cowpea cultivars grown (Shoyinka et al., 1997).
Symptoms of the CABMV isolate from Iran on cowpea cv. Early Ramshorn were mosaic, leaf deformation, puckering and stunting (Kaiser and Mossahebi, 1975), whereas cowpea plants infected by the Moroccan isolate (CABMV-Mor) showed mosaic pattern, leaf bumping and distortion, and serious stunting, which resulted in extreme yield reduction (Fisher and Lockhart, 1976). Mazyad et al. (1981) observed severe mosaic on field-grown cowpea plants, whereas yellow mottling with slight distortion of leaflets was observed on plants from Egypt inoculated with CABMV. Bashir and Hampton (1996b) recorded variable symptoms when they tested 51 cowpea genotypes against seven different CABMV isolates. Each isolate induced a wide range of symptoms in plants of susceptible genotypes. The characteristic symptoms of CABMV appeared as vein-banding, interveinal chlorosis, distortion, blistering and stunting of leaves. Some isolates induced necrotic local lesions, sometimes but not always followed by systemic spread of the virus in the form of mosaic pattern. In their host test experiment (Bashir and Hampton, 1996b), CABMV-Mor induced necrotic lesions in the inoculated plants of germplasm accession PI 218123 followed by whole plant necrosis; these plants were therefore said to exhibit lethal susceptibility (LS). In some genotypes symptom expression was delayed. These genotypes remained symptomless for 2-3 weeks, and then developed mild to severe systemic mosaic at a later stage.
Williams (1975) observed widespread mottling, interveinal chlorosis and vein-banding when the field-grown cowpea plants were infected with the CABMV-Nigeria isolate. As the disease developed, leaf cupping occurred; later, leaves became further distorted and developed necrotic lesions. Infected plants remained stunted and bushy, and flowering was retarded or inhibited. Thottappilly and Rossel (1997) reported prominent mosaic, mottle and stunting due to CABMV in bambara groundnut (Vigna subterranea) in Nigeria. Recently Pappu et al. (1997) reported mosaic type symptoms induced in field-grown sesame (Sesamum indicum) plants by a CABMV-SES isolate from the USA.
CABMV isolates obtained from yellow passionfruit plants were able to systemically infect yellow passionfruit (P. edulis f. flavicarpa), Nicotiana. benthamiana, N. clevelandii, bean cv. Preto 153, and cowpea cvs. Pitiúba and Clay (Nascimento et al., 2006). Woodiness symptoms were reproduced in plants of yellow passionfruit (including the fruit), accompanied by severe mosaic and leaf distortion. Both cowpea cultivars displayed mild mosaic symptoms when infected by isolates PB-Alh, PB-Cnd, PE-Bcs1, and PE-Bcs2, and severe mosaic symptoms when infected by other isolates. The bean cultivar Preto 153 reacted in a similar way, except that isolates PE-Ptr and SE-Nps also induced mild symptoms.
Symptoms resulting from seedborne infection
Seedborne infection is expressed in the primary leaves which show vein-clearing, vein-yellowing, diffused chlorotic spots or patches, or an intense chlorosis (Phatak, 1974; Bashir, 1992). Later, in trifoliate leaves, the symptoms are usually more distinct and include vein-yellowing, or variable degrees of yellow mosaic with or without dark-green, or somewhat irregular vein-banding and blistering.
List of Symptoms/SignsTop of page
|Fruit / abnormal shape|
|Fruit / discoloration|
|Fruit / lesions: on pods|
|Growing point / distortion|
|Growing point / lesions|
|Inflorescence / blight; necrosis|
|Leaves / abnormal colours|
|Leaves / abnormal forms|
|Leaves / abnormal patterns|
|Leaves / leaves rolled or folded|
|Leaves / necrotic areas|
|Leaves / yellowed or dead|
|Seeds / discolorations|
|Seeds / lesions on seeds|
|Stems / distortion|
|Stems / stunting or rosetting|
|Whole plant / distortion; rosetting|
|Whole plant / dwarfing|
Biology and EcologyTop of page
The seed transmissibility of CABMV reflects its wide geographical distribution, and probably also virus survival in the off-crop season (Allen, 1983; Rossel and Thottappilly, 1990). The role of weeds and wild legumes as reservoirs of CABMV infection has yet to be determined; however, there is evidence that irrigation and perennially damp areas provide reservoirs of CABMV in the semi-arid savannah of West Africa (Raheja and Leleji, 1974; Rossel, 1977). Infection from infected seeds plays an important role in initiation of the disease, whereas aphids are important in the secondary spread of the disease under field conditions. Cultivation of virus-susceptible cowpea cultivars in a large area is another factor which favours disease spread (Thottappilly, 1992).
CABMV is transmitted readily by mechanical inoculation, by several aphid species and through cowpea seeds.
CABMV is readily transmitted by sap inoculation on cowpea and by aphid vector (Atiri, 1982). The virus has been reported to be transmitted by several aphid species in a stylet-borne non-persistent manner, but Aphis craccivora is reported to be the most efficient vector (Bock, 1973; Atiri et al., 1984, 1986). The aphid species reported to be vectors of CABMV are Aphis craccivora, A. gossypii, A. spiraecola, A. fabae, A. sesbaniae, Macrosiphum euphorbiae, Myzus persicae, Rhopalosiphum maidis and Acyrthosiphon pisum (Bock, 1973; Bock and Conti, 1974; Mazyad et al., 1981; Atiri et al., 1984, 1986; Dijkstra et al., 1987; Mali et al., 1988; Thottapilly, 1992; Thottapilly and Rossel, 1992; Roberts et al., 1993; Bashir and Hampton, 1994). Aphis craccivora is a widespread and common pest of cowpea in many countries of Africa and in India (Singh and Allen, 1979). Transmission of CABMV by A. craccivora has been reported as 57% (Bashir and Hampton, 1994) and 64-71% with A. craccivora and A. gossypii. Both aphid species were reported as efficient vectors of CABMV (Roberts et al., 1993). When single aphids were used, the transmission was erratic, but batches of 10-15 individuals invariably transmitted the virus (Phatak, 1974). Mazyad et al. (1981) found that M. persicae was a more efficient vector for transmitting CABMV isolate than A. craccivora.
Both the colonizing and transient aphid species are important in the epidemiology of CABMV, but colonial species of Aphis are principally responsible for secondary spread. Other species, such as R. maidis, play an important role in the development of infection foci (Atiri et al., 1986). The feeding behaviour of A. craccivora, the most important species on the cowpea crop in Africa, is influenced by the cowpea cultivar, with populations and sizes of aphids being less on aphid-resistant cowpea than on aphid-tolerant or susceptible cultivars (Atiri et al., 1984).
A significant positive correlation between alatae forms of A. craccivora and CABMV incidence in aphid-susceptible and aphid-tolerant cowpea lines was observed, whereas this correlation was negative in the aphid-resistant lines. However, the incidence of CABMV was not significantly different in any of the lines, indicating that aphid activity (e.g., wide dispersal) was more important in the spread of CABMV than the absolute number of viruliferous alatae (Atiri et al., 1984).
Cowpea resistance to aphids has been reported (Chari et al., 1976; Singh and Allen, 1979, 1980).
Three CABMV isolates have been completely sequenced: CABMV-Z from Zimbabwe, obtained from cowpea (GenBank Access # AF348210); CABMV-BR1 from Brazil, obtained from peanut (HQ880242), and CABMV-MG-Avr, also from Brazil, obtained from passionfruit (HQ880243). The genome of CABMV-Z is 9465 nucleotides (nt) long and contains one long open reading frame (ORF), starting at nt 76 and finishing at nt 9237, encoding a polyprotein with 3053 deduced amino acids (aa) (Mlotshwa et al., 2002). The two Brazilian isolates have significantly longer genomes (9894 and 9930 nt for BR1 and MG-Avr, respectively), and the polyproteins encoded have 3174 (BR1) and 3182 (MG-Avr) aa. This difference is due to a 129 aa deletion in the P1 coding region of isolate CABMV-Z (Barros et al., 2011). The small ORF PIPO and the conserved G(1-2)A(6-7)
motif associated with its putative frameshift translation are also present in the three isolates (Barros et al., 2011). A total of nine cleavage sites are present in the polyproteins of the three isolates. Seven out of those nine are conserved among the two Brazilian isolates, and three (HC-Pro/P3, 6K2/NIa-VPg and NIb/CP) are conserved among the three isolates (BR1, MG-Avr and Z). The differences observed at the cleavage sites for 6K1/CI and CI/6K2 involve amino acid residues that probably do not affect recognition by the viral protease NIa-Pro. However, the sequences at the cleavage sites for P3/6K1 and NIa-Pro/Nib differ at several positions, including the highly conserved valine residue at position -4, which is present in the sequence of the Brazilian isolates but not of isolate Z. The sequences of the Brazilian isolates are identical at both sites (EHVETQ//A for P3/6K1, and DGVATQ//S for NIa-Pro/NIb) and are in full accordance with the consensus sequences for NIa-Pro cleavage sites, XXVXXQ/E//S/G/A/V (Martín et al., 1990; Carrington et al., 1993; Barros et al., 2011).
