Fusarium oxysporum f.sp. vasinfectum (Fusarium wilt)
- 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
- Seedborne Aspects
- Plant Trade
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Fusarium oxysporum f.sp. vasinfectum (G.F. Atk.) W.C. Snyder & H.N. Hansen
Preferred Common Name
- Fusarium wilt
Other Scientific Names
- Fusarium vasinfectum G.F. Atk.
International Common Names
- English: Fusarium wilt of cotton
- Spanish: marchitez del algodonero; pudricion de la raiz
- French: flétrissement du cotonnier; flétrissement fusarien du cotonnier; fusariose
Local Common Names
- Germany: Baumwollewelke
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Fungi
- Phylum: Ascomycota
- Subphylum: Pezizomycotina
- Class: Sordariomycetes
- Subclass: Hypocreomycetidae
- Order: Hypocreales
- Family: Nectriaceae
- Genus: Fusarium
- Species: Fusarium oxysporum f.sp. vasinfectum
Notes on Taxonomy and NomenclatureTop of page
The causal organism of Fusarium wilt of cotton is Fusarium oxysporum f.sp. vasinfectum. The species, Fusarium oxysporum, is variable and contains a number of saprophytic and pathogenic forms which have morphological features in common and cannot be distinguished without the use of molecular tools and/or pathogenicity tests. The parasitic forms were grouped into formae speciales (f.sp.) by Snyder and Hansen (1940) on the basis of their selective pathogenicity to a particular plant species. The concept of formae speciales continues to evolve, especially with the advent of molecular approaches to characterize isolates (Edel-Hermann and Lecomte, 2019).
Within F. oxysporum f. sp. vasinfectum, six races of the pathogen are recognized on the basis of their selective pathogenicity to a range of differential hosts (Armstrong and Armstrong, 1958, Armstrong and Armstrong, 1960; Ibrahim, 1966; Armstrong and Armstrong, 1978; Sun et al., 1999; Edel-Hermann and Lecomte, 2019): 1, 2, 3, 4, 6 and 8. Previously-described races 5 and 7 were subsequently determined to be identical to races 3 and 4, respectively (Cianchetta and Davis, 2015). Some workers have used race A as a genetically-based combined designation of races 1, 2 and 6, as these races are differentiated on the basis of reactions of plants other than Gossypium spp. (Davis et al., 2006). There are two distinct biotypes of F. oxysporum f.sp. vasinfectum which are different from the races in other parts of the world (Kim et al., 2005). RAPD markers have been used to differentiate isolates of F. oxysporum f.sp. vasinfectum: 46 isolates from different geographical areas were tested and clustered into three groups corresponding to three of the races identified by inoculation into the differential hosts (Assigbetse et al., 1994). Isolates in China were divided into three RAPD sections one of which was similar to race 3 and two other races different from races known elsewhere, designated races 7 and 8. Isolates of race 7, the predominant race, were all placed in a single section while race 8 was divided into two RAPD sections (Feng et al., 1999). Vegetative compatibility groups have also been used to genetically distinguish isolates within races (Fernandez et al., 1994; Abo et al., 2005; Bell et al., 2016). Bell et al. (2016) recognized two pathotypes on the basis of plant symptoms and stem colonizing ability, specifically causing root rot but not colonizing the stem, or colonizing the stem but not causing root rot. The vascular-colonizing (‘vascular-competent’) pathotypes include races 1, 2, 6 and 8, while the root-rotting pathotypes include races 3, 4 and the Australian biotypes.