Means of Movement and DispersalTop of page
CABMV is transmitted readily by mechanical inoculation, by several aphid species and via cowpea seeds. The presence of CABMV in cowpea seeds allows for long-distance dispersal via the international seed exchange (Hampton et al., 1992)
Aphids play an important role in the secondary spread of the disease. The aphid species reported to be vectors of CABMV are Aphis craccivora, A. gossypii, A. spiraecola, A. fabae, A. sesbaniae, Macrosiphum euphorbiae, Myzus persicae, Rhopalosiphum maidis and Acyrthosiphon pisum (Bock, 1973; Bock and Conti, 1974; Mazyad et al., 1981; Atiri et al., 1984, 1986; Dijkstra et al., 1987; Mali et al., 1988; Thottapilly, 1992; Thottapilly and Rossel, 1992; Roberts et al., 1993; Bashir and Hampton, 1994). A. craccivora is reported to be the most efficient vector.
Seedborne AspectsTop of page
Transmission of CABMV through cowpea seeds tends to ensure viral perennation and endemicity, together with long-distance dissemination by international seed exchange (Hampton et al., 1992). Seed transmission depends mainly on gamete infection by the virus (Tsuchizaki et al., 1970) and plays an important role in the epidemiology of CABMV.
In comparing three periods of infection, it was found that the earliest inoculation (7 days after sowing) led to highest incidence of infection. No correlation was found between infection incidence and severity of symptoms, or between infection incidence and seed transmission rate. However, the severity of symptoms and rate of seedborne infection were positively correlated (Aboul-Ata et al., 1982).
Location of Virus in Seed
Phatak (1974) reported that CABMV was detected in the plumule of the growing infected seeds, although in some cases the cotyledons had virus; virus was not detected in the seed coat. Tsuchizaki et al. (1970) detected virus from pollens, anthers and ovaries of CABMV-infected cowpea plants. Virus concentration was higher in anthers, but lower in ovaries in CABMV-infected plants. They also studied the effect of time of infection in seed transmission, concluding that seed transmission occurred only when the mother plants were inoculated with CABMV at least 20 days before flowering whereas CABMV was recovered from pollen only when the mother plants were inoculated at least 17 days before flowering.
In cowpea variety Zairai-Tsurunashi-Kintoki infected with CABMV, both virus particles and cytoplasmic inclusions were seen in the meristemic tissues 0.1-0.2 mm from the apex. No virus was detected in the cell layer above this. Electron microscopic examination of individual pollen for virus particles revealed that 6.8% and 18.6% of pollen grains from cowpea plants infected with strains CABMV-1 and CABMV-3, respectively, contained virus particles. Developing immature embryos are not liable to infection through direct invasion by the virus from mother plants in a non-seed transmission virus-host combination, but are infected through direct invasion by virus from the mother plant in a seed-transmitted virus-host combination (Tsuchizaki and Hibino, 1971).
The incidence of CABMV transmission through seed varies with strains/isolates and cowpea genotype interaction, and has been reported to range from 0 to 40% (Bock, 1973; Phatak, 1974; Ladipo, 1977, Aboul-Ata et al., 1982; Hampton et al., 1992). Bashir and Hampton (1994) studied the rate of seed transmission of 12 isolates of CABMV under greenhouse conditions by mechanical inoculation in three cowpea genotypes. Variable seed-transmission rates were obtained depending upon the interaction of virus isolate and cowpea genotype. Maximum transmission (55%) of CABMV was recorded with isolate RN-27C in genotype 58-57. Three isolates (RN-27C, RN-28C and PI-44C) were not transmitted in any cowpea genotype. The transmission in naturally infected seeds has been reported in the range 1.1-13.3% in cowpea (Kaiser and Mosshebi, 1975; Mazyad et al., 1981; Mali et al., 1983, 1988, 1989; Hampton. et al., 1992; Bashir and Hampton, 1993; Bashir and Hampton, 1996a).
Effect on Seed Quality
CABMV causes deformation of pods, reduction in seed size, discoloration of seed and reduction in seed germination, but there is no correlation between yield reduction and percentage seed transmission (Kaiser and Mossahebi, 1975).
Symptoms Expressed from Seedborne Infection
Seedborne infection is expressed in the primary leaves, which show vein-clearing, vein-yellowing, diffused chlorotic spots or patches, or an intense chlorosis (Phatak, 1974; Bashir, 1992). Later, in trifoliate leaves, the symptoms are usually more distinct and include vein-yellowing, or variable degrees of yellow mosaic with or without dark-green, or somewhat irregular vein-banding and blistering.
The virus survives in infected seed, volunteer host plants and in viruliferous aphids. The infected seeds provide primary inoculum foci in the field and the aphid species play an important role in the secondary spread of the disease (Thottappilly, 1992).
No seed treatment has yet been reported to eliminate CABMV directly from seed.
Seed Health Tests
Correct identification of seedborne viruses is a fundamental step in any method adopted for control of a virus disease or seed certification programme. Significant progress has been made in developing and applying methods for the rapid detection and identification of virus in seed. Most of these methods are based on serology, while various blot tests (e.g., Southern and Northern) permit the detection of specific virus nucleic acid by the use of isotope-labelled complementary DNA (cDNA) (Bashir, 1995a).
Various methods commonly used for detecting seed-transmitted viruses (Maury and Khetarpal, 1989; Bashir, 1995a; Bashir and Hassan, 1998) are discussed below.
This test is based on the appearance of virus symptoms in the seedlings growing from the virus-infected seeds. In a controlled environment room test, seedlings are grown in a controlled chamber or greenhouse and the presence of virus is determined after 2-3 weeks when the seedlings express virus symptoms, or by ELISA.
In this method the presence of virus is assayed by inoculating extracts of seed or seedlings on to indicator hosts such as Chenopodium amaranticolor, C. quinoa, Phaseolus vulgaris, Vigna unguiculata, Datura stramonium. This method can also reveal the symptomless or latent infection of plants as in a grow-out test. However, it has the following limitations: (a) it needs a span of time to standardize the indicator hosts and to record the symptoms; (b) a large space in the form of a controlled-environment chamber or greenhouse is required; and (c) the test is laborious and time-consuming.
These tests are based on the specific reaction between antibodies and antigens. The antiserum is used to perform serological tests, which are the most reliable and effective methods for the detection of seedborne viruses and virus from plant tissues.
The important serological methods which are used for detecting viruses in seed are the agar gel double diffusion test, microprecipitin test, dot-immunobinding assay (DIBA), immunoelectron microscopy (IEM) and enzyme-linked immunosorbent assay (ELISA) (Taiwo et al., 1982; Dijkstra et al., 1987; Bashir, 1995b; Bashir and Hassan, 1998).
Agar gel double diffusion test
In this test the antigens and antibodies are added to wells punched in agar or agarose. They diffuse through the agarose and form a thin precipitin line where they meet in optimum proportions. This test is less sensitive, requires more antiserum and is difficult to perform with elongated viruses.
This test involves placing small drops of antiserum on plastic or plastic-coated Petri dishes or in small wells in plastic trays. Plant or seed extract is added to the antiserum and the drops are viewed under the microscope to observe precipitation. Different antiserum antigen ratios can be tested. The drops in the Petri dishes are covered with liquid paraffin to prevent evaporation.