DescriptionTop of page
F. oxysporum f.sp. vasinfectum is an anamorphic fungus. No teleomorph is known. On potato dextrose agar (PDA), it produces a dense, white aerial mycelium and a red/purple pigment is produced in the medium. It produces two types of conidia. The microconidia are borne in false heads on short monophialides and are single celled or one-septate, oval, elliptical or kidney-shaped, and measure 5-20 x 2-3.5 µm. Macroconidia are fusiform-falcate in shape, three- to five-septate and 27-48 x 2.5-4.5 µm, with a foot-shaped basal cell and a curved, pointed apical cell, and are borne on monophialides on branched or unbranched conidiophores (Booth and Waterson, 1964; Booth, 1971). Macroconidia may be produced in orange sporodochia or from monophialides on hyphae. Chlamydospores are formed singly or in pairs and are roughly spherical in shape, 7-13 µm in diameter, with a thick wall. In PDA, brown, blue or black sclerotia may be formed by some isolates (Leslie and Summerell, 2006). F. oxysporum f.sp. vasinfectum cannot be morphologically distinguished from other formae speciales.
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.Last updated: 05 May 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Angola||Present, Localized||UK, CAB International (1982); Ragazzi et al. (1995)|
|Central African Republic||Present||IRCT (1977); UK, CAB International (1982)|
|Congo, Democratic Republic of the||Present||UK, CAB International (1982)|
|Côte d'Ivoire||Present, Widespread||Abo et al. (2005)|
|Egypt||Present, Localized||UK, CAB International (1982); Nirenberg et al. (1994)|
|Ethiopia||Present||UK, CAB International (1982)|
|Morocco||Present||UK, CAB International (1982)|
|Somalia||Present||UK, CAB International (1982)|
|South Africa||Present||UK, CAB International (1982)|
|Sudan||Present, Localized||UK, CAB International (1982); Yassin and Daffalla (1982)|
|Tanzania||Present, Widespread||UK, CAB International (1982); Hillocks (1984)|
|Togo||Present||Goebel and Vaissayre (1986)|
|Uganda||Present||UK, CAB International (1982)|
|Zimbabwe||Present||UK, CAB International (1982); Hillocks (1992)|
|Afghanistan||Present||UK, CAB International (1982)|
|Bangladesh||Present||UK, CAB International (1982)|
|China||Present||CABI (Undated a)||Present based on regional distribution.|
|-Anhui||Present||Guo et al. (1993)|
|-Hebei||Present||Guo et al. (1993)|
|-Henan||Present||Yang ZhiWei et al. (1995)|
|-Jiangsu||Present||Guo et al. (1993)|
|-Shaanxi||Present||Yang ZhiWei et al. (1995)|
|-Shandong||Present||Yang ZhiWei et al. (1995)|
|-Sichuan||Present, Widespread||He et al. (1994)|
|India||Present||UK, CAB International (1982)|
|-Haryana||Present||Chopra and Chauhan (1993)|
|-Maharashtra||Present||Khetmalas et al. (1989)|
|-Punjab||Present||Chopra and Chauhan (1993)|
|-Tamil Nadu||Present, Localized||Shanmugam et al. (1977)|
|Indonesia||Present||UK, CAB International (1982)|
|Iran||Present||UK, CAB International (1982)|
|Iraq||Present||UK, CAB International (1982)|
|Israel||Present||Dishon and Nevo (1970)|
|Japan||Present||UK, CAB International (1982)|
|-Honshu||Present||UK, CAB International (1982)|
|Myanmar||Present||UK, CAB International (1982)|
|North Korea||Present||Chung and Ser (1992)|
|Pakistan||Present||Anwar and Khan (1973); UK, CAB International (1982)|
|Saudi Arabia||Present||UK, CAB International (1982)|
|South Korea||Present||Chung and Ser (1992)|
|Turkey||Present||UK, CAB International (1982)|
|Turkmenistan||Present||Sidorova and Akmuradov (1981); UK, CAB International (1982)|
|Uzbekistan||Present||UK, CAB International (1982); Tursanov and Aliev (1995)|
|Vietnam||Present||UK, CAB International (1982)|
|Yemen||Present||UK, CAB International (1982)|
|Federal Republic of Yugoslavia||Present||UK, CAB International (1982)|