Dot-immmunobinding assay (DIBA)
The basic principle is similar to that of plate ELISA. In this assay antigens are first immobilized on nitrocellulose membranes. Because these membranes have a high affinity for proteins, it is essential to block the free protein-binding sites of the membrane. This 'blocking' is usually done with non-fat dry milk, or bovine serum albumin or gelatin. Immobilized antigen is then exposed to solutions of unconjugated virus-specific antibody. Crude antiserum is usually used for this purpose. Trapped antibody is probed with alkaline phosphatase (ALP), or horseradish peroxidase (HRP)-labelled protein A, or anti IgG. Finally, the conjugated treated membrane is exposed to substrate solution, i.e., naphthol phosphate for ALP. Naphthol phosphate in the presence of ALP is converted to phosphoric acid and naphthol. Naphthol is detected by adding a diazonium salt; together these form a coloured, insoluble product which can be detected visually.
Immunoelectron microscopy (IEM)
In this method virus and antiserum are reacted together, and the results are viewed in the electron microscope. Carbon-coated copper grids are first coated with virus-specific antiserum, and then with virus-containing extract of seed or seedlings. Viruses reacting with the antiserum remain attached to the surface of the grid and are then detected by negative staining upon viewing in the electron microscope. This method can give 1000-fold or more increase in sensitivity over conventional electron microscopy in detecting viruses (Roberts and Harrison, 1979). This is the most sensitive of all the serological tests, is rapid, and a very small amount of antiserum is required. The only limitation is the high cost of an electron microscope.
Enzyme-linked immunosorbent assay (ELISA)
ELISA is one of the most widely used serological tests for the detection of seedborne infection. In this test, the immunospecific activity is recognized through the action of associated enzyme label rather than by observing the formation of an insoluble antigen-antibody complex as in other serological tests. The basic principle of ELISA is to immobilize the antigen on the solid surface and probe with a specific immunoglobulin carrying an enzyme label. There are two main categories of ELISA procedures, direct ELISA (DAS-ELISA) and indirect ELISA (DAC-ELISA). DAS-ELISA is highly strain specific, whereas DAC-ELISA is more sensitive but less specific than direct ELISA. Indirect ELISA is mostly used to detect viruses from seeds.
In the DAC-ELISA procedure the plates are first coated with antigen. The immobilized antigen is then targeted by unconjugated virus-specific antibody in crude antiserum. The trapped antibody is detected by an enzyme-labelled secondary antibody, e.g., goat anti-rabbit IgG. The main advantage of DAC-ELISA is that one enzyme conjugate can be used for all systems. This form of ELISA is particularly suitable for virus detection during disease surveys, for testing for the presence of virus in seeds and for establishing serological relationships among viruses. It is also more economical than DAS-ELISA (Bashir and Hampton, 1995b).
Pathway CausesTop of page
|Seed trade||CABMV is seedborne in cowpea seeds||Yes||Yes|
Pathway VectorsTop of page
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|True seeds (inc. grain)||Yes|
Vectors and Intermediate HostsTop of page
|Aphis craccivora||Bock, 1973.||Insect|
|Aphis fabae||Bock and Conti, 1974.||Insect|
|Aphis gossypii||Bock and Conti, 1974.||Insect|
|Aphis medicaginis||Bock and Conti, 1974.||Insect|
|Macrosiphum euphorbiae||Bock and Conti, 1974.||Insect|
|Myzus persicae||Bock and Conti, 1974.||Insect|
Impact SummaryTop of page
ImpactTop of page
Raheja and Leleji (1974) reported the complete loss of a cowpea crop in northern Nigeria resulting from CABMV attack under irrigated field conditions. Kaiser et al. (1968) reported 44-80% seed yield reduction in cowpea in greenhouse studies. A yield loss of 13-87% due to natural infection of cowpea by CABMV was reported in Iran (Kaiser and Mossahebi, 1975), and 48-60% loss in cowpea was reported in Zambia (Kannaiyan and Haciwa, 1993).
Passionfruit woodiness caused by CABMV is a devastating disease, and one of the main limiting factors to passionfruit yield in South America and Africa. The complete absence of control measures forces growers in Brazil, the world's largest producer of passionfruit, to grow passionfruit as an annual crop, replanting it every year, despite the plant’s potential to remain productive for many years.
Risk and Impact FactorsTop of page Invasiveness
- Proved invasive outside its native range
- Negatively impacts agriculture
- Negatively impacts livelihoods
- Pest and disease transmission
- Highly likely to be transported internationally accidentally
DiagnosisTop of page
Proper detection and identification of a virus is a prerequisite in developing strategies for its control. Several reliable and quick diagnostic methods have been developed by many workers to detect and identify CABMV in host-plant tissue, vector and seed (Huguenot et al., 1993; McKern et al., 1994; Huguenot et al., 1996; Sithol-Niang et al., 1996; Bashir and Hampton, 1996b). Some indicator host plants are used for diagnosis in which the virus induces characteristic necrotic or chlorotic local lesions on primary inoculated leaves. The diagnostic species are listed below under Diagnostic Hosts.
Virus identification is mainly based on the study of physical, biological and chemical properties. This includes the study of mechanical or vector transmission, a host-range test, infectivity assay, in vitro properties, serological tests, molecular analysis of capsid protein and nucleic acid analysis. The virus in seed is usually detected by a growing-out test, infectivity assay, or by enzyme-linked immunosorbent assay (ELISA) (Phatak, 1974; Bashir and Hampton, 1993; Gillaspie et. al., 1995). See also Seedborne Aspects.
Seedborne isolates of CABMV were detected and identified from cowpea germplasm accessions by direct antigen coating enzyme-linked immunosorbent assay (DAC-ELISA) (Hampton et al., 1992; Bashir and Hampton, 1996a). Konate and Neya (1996) used a biotin/streptavidin ELISA technique to detect CABMV in cowpea seeds. Joshi and Albrechtsen (1992) used a mixed antisera technique in ELISA to detect CABMV in virus-infected plant tissue. A mixed panel of monoclonal antibodies has been used to detect virus in plant tissue (Mink and Silbernagel, 1992; Huguenot et al., 1993). Other tests which have been employed to detect and identify CABMV isolates are SDS-immunodiffusion, immunosorbent electron microscopy (IEM), slide-agglutination, dot-immunobinding assay (DIBA), tissue blot test (TBT), capsid protein immunoblotting, HPLC peptide profile analysis and nucleic acid sequence analysis (Taiwo et al., 1982; Dijkstra et al., 1987; Huguenot et al., 1994; McKern et al., 1994; Bashir, 1995a; Bashir and Hampton, 1996b; Huguenot et al., 1996; Bashir and Hassan, 1998). Of all these tests, ELISA is the most commonly used to analyse seed or plants samples collected during surveys (Bashir and Hassan, 1998).
Virus propagation species
CABMV virus cultures are best maintained in cowpea (Vigna unguiculata), common bean (Phaseolus vulgaris), soyabean (Glycine max) and/or tobacco (Nicotiana benthamiana). However, for virus purification, cowpea cultivars Pusa Phalguni, California blackeye, Vigna unguiculata subsp. sesquipedalis or Phaseolus vulgaris cv. Red Kidney have been used (Bock and Conti, 1974; Dijkstra et al., 1987; Huguenot et al., 1994; Bashir and Hampton, 1995a; Huguenot et al., 1996; Pappu et al., 1997).
The following diagnostic species have been reported for CABMV: Chenopodium amaranticolor; Glycine max (soyabean); Ocimum basilicum (basil); Petunia hybrida; Phaseolus vulgaris cvs Prince, Canadian Wonder, Saxa, Scotia, Black Turtle Soup and Bountiful; Vigna radiata cv. Oklahoma 12; Nicotiana glutinosa; N. tabacum; Chenopodium album; Cucumis sativus; Gomphrena globosa; Datura stramonium; Pisum sativum cvs Greenfeast, Koroza and Laxaton; Chenopodium quinoa (Lovisolo and Conti, 1966; Kaiser and Mossahebi, 1975; Ladipo, 1976; Mazyad et al., 1981; Mali et al., 1988; Nascimento et al., 2006).