|Union of Soviet Socialist Republics||Present||UK, CAB International (1982)|
|France||Present||UK, CAB International (1982)|
|Greece||Present||UK, CAB International (1982)|
|Italy||Present||UK, CAB International (1982); CABI (Undated)|
|Romania||Present||UK, CAB International (1982)|
|Cuba||Present||UK, CAB International (1982)|
|El Salvador||Present||UK, CAB International (1982)|
|Guatemala||Present||UK, CAB International (1982)|
|Mexico||Present||UK, CAB International (1982)|
|Nicaragua||Present||UK, CAB International (1982)|
|Puerto Rico||Present||UK, CAB International (1982)|
|Saint Kitts and Nevis||Present||UK, CAB International (1982)|
|Saint Lucia||Present||Schotman (1989)|
|Saint Vincent and the Grenadines||Present||UK, CAB International (1982)|
|Trinidad and Tobago||Present||Schotman (1989)|
|United States||Present||CABI (Undated a)||Present based on regional distribution.|
|-Alabama||Present, Widespread||Blasingame (1990)|
|-Arizona||Present||UK, CAB International (1982); Blasingame (1990)|
|-California||Present, Widespread||Blasingame (1990); Kim et al. (2005)|
|-Georgia||Present, Localized||Blasingame (1990)|
|-Louisiana||Present, Localized||Blasingame (1990)|
|-Missouri||Present, Widespread||Blasingame (1990)|
|-New Mexico||Present, Localized||Zhu et al. (2020)|
|-North Carolina||Present||Blasingame (1990)|
|-South Carolina||Present, Widespread||Blasingame (1990)|
|-Texas||Present, Localized||Blasingame (1990); Halpern et al. (2018)|
|Australia||Present||CABI (Undated a)||Present based on regional distribution.|
|Fiji||Present||UK, CAB International (1982)|
|Argentina||Present||UK, CAB International (1982)|
|Bolivia||Present||UK, CAB International (1982)|
|Brazil||Present, Localized||Cia (1977); UK, CAB International (1982)|
|-Minas Gerais||Present||Chalfoun (1979)|
|-Sao Paulo||Present, Widespread||Gridi-Papp et al. (1984)|
|Chile||Present||UK, CAB International (1982)|
|Colombia||Present||UK, CAB International (1982)|
|Guyana||Present||UK, CAB International (1982)|
|Paraguay||Present||Mathieson and Follin (1981)|
|Peru||Present, Localized||UK, CAB International (1982); Delgado and Agurto (1984)|
|Uruguay||Present||UK, CAB International (1982)|
|Venezuela||Present||UK, CAB International (1982)|
Risk of IntroductionTop of page
Risk Criteria Category
Economic Importance Moderate
Seedborne Incidence Moderate
Seed Transmitted Yes
Seed Treatment Yes
Overall Risk Moderate
Notes on Phytosanitary Risk
Fusarium wilt is spread locally when soil carrying chlamydospores is moved from one field to another on farm implements and crop residues, or movement of soil as a result of furrow irrigation or flood water movement. Long-distance spread occurs when infected seed is used for planting (see Seedborne Aspects) and it is essential to ensure that seed multiplication does not occur on land infested with the disease and that seed for planting is stored and ginned separately from that harvested from infested sites. This is particularly important in countries such as Tanzania, where farmers' fields are used for seed multiplication. Seed from an unknown source should not be used for planting. This is a potential problem in East Africa where seed cotton is sometimes carried across international boundaries for sale. In countries affected by the disease where seed is used for oil extraction by pressure and the remaining husk used for cattle feed, there is a risk that the husk may still contain viable chlamydospores. There is a risk of introducing another race of the pathogen by importing seed from growing areas in which wilt occurs, but the pathogen is of a different race to that of the importing growing area. This is particularly true of China and Australia, which appear to have races of the pathogen that do not occur elsewhere.