Inclusion bodies induced by CABMV
Comparison of the cytoplasmic inclusion bodies induced by plant viruses in their host cells is another criterion by which to distinguish between viruses or virus strains (Edwardson et al., 1972; Bashir, 1995b). Some viruses form distinct cytoplasmic structures in their host cells and these can be used for rapid identification of the virus by light or electron microscopy. The diversity of potyvirus-induced cytological alteration is so great that several can serve as markers in diagnostic work. In certain cases strains of potyviruses may be differentiated by the presence of inclusion bodies, although they are very similar (Bashir, 1995b). CABMV induces numerous granular inclusions in Petunia (Bos, 1970). Light microscopy (Bos, 1970; Lima et al., 1981) and electron microscopy of CABMV infected cells (Tsuchizaki and Hibino, 1971; Behncken and Maleevsky, 1977; Lima and Purcifull, 1979) revealed cytoplasmic cylindrical inclusions consisting of pinwheels and bundles with associated scrolls (subdivision-1). CABMV cylindrical inclusions have the same morphology as those induced by BlCMV (type-1) (Edwardson and Christie, 1986). The Moroccan isolate of CABMV and BlCMV induce similar types of cylindrical inclusion bodies in their hosts, but the former can be distinguished from BlCMV by the DSD-immunodiffusion test (Lima et al., 1978; Lima et al., 1979; Taiwo and Gonsalves, 1982).
Potyviruses are difficult to purify because of their tendencies towards irreversible aggregation during extraction and concentration, with consequent virus loss during low-speed centrifugation (Bashir and Hampton, 1995a). CABMV can be purified from systemically infected leaves of cowpea (Vigna unguiculata), V. unguiculata subsp. sesquipedalis or Nicotiana benthamiana (Dijkstra et al., 1987; Bashir and Hampton, 1995a). Several protocols have been described for the purification of CABMV (Bock and Conti, 1974; Lima et al., 1979; Taiwo et al., 1982; Dijkstra et al., 1987; Zhao et al., 1991). Virus yields per 100 g of leaf tissue of virus-infected cowpea averaged about 1-2 mg (Dijkstra et al., 1987). Bashir and Hampton (1995a) compared three different methods of virus purification of two CABMV isolates (RN-7C and PI-23C) using two species, V. unguiculata cv. Pusa Phalguni and N. benthamiana. The highest average yield of 4.6 mg per 50 g leaf tissue of tobacco was obtained with the RN-7C isolate. Carbon tetrachloride (CCl<(sub)3>) when used in combination with chloroform as a clarifying agent improved the yield of BlCMV, but not that of CABMV isolates.
Detection and InspectionTop of page
CABMV symptoms observed on cowpea under field conditions are extremely variable. Factors such as genetic variability of cowpea cultivars grown and stage of growth at the time of infection seemed to influence the type and severity of symptoms produced. Apart from the characteristic vein-banding symptoms that distinguish CABMV from other virus symptoms, conspicuous mild mosaic and mosaic mottle are sometimes observed on plants infected by CABMV under field conditions, depending upon the cowpea cultivars grown (Shoyinka et al., 1997).
For further details see Symptoms.
Similarities to Other Species/ConditionsTop of page
Virus isolates having similarity to CABMV in symptomatology, in vitro properties, aphid and seed transmissibility, but not fully characterized, include the cowpea mosaic viruses of McLean (1941), Yu (1946), van Velsen (1962), Abeygunawardena and Perea (1964) and Shankar et al. (1973), and the asparagus bean mosaic virus of Synder (1942).
CABMV and a BlCMV strain (BCMV-BLC) are closely related, but apparently distinct, potyviruses. Both these viruses occur worldwide on cowpea, induce similar symptoms, and are so similar for in vitro and transmission properties that the taxonomic status of CABMV remained a controversial issue for a long time (Taiwo et al., 1982; Dijkstra et al., 1987; Bashir and Hampton, 1996b). Isolates identified as CABMV from Kenya and Nigeria are now thought to be more correctly assigned to BlCMV, to which isolates from the USA are also referred (Taiwo and Gonsalves, 1982; Taiwo et al., 1982). It has now been established that CABMV and BCMV-BLC are two distinct potyviruses (Lima et al., 1979; Taiwo et al., 1982; Shukla et. al., 1991; Hugenot et al., 1993; Bashir and Hampton, 1996b).
Among the African strains, the neotype and the mild strains are serologically identical; the vein-banding strain is distinguishable, although related. European strains were considered to be similar based on a distinct serological relationship with BCMV (Lovisolo and Conti, 1966; Bock, 1973). Within the potyvirus group, CABMV was considered to be distinctly related to BCMV and apparently shares no antigens with bean yellow mosaic virus (BYMV), pea seedborne mosaic virus (PSbMV), clover yellow vein virus (ClYVV), soybean mosaic virus (SMV), potato virus Y (PVY), tobacco severe etch virus (TSEV), sugarcane mosaic virus (ScMV), or iris mosaic virus (Bock, 1973). Some CABMV isolates did not show any serological relationship with BCMV (Kaiser and Mossahebi, 1975; Fisher and Lockhart, 1976).
Lima et al. (1979), Behncken and Maleevsky (1977) and Lima et al. (1981) found a serological relationship of CABMV with BCMV and BlCMV, but no relationship with BYMV. Taiwo and Gonsalves (1982) also reported no serological relationship of CABMV with the potyviruses BYMV and SbMV; however, a serological relationship of partially purified preparations of the SMV-CS isolate was found with antiserum to SMV-CS obtained from the USA (Nain et al., 1994).
On the basis of HPLC and immunoblot analysis of crude sap from virus-infected plants using specific monoclonal antibodies, CABMV and bean common mosaic necrotic virus (BCMNV-NL3) were found to be serologically related, suggesting a possible similarity between them (Huguenot et al., 1996). HPLC profiles of tryptic peptides, and amino acid partial sequences of CABMV-Mor were compared with those of the coat proteins of BlCMV-type, BlCMB-W, mild mottle strain of peanut stripe virus (PStV-MM), and NY-15 and NL3 strains of BCMV, all of which are strains of BCMV. The HPLC peptide profiles indicated that CABMV-Mor was distinct from BCMV and BCMNV. Amino-acid sequence analysis of peptides confirmed that CABMV-Mor was not a strain of BCMNV or BCMV (McKern et al., 1994). Partial sequence data of coat protein suggested that CABMV-Mor was very similar to South African Passiflora virus (SAPV) (McKern et al., 1994), and in fact SAPV is now classified as a strains of CABMV, CABMV-SAP (Adams et al., 2012).
Immunoblot analysis of coat protein (CP) showed a serological relationship between CABMV and the necrotic strain of BCMV (BCMNV-NL3), and provided evidence of a possible similarity between CABMV and BCMV-NL3. Trypsin digest peptide profiles of the coat proteins of various CABMV serotypes and BCMV-NL3 were distinct (Huguenot et al., 1994). Phylogenetic analysis of the 3' terminal region of a potyvirus isolated from naturally infected sesame (Sesamum indicum) in Georgia, USA, indicated that the virus (CABMV-GA) is closely related to CABMV and SAPV. Based on sequence identity, the virus was recognized as a strain of CABMV (Pappu et al., 1997).
Using indirect ELISA, Mink and Silbernagel (1992) compared eight isolates of BCMV, five of BlCMV and four of CABMV against a panel of 13 monoclonal antibodies (MAbs) raised against BCMV, BlCMV CABMV and PStV. Four MAbs detected all isolates, suggesting their coat proteins have at least one epitope in common. Dijkstra and Khan (1992) compared CABMV-Mor, three strains of BCMV (NL1, NL3 and NY15) and four strains of BlCMV (Fla, Ind, NR and W); based on host range, antigenic properties and N-terminal peptide profiles it was concluded that Fla, NR and W are strains of BCMV, whereas NL3, Ind and CABMV-Mor (and possibly NL1 and NY15) are separate viruses (Dijkstra and Khan, 1992; Mink et al., 1994).
Strains of CABMV
A range of CABMV strains differing widely in symptoms and host range has been identified (Bock, 1973); for example, the European (type) strain (Lovisolo and Conti, 1966), African (neotype) strain (Bock, 1973), African mild strain (Bock, 1973) and African vein-banding strain (Bock, 1973; Ladipo, 1976). Taiwo et al. (1982) reported three distinct pathogenic strains of CABMV. A distinct strain of CABMV from Senegal was reported by Ndiaye et al. (1993). The other CABMV strains are South African Passiflora strain (CABMV-SAP) from South Africa (McKern et al., 1994), Zimbabwe strain (CABMV-Z) from Zimbabwe (Sithole-Niang et al., 1996), Brazilian strain from Brazil (Sousa et al., 1996; Nascimento et al., 1994; 1996) and Moroccan strain from Morocco (Fisher and Lockhart, 1976). Taiwo and Gonsalves (1982) have shown that the viruses described by Bock (1973) and Ladipo (1976) as CABMV strains are serologically identical to BlCMV; therefore they should no longer be regarded as strains of CABMV. The potyvirus designated PTY+ by Ndiaye et al. (1993) was later determined to be a distinct virulent strain of CABMV; this strain clearly differed from CABMV-Mor (Hampton et al., 1997).
Serotypes of CABMV
Seven distinct CABMV serotypes have been reported on the basis of biotin-labelled monoclonal antibodies and double-antibody sandwich enzyme-linked immunosorbent assay (DAS-ELISA) (Huguenot et al., 1993; Ndiaye et al., 1993; Huguenot et al., 1996).
Pathotypes of CABMV
Considerable evidence of pathogenic variability among CABMV isolates has been reported (Bashir and Hampton, 1992; Gumedzoe, 1995; Bashir and Hampton, 1996b,c).
Prevention and ControlTop of page
Because CABMV spreads under field conditions through increased aphid populations, availability of susceptible host plants and the presence of initial infection foci through seedborne infection, appropriate control can be achieved by limiting these three factors. CABMV is transmitted by several species of aphid in a non-persistent manner; therefore the use of insecticides may not be an appropriate approach. However, chemical control of aphids at proper times may be useful in keeping the vector population at a low level to avoid secondary spread of the disease. Until genetic engineering is further refined, breeding for host-plant resistance remains the most practical approach for the control of CABMV (Rossel and Thottappilly, 1985).
Excellent sources of resistance to CABMV have been identified among cowpea germplasms in the USA (Taiwo et al., 1982), Brazil (Lima et al., 1981), Nigeria (Williams, 1977; Ladipo and Allen, 1979a), Tanzania (Patel et al. 1982), Iran (Kaiser and Mossahebi, 1975), India (Mali et al., 1981), Zambia (Kannaiyan et al., 1987) and Senegal (Ndiaye et al., 1993).
Cisse et al. (1997) reported an extra early maturing cowpea line PI 596353 that was not only resistant to CABMV but also to the aphid vector (A. craccivora) and bacterial blight disease. Bashir and Hampton (1996c) tested 51 cowpea lines by mechanical inoculation under greenhouse conditions against seven CABMV geographical diverse isolates, and identified TVU-410, TVU-1582 and TVU-1593 as immune to all seven isolates. Several immune, resistant and tolerant genotypes were identified against individual isolates. Mligo (1989) developed a cowpea cultivar VU 1-1 through breeding resistance to CABMV and bacterial blight. Ladipo and Allen (1979b) identified some cowpea lines resistant to CABMV and A. craccivora, and to two other cowpea viruses (southern bean mosaic virus and cowpea mosaic virus). Mih et al. (1991) reported TVU-15656 to be a highly resistant line to both CABMV and cucumber mosaic virus. Sources of CABMV resistance, including those with combined resistance against several distinct viruses (Allen, 1980, 1983) have now been widely utilized in cowpea breeding, both in Nigeria (Singh et al. 1987) and elsewhere in Africa (Kannaiyan and Haciwa, 1993).
Three types of resistance to CABMV are recognized: immunity, hypersensitive resistance and viral tolerance; immunity is more common than others (Bashir and Hampton, 1996c). Resistance is also expressed as development of very mild mosaic without adverse effects on plant growth (latent infection) (Patel. et al., 1982) and also by tolerance in which systemic infection occurs without the appearance of symptoms (Ladipo and Allen, 1979a; Bashir and Hampton, 1996c). Several cowpea lines with isolate-specific resistance have been identified (Bashir and Hampton, 1996c).
No cowpea genotypes have been found to possess resistance to both CABMV and BlCMV; however, cowpea genotypes including TVUs 22, 410, 1582, 1593, 612, 1453, 1948, 2331, 2480, 2657, 2740, 3433, Big Boy, Corona and Serodo have been identified as useful differentials (Ladipo and Allen, 1979a; Patel et al., 1982; Taiwo et al., 1982; Bashir and Hampton, 1996b, c).
To date, no sources of resistance to CABMV have been identified in Passiflora.
Genetics of Resistance
Inheritance of resistance in cowpea to CABMV is governed by a single dominant or recessive gene (Taiwo et al., 1981; Fisher and Kyle, 1994, 1996), sometimes in association with minor and/or modifier genes (Patel et al., 1982). Provvidenti et al. (1983) reported that resistance to CABMV and BlCMV in common bean (Phaseolus vulgaris) was conferred independently by a single dominant factor that appears to be closely linked. Systemic resistance to CABMV in P. vulgaris cv. Great Northern 1140 (GN1140) is conditioned by a dominant allele which has been designated Cam2. Under some environmental conditions, a recessive allele at an unlinked locus Cam3 also controls a resistant response to CABMV (Fisher and Kyle, 1996). Monogenic dominant resistance to CABMV in P. vulgaris cv. Black Turtle Soup was reported by Fisher and Kyle (1994).
The virus can be controlled through cultural practices which include early sowing and intercropping of cowpea with cereals, possibly leading to decreased virus incidence (Kannaiyan and Haciwa, 1993). The use of virus-free seed is potentially important, particularly in preventing spread to new areas (Zettler and Evans, 1972).
Production of Virus-Free Seed
The production of virus-free seed is a potential measure for the control of CABMV, particularly if certified seed can be produced in areas where the virus is not known to occur (Zettler and Evans, 1972). Field inspection and roguing of diseased plants may help to eliminate seedborne inoculum, but because there is evidence that CABMV may also occur and occasionally be seed-transmitted in symptomless plants (Aboul-Ata et al., 1982; Bashir and Hampton, 1996c), there is potential value in implementing a rapid indexing procedure for the detection of CABMV in seedlots. It is well established that the level of seed transmission varies with cultivar (Bock and Conti, 1974; Kaiser and Mossahebi, 1975; Ladipo, 1977; Aboul-Ata et al., 1982). Cowpea genotypes resistant to seed transmission have been identified (Ladipo, 1977; Mali et al., 1981, 1983; Bashir and Hampton, 1994). Because the detection of virus in seed depends partly on the sensitivity of the assay (Konate and Neya, 1996), the incidence of seed transmission in some cultivars might be sufficiently low to control the disease significantly. As the minimum level of seedborne inoculum at which an epidemic may be initiated is still unknown, selection for resistance to seed transmission could prove a useful strategy for CABMV control (Aboul-Ata et al., 1982).
Resistance to Seed Transmission
The identification of cowpea cultivars/lines that prevent seed transmission of the virus or permit only minimum seed transmission would help to decrease the transmission rate. Cowpea lines resistant to seed transmission have been identified (Ladipo, 1977; Mali et al., 1983).
Control Through Seed Certification
Certification against seedborne viruses such as CABMV is one of the methods which minimizes their spread and it must be used in the production of certified seed. The seed certification programme should be started at the basic level of the germplasm collection available to the plant breeders, and continue through the subsequent development of varieties. Moreover, such programmes must also take into consideration other means by which a particular seedborne virus may be disseminated in the standing crop. The major method of monitoring the presence of seedborne viruses, i.e., visual inspection, should be followed in the standing crop. A seed certification programme for CABMV is being practised at IITA, Ibadan, Nigeria. Samples of 1000 seeds are planted and the resulting seedlings are examined visually for the presence of seed-transmitted viruses. Differential hosts and serological tests are used to facilitate identification where appropriate. Seed lots showing 2% seed transmission or greater are not distributed; those with less than 2% seed transmission are so indicated and the recipient is advised. ELISA is the most reliable method for virus detection from seed and plant tissue (Hamilton, 1983).
Resistance to Aphid Vector
Cowpea cultivars possessing resistance or tolerance to the CABMV aphid vector (Aphis craccivora) have been identified (Chari et al., 1976; Singh and Allen, 1979, 1980), but it is not known whether their use would limit the rate of CABMV spread.
Chemical Control of Virus Vector
Although control of disease through chemical treatment of aphids in the case of potyviruses is not effective, certain insecticides may possibly have potential in controlling CABMV. Organophosphates and carbamate had no effect. Atiri et al. (1987) found synthetic pyrethroid cypermethrin restricts the acquisition and inoculation of the virus, and protects against its transmission; however, the initial virus introduction was not prevented by these synthetic pyrethroids when the incidence of incoming alatae aphids was high, and virus incidence was higher in sprayed plots relative to unsprayed controls (Roberts et al., 1993).
ReferencesTop of page
Abeygunawardena; DVW; Perera SMD, 1964. Virus diseases affecting cowpea in Ceylon. Tropical Agriculturist, Ceylon Agriculture Society, 120:181-204.
Aboul-Ata AE; Allen DJ; Thottappilly G; Rossel HW, 1982. Variation in rate of seed transmission of cowpea aphid-borne mosaic virus in cowpea. Tropical Grain Legume Bulletin, 25:2-7.
Adams MJ; Zerbini FM; French R; Rabenstein F; Stenger DC; Valkonen JPT, 2012. Family Potyviridae. Virus Taxonomy. 9th Report of the International Committee on Taxonomy of Viruses [ed. by King, A. M. Q. \Adams, M. J. \Carstens, E. B. \Lefkowitz, E. J. (.]. London, UK: Elsevier Academic Press, 1069-1089.
Allen DJ, 1983. Disease resistance in crop improvement. In: The Pathology of Tropical Food Legumes. Chichester, UK: John Wiley and Sons, 210-213.
Anderson CW, 1955. Vigna and Crotalaria viruses in Florida. II. Notations concerning cowpea mosaic virus (Marmor vignae). Plant Disease Reporter, 39:349-353.
Anonymous, 1981. Viruses, virus-like diseases and mycoplasma-like diseases. In: Pest Control in Tropical Grain Legumes. London: Centre for Overseas Pest Research (COPR), 95-110.
Atiri GI, 1982. Virus-vector-host relationship of cowpea aphid-borne mosaic virus in cowpea [Vigna unguiculata (L.) Walp.]. PhD thesis, University of Ibadan, Nigeria.
Atiri GI; Ekpo EJA; Thottappilly G, 1984. The effect of aphid-resistance in cowpea on infestation and development of Aphis craccivora and the transmission of cowpea aphid-borne mosaic virus. Annals of Applied Biology, 104(2):339-346
Atiri GI; Thottappilly G; Ligan D, 1987. Effects of cypermethrin and deltamethrin on the feeding behaviour of Aphis craccivora and transmission of cowpea aphid-borne mosaic virus. Annals of Applied Biology, 110(3):455-461
Barros DR; Alfenas-Zerbini P; Beserra JEA; Antunes TFS; Zerbini FM, 2011. Comparative analysis of the genomes of two isolates of cowpea aphid-borne mosaic virus (CABMV) obtained from different hosts. Archives of Virology, 156(6):1085-1091. http://www.springerlink.com/content/v4468l800667g717/
Bashir M, 1992. Serological and biological characterization of seed-borne isolates of blackeye cowpea mosaic and cowpea aphid-borne mosaic potyviruses in Vigna unguiculata (L.) Walp. PhD Thesis, Oregon State University, Corvallis, Oregon, USA.
Bashir M, 1995. Inclusion bodies of plant viruses - a review. Journal of Agricultural Research (Pakistan), 33:223-234.
Bashir M, 1995. Seed-borne viruses of legume crops: symptomatology, epidemiology, detection and control. 31-47. In: Ahmed SI, ed., Legume Seed Health Testing. Proceedings of the Training Course on Legume Seed Health Testing. 4-20 March 1995. Organized in Collaboration with ICARDA, Aleppo, Syria and Federal Seed Certification Department. Ministry of Food, Agriculture and Livestock. Government of Pakistan, Islamabad, Pakistan.
Bashir M; Hampton RO, 1992. Biological characterization of pathotypes of blackeye cowpea mosaic and cowpea aphid-borne mosaic potyviruses. Abstracts of APS/MSA Joint Annual Meeting held in Portland, Oregon, USA, 8-12 August 1992, Abstract No. A-392.
Bashir M; Hampton RO, 1995. Antiserum production against cowpea aphid-borne mosaic virus and standardization for enzyme-linked immunosorbent assays. Sarhad Journal of Agriculture, 11(4):505-512; 16 ref.
Bashir M; Hampton RO, 1996. Detection and identification of seed-borne viruses from cowpea (Vigna unguiculata (L.) Walp.) germplasm. Plant Pathology, 45:54-58.
Bashir M; Hampton RO, 1996. Serological and biological comparisons of blackeye cowpea mosaic and cowpea aphid-borne mosaic potyvirus isolates seed-borne in Vigna unguiculata (L.) Walp. germplasm. Journal of Phytopathology, 144(5):257-263; 39 ref.
Bashir M; Hampton RO, 1996c. Sources of genetic resistance in cowpea (Vigna unguiculata (L.) Walp. to cowpea aphid-borne mosaic potyviruses. European Journal of Plant Pathology, 102:411-419.
Bashir M; Hassan S, 1998. Diagnostic Methods for Plant Viruses. Islamabad, Pakistan: Pakistan Agricultural Research Council.
Bock KR; Conti M, 1974. Cowpea aphid-borne mosaic virus, No. 134. In: CMI/AAB Descriptions of Plant Viruses. Wellesbourne, UK: Association of Applied Biology, 4 pp.
Bos L, 1970. The identification of three new viruses isolated from Wisteria and Pisum in the Netherlands, and the problem of variation within the potato virus Y group. Netherlands Journal of Plant Pathology, 76:8-46.
Brand RJ; Burger JT; Rybicki EP, 1993. Cloning, sequencing, and expression in Escherichia coli of the coat protein gene of a new potyvirus infecting South African Passiflora. Archives of Virology, 128(1-2):29-41.
Brandes J, 1994. Eine eletronenmikroskopisch Schnellmethode Zum Nachweis faden-und stabchenformiger Viren, ins besondere in Kartoffelddumnkelkeimen. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes Braunschweig, 9:151-152.
Carrington JC; Haldeman R; Dolja VV; Restrepo-Hartwig MA, 1993. Internal cleavage and trans-proteolytic activities of the VPg-proteinase (NIa) of tobacco etch potyvirus in vivo. Journal of Virology, 67(12):6995-7000.
Chang CA, 1991. Virus diseases and their transmission of legume crops in Taiwan. Integrated control of plant virus diseases. Proceedings of the International Workshop TARI, Taichung, Taiwan, April 9-14, 1990 [edited by Kiritani, K.; Su, H.J.; Chu, Y.I.] Taipei, Taiwan; Food and Fertilizer Technology Center for the Asian and Pacific Region, 99-110
Chang CA; Kuo YJ, 1988. Cowpea aphid-borne mosaic virus and its effect on the yield and quality of asparagus bean. Journal of Agricultural Research of China (Taiwan), 32:270-278.
Damiri BV; Al-Shahwan IM; Al-Saleh MA; Abdalla OA; Amer MA, 2013. Identification and characterization of Cowpea aphid-borne mosaic virus isolates in Saudi Arabia. Journal of Plant Pathology, 95(1):79-85. http://sipav.org/main/jpp/index.php/jpp/article/view/2695
Dijkstra J; Bos L; Bouwmeester HJ; Hadiastono T; Lohuis H, 1987. Identification of blackeye cowpea mosaic virus from germplasm of yard-long bean and from soybean, and the relationships between blackeye cowpea mosaic virus and cowpea aphid-borne mosaic virus. Netherlands Journal of Plant Pathology, 93(3):115-133
Edwardson JR; Christie RG, 1986. Viruses infecting forage legumes. Vol. II. Monograph No. 14. Agriculture Experimental Station, University of Florida, Gainesville, Florida, USA.
Edwardson JR; Zettler FW; Christie RG; Evans IR, 1972. A cytological comparison of inclusions as a basis for distinguishing two filamentous legume viruses. Journal of General Virology, 15:113-118.
Fang GW; Allison RF; Zambolim EM; Maxwell DP; Gilbertson RL, 1995. The complete nucleotide sequence and genome organization of bean common mosaic virus (NL-3 strain). Virus Research, 39:13-23.
Fauquet C; Dejardin J; Thouvenel JC, 1986. Evidence that the amino acid composition of the particle proteins of plant viruses is characteristic of the virus group. II. Discriminant analysis according to structural biological and classification properties of plant viruses. Intervirology, 25(4):190-200
Fidan U; Yorganci U, 1990. Investigation on the detection and seed transmission of the virus diseases occurring on pulse crops in pgean Region. 2. Seed transmission of virus diseases by grower seeds and seeds of artificially infected pulse crops. Journal of Turkish Phytopathology, 19(1):1-5
Fisher ML; Kyle MM, 1994. Inheritance of resistance to potyviruses in Phaseolus vulgaris L. III. Cosegregation of phenotypically similar dominant responses to nine potyviruses. Theoretical and Applied Genetics, 89(7/8):818-823
Fisher ML; Kyle MM, 1996. Inheritance of resistance to potyviruses in Phaseolus vulgaris L. IV. Inheritance, linkage relations, and environmental effects on systemic resistance to four potyviruses. Theoretical and Applied Genetics, 92(2):204-212; 41 ref.
Gumedzoe MY, 1985. Studies of the variability of the cowpea aphid-borne mosaic (CAbMV) complex in Nigeria. PhD Thesis, University of Laval, Quebec, Canada.
Gumedzoe MY; Sundu DY; Thottappilly G, 1989. Inventarie des principaux viroses du niebe Papier presente a la Conference biannale de I'Association Scientifique Ouest Africaine, Cotonou, 10-15 September 1989.
Hamilton RI, 1983. Certification scheme against seed-borne viruses in leguminous hosts; present status and further areas for research and development. Seed Science and Technology, 11:1051-1062.
Hampton RO, 1983. Seed-borne viruses in crop germplasm resources: disease dissemination risks and germplasm-reclamation technology. Seed Science and Technology, 11:535-546.
Hampton RO; Thottappilly G; Rossel HW, 1997. Viral diseases of cowpea and their control by resistance-conferring genes. In: Sing BB, Mohan Raj DR, Dashiell KE, and Jacka LEN, eds. Advances in Cowpea Research. Ibadan, Nigeria: International Institute of Tropical Agriculture (IITA) and Japan International Research Centre for Agricultural Sciences (JIRCAS), 159-175.
Harrison BD; Finch JT; Gibbs AJ; Hollings M; Shepherd RJ; Valenta V; Wetter C, 1971. Sixteen groups of plant viruses. Virology, 45:356-363.
Hino T, 1960. Studies on asparagus bean mosaic virus. Annals of the Phytopathological Society of Japan, 25:178-186.
Hollings M; Brunt AA, 1981. Potyviruses. In: Kurstak E, ed. Handbook of Plant Virus Infections and Comparative Diagnosis. Amsterdam, North Holland: Elsevier, 731-807.
Huguenot C; Furneaux MT; Clare JA; Hamilton RI, 1996. Improved diagnosis of cowpea aphid-borne mosaic virus in Africa: significance for cowpea seed-indexing, breeding programs and potyvirus taxonomy. Archives of Virology, 141(1):137-145; 18 ref.
Huguenot C; Furneaux MT; Hamilton RI, 1994. Capsid protein properties of cowpea aphid-borne mosaic virus and blackeye cowpea mosaic virus confirm the existence of two major subgroups of aphid-transmitted, legume-infecting potyviruses. Journal of General Virology, 75(12):3555-3560
Huguenot C; Furneaux MT; Thottappilly G; Rossel HW; Hamilton RI, 1993. Evidence that cowpea aphid-borne mosaic and blackeye cowpea mosaic viruses are two different potyviruses. Journal of General Virology, 74(3):335-340
Iwai H; Yamashita Y; Nishi N; Nakamura M, 2006. The Potyvirus associated with the dappled fruit of Passiflora edulis in Kagoshima prefecture, Japan is the third strain of the proposed new species East Asian Passiflora virus (EAPV) phylogenetically distinguished from strains of Passion fruit woodiness virus. Archives of Virology, 151(4):811-818. http://springerlink.metapress.com/content/y10129467867132x/?p=9a7412d353c94cd7b4cfd577138f4c92&pi=14
Iwaki M; Roechan M; Tantera DM, 1975. Virus diseases of legume plants in Indonesia. 1. Cowpea aphid-borne mosaic virus. Contributions from the Central Research Institute for Agriculture, Bogor, Indonesia, 13:14 pp.
Joshi S; Albrechtsen SE, 1992. Use of mixed individual antisera in ELISA-detection of viruses infecting cowpea and soybean. Journal of the Institute of Agriculture and Animal Science, 13:83-87.
Kaiser WJ; Danesh D; Okhovat M; Mossahebi H, 1968. Diseases of pulse crops (edible legumes) in Iran. Plant Disease Reporter, 52:687-689.
Kannaiyan J. Haciwa HC, 1993. Diseases of food legume crops for the scope of their management in Zambia. FAO Plant Protection Bulletin, 41:73-90.
Khan JA; Lohuis D; Goldbach R; Dijkstra J, 1993. Sequence data to settle the taxonomic position of bean common mosaic virus and blackeye cowpea mosaic virus isolates. Journal of General Virology, 74(10):2243-2249
Kitajima EW; Alcântara BKde; Madureira PM; Alfenas-Zerbini P; Rezende JAM; Zerbini FM, 2008. A mosaic of beach bean (Canavalia rosea) caused by an isolate of Cowpea aphid-borne mosaic virus (CABMV) in Brazil. Archives of Virology, 153(4):743-747. http://springerlink.metapress.com/content/c690469832402204/?p=c8ecb57db134444dadd757190d83f52c&pi=15
Ladipo JL, 1976. A vein-banding strain of cowpea aphid-borne mosaic virus in Nigeria. Nigerian Journal of Science, 10:77-86.
Lima JAA; Purcifull DE; Edwardson JR, 1978. Serological, biological and cytological distinctions between three legume viruses. International Virology, 4:586.
Lin MT; Rios GP, 1985. Cowpea diseases and their prevalence in Latin America, In: Singh, SR, Rachie JO, eds. Cowpea Research, Production and Utilization. Chichester, UK: John Wiley and Sons, 199-204.
Lovisolo O; Conti M, 1966. Identification of an aphid-transmitted cowpea mosaic virus. Netherlands Journal of Plant Pathology, 72:265-269.
Mali VR; Mundhe GE; Patil NS; Kulthe KS, 1988. Detection and identification of blackeye cowpea mosaic and cowpea aphid borne mosaic viruses in India. International Journal of Tropical Plant Diseases, 6(2):159-173
Mali VR; Thottappilly G, 1986. Virus diseases of cowpeas in the Tropics. Review of tropical plant pathology. Volume 3. [edited by Raychaudhuri, S.P.; Verma, J.P.] New Delhi, India; Today and Tomorrow's Printers and Publishers, 361-403
Martín MT; Otín CL; Laín S; García JA, 1990. Determination of polyprotein processing sites by amino terminal sequencing of nonstructural proteins encoded by plum pox potyvirus. Virus Research, 15(2):97-106.
Matthews REF, 1979. Classification and nomenclature of viruses. Third Report of the International Committee on Taxonomy of Viruses. Intervirology, 12:131-296.
Maury Y; Khetarpal RK, 1989. Testing seed for viruses using ELISA. Perspectives in phytopathology [edited by Agrihotri, V.P.; Singh, N.; Chaube, H.S.; Singh, U.S.; Dwivedi, T.S.] New Delhi, India; Today and Tomorrow's Printers & Publishers, 31-49
McKern NM; Mink GI; Barnett OW; Mishra A; Whittaker LA; Silbernagel MJ; Ward CW; Shukla DD, 1992. Isolates of bean common mosaic virus comprising two distinct potyviruses. Phytopathology, 82(9):923-929
McKern NM; Strike PM; Barnett OW; Dijkstra J; Shukla DD; Ward CW, 1994. Cowpea aphid borne mosaic virus
McLean DM, 1941. Studies on mosaic of cowpea, Vigna sinensis. Phytopathology, 31:420-429.
Mih AM; Atiri GI; Thottappilly G, 1991. Relationships between co-infection with cowpea aphid-borne and cucumber mosaic viruses and yield of cowpea lines with varying resistance to these viruses. Phytoparasitica, 19(1):65-72
Mink GI; Silbernagel MJ, 1992. Serological and biological relationships among viruses in the bean common mosaic virus subgroup. Vienna, Austria: Springer-Verlag/Wien. Archives of Virology, Supplementum, 5:397-406
Mink GI; Vetten HJ; Ward CW; Berger PH; Morales FJ; Myers JM; Silbernagel MJ; Barnett OW, 1994. Taxonomy and classification of legume-infecting potyviruses. A proposal from the Potyviridae Study Group of the Plant Virus Subcommittee of the ICTV. Archives of Virology, 139:231-235.
Mlotshwa S; Verver J; Sithole-Niang I; Kampen Tvan; Kammen Avan; Wellink J, 2002. The genomic sequence of cowpea aphid-borne mosaic virus and its similarities with other potyviruses. Archives of Virology, 147(5):1043-1052.
Nariani TK; Kandaswamy TK, 1961. Studies on a mosaic disease of cowpea. Indian Phytopathology, 14:777-782.
Nascimento AVS; Santana EN; Braz ASK; Alfenas PF; Pio-Ribeiro G; Andrade GP; Carvalho MG; Zerbini FM, 2006. Cowpea aphid-borne mosaic virus (CABMV) is widespread in passionfruit in Brazil and causes passionfruit woodiness disease. Archives of Virology, 151:1797-1809.
Nascimento AVS; Souza ARR; Alfenas PF; Andrade GP; Carvalho MG; Pio-Ribeiro G; Zerbini FM, 2004. Phylogenetic analysis of potyviruses causing passionfruit woodiness in northeastern Brazil (in Portuguese). Brazilian Phytopathology, 29:378-383.
Ndiaye M; Bashir M; Keller KE; Hampton RO, 1993. Cowpea viruses in Senegal, West Africa: identification, distribution, seed transmission and sources of genetic resistance. Plant Disease, 77(10):999-1003
Pappu HR; Pappu SS; Sreenivasulu P, 1997. Molecular characterization and interviral homologies of a potyvirus infecting sesame (Sesamum indicum) in Georgia. Archives of Virology, 142(9):1919-1927; 18 ref.
Patel PN; Mligo JK, Leya. HK, Kuwite. C, Mmbaga ET, 1982. Sources of resistance, inheritance and breeding of cowpea for resistance to a strain of cowpea aphid-borne mosaic virus from Tanzania. Indian Journal of Genetics, 42:221-229.
Phatak HC, 1974. Seed-borne plant viruses - Identification and diagnosis in seed health testing. Seed Science and Technology, 2:3-155.
Roberts M; Harrison BD, 1979. Detection of potato roll and potato mop top viruses by immunoelectron microscopy. Annals of Applied Biology, 93:289-297.
Rossel HW; Thottappilly G, 1990. Possible dependence of geographical distribution of virus diseases of cowpea in African agroecological parameters. In: Allen DJ, ed. Proceedings of Working Group Meeting on Virus Diseases of Beans and Cowpeas in Africa. CIAT Africa Workshop Series No. 13. Cali, Colombia: Centro Internacional de Agricultura Tropical, 33-37.
Shukla DD; McKern MM, Ward. CW, Ford RE, 1991. Molecular parameters suggest that cowpea aphid-borne mosaic virus is a distinct potyvirus and bean common mosaic virus consists of at least three distinct potyviruses. Paper presented at the APS Annual Meeting, St. Louis, Missouri, 17-21 August 1991. Abstract no. 238.
Singh BB; Thottappilly G; Rossel HW, 1987. Breeding for multiple virus resistance in cowpea. Agronomy Abstracts, 79.
Singh SR; Allen DJ, 1979. Cowpea Pests and Diseases. Manual Series No.2. Ibadan, Nigeria: IITA.
Singh SR; Allen DJ, 1980. Pests, diseases, resistance and protection in cowpea. In: Summerfield RJ, Bunting AH, eds. Advances in Legume Science. London, UK: Her Majesty's Stationery Office, 419-443.
Sithole-Niang I; Nyathi T; Maxwell DP; Candresse T, 1996. Sequence of the 3
Snyder WC, 1942. A seed-borne mosaic of asparagus bean, Vigna sesquipedalis. Phytopathology, 32:518-523.
Taiwo MA; Gonsalves D; Provvidenti R; Thurston HD, 1982. Partial characterization and grouping of isolates of blackeye cowpea mosaic and cowpea aphidborne mosaic viruses. Phytopathology, 72(6):590-596
Thottappilly G; Hamilton RI; Huguenot C; Rossel HW; Furneaux MT; Gumedzoe MY; Shoyinka SA; Naik DM; Konate G; Atcham-Agneroh T; Haciwa HC; Anno-Nyako FO; Saifodine N; Wangai A; Lamptey P; Gubba A; Mbwaga AM; Neya J; Offei SK, 1995. Identification of cowpea viruses and their strains in tropical Africa - an international pilot project. IITA Research, No. 10:12-15; 11 ref.
Thottappilly G; Rossel HW, 1985. Worldwide occurrence and distribution of virus diseases, In: Singh SR., Rachie RO, eds. Cowpea Research, Production and Utilization. Chichester, UK: John Wiley and Sons, 155-171.
Thottappilly G; Rossel HW, 1997. Identification and characterization of viruses infecting bambara groundnut (Vigna subterranea) in Nigeria. International Journal of Pest Management, 43(3):177-185; 56 ref.
Tsuchizaki T; Hibino H, 1971. Seed transmission of viruses in cowpea and azuki bean plants. 4. Relation between seed transmission and virus distribution in apical meristem of flower bud. Annals of the Phytopathological Society of Japan, 37:17-21.
Tsuchizaki T; Senboku T; Iwaki M; Pholauporn S; Srithongchi W; Deema N; Ong CA, 1984. Blackeye cowpea mosaic virus from asparagus bean (Vigna sesquipedalis) in Thailand and Malaysia, and their relationships to a Japanese isolate. Annals of the Phytopathological Society of Japan, 50(4):461-468
Tsuchizaki T; Yora K; Asuyama H, 1970. The viruses causing mosaic of cowpea and azuki bean, and their transmissibility through seeds. Annals of the Phytopathological Society of Japan, 36:112-120.
Velsen RJ van, 1962. Cowpea mosaic, a virus disease of Vigna sinensis in New Guinea. Papua New Guinea Journal of Agriculture, 14:153-161.
Vidano C; Conti M, 1965. Transmission con afidi d'um "cowpea mosaic virus" isolate da Vigna sinensis Endl. in Italia. Atti dell'Accademia del Scienze di Torino, 99:1041-1050.
Wylie SJ; Jones MGK, 2011. The complete genome sequence of a Passion fruit woodiness virus isolate from Australia determined using deep sequencing, and its relationship to other potyviruses. Archives of Virology, 156(3):479-482. http://springerlink.metapress.com/content/4815uh5058304601/
Zettler FW; Evans IR, 1972. Blackeye cowpea mosaic virus in Florida: host range and incidence in certified cowpea seed. Proceedings of the Florida State Horticultural Society, 85:99-101.
Zhao GS; Baltensperger DD; Purcifull DE; Christie RG; Hiebert E; Edwardson JR, 1991. Host range, cytology, and transmission of an alyce-clover isolate of blackeye cowpea mosaic virus. Plant Disease, 75(3):251-253
ContributorsTop of page
22/12/98 Original text by:
Muhammad Bashir, National Agricultural Research Centre, Pakistan.
01/08/13 reviewed by:
Murilo Zerbini, Universidade Federal de Viçosa, Brazil
Distribution MapsTop of page
Unsupported Web Browser:
One or more of the features that are needed to show you the maps functionality are not available in the web browser that you are using.
Please consider upgrading your browser to the latest version or installing a new browser.
More information about modern web browsers can be found at http://browsehappy.com/