Hosts/Species AffectedTop of page
Investigations into the host range of vascular wilt fusaria revealed that they had a wider host range than was originally allowed for in the system of Snyder and Hansen (1940). A concept of primary and secondary hosts was developed for the cotton wilt Fusarium (Armstrong and Armstrong, 1968) in which cotton is the primary host, with numerous secondary hosts on which the fungus can multiply but produces only mild symptoms or none at all. However, a plant species may be a secondary host for one race of the pathogen but a non-host for another race. For example, natural infections by race 1 have been recorded on Abelmoschus esculentus, but races 3 and 4 did not reproduce on this host (Grover and Singh, 1970). Results from artificial inoculation may suggest a much wider host range (e.g. Wood and Ebbels, 1972) but should be interpreted with caution unless the same species has also been shown to support the growth of the pathogen in the field. Reports on new hosts for the cotton wilt pathogen are only useful where the race of the pathogen is specified and isolations have been made consistently from naturally infected plants. A complete list of the known hosts up until 1974 based on natural and artificial infection, is given by Ebbels (1975).
Host Plants and Other Plants AffectedTop of page
|Abelmoschus esculentus (okra)||Malvaceae||Other|
|Cajanus cajan (pigeon pea)||Fabaceae||Main|
|Capsicum annuum (bell pepper)||Solanaceae||Main|
|Hevea brasiliensis (rubber)||Euphorbiaceae||Other|
|Hibiscus cannabinus (kenaf)||Malvaceae||Main|
|Nicotiana tabacum (tobacco)||Solanaceae||Other|
|Sesamum indicum (sesame)||Pedaliaceae||Other|
Growth StagesTop of page Flowering stage, Seedling stage, Vegetative growing stage
SymptomsTop of page
Symptoms of Fusarium wilt can appear at any stage of crop development. At high inoculum density or when infection initiates from the seed, plants may be killed at the seedling stage. In older plants, symptoms are first seen on the lower leaves. Leaf chlorosis begins at the margin and spreads between the main veins. More leaves become chlorotic as the disease spreads upwards in the plant and the leaves become flaccid, giving the plant a wilted appearance during the middle of the day. Symptomatic leaves may fall from the plant. Infected plants may also be stunted. As the infection progresses, all the leaves are affected and chlorosis turns to necrosis as the wilt becomes permanent and the plant dies from moisture stress.
With races that colonize the vascular system of the stem, plants with foliar or wilting symptoms will have brown to black discoloration of the xylem, seen when stems are cut. Roots will not be affected.
With root-rotting races, plants with foliar or wilting symptoms will have very limited vascular discoloration of the stem, limited to the lower stem and seen when there is also substantial root rot. Root rot can be seen as early as the one to true leaf growth stage and range from brown to black streaks in the centre of the root, to more extensive internal root rot, along with prominent external root rot symptoms. Often, there is no external root rot and diagnosis requires slicing the roots lengthwise. Susceptible Pima varieties tend to show more severe symptoms than susceptible Upland varieties. In some susceptible varieties, internal root rot may be present when there is no visible foliar or external root symptoms.
List of Symptoms/SignsTop of page
|Leaves / abnormal colours|
|Leaves / necrotic areas|
|Whole plant / dwarfing|
Biology and EcologyTop of page
F. oxysporum f.sp. vasinfectum is a soil-invading (Garrett, 1956), weak saprophyte that can remain dormant in the soil for long periods in the form of chlamydospores. The fungus is spread locally when soil carrying chlamydospores is moved from one field to another on farm implements and crop residues, or by water from flooding or furrow-irrigation. Spores in the soil germinate when fungistasis is overcome by exogenous nutrients provided by root exudates from a nearby root. The germ tube grows towards the cotton root and enters through a fissure in the epidermis or by direct penetration.
The fungus grows through the cortex to the stele and sporulates only when it has invaded the xylem. Conidia are then carried upwards in the transpiration stream, the fungus grows through the vessel end plates and then sporulates again in the adjoining vessel. Systemic spread within the plant occurs by spore transport; mycelial growth occurs in the vessels and later, to some extent, in the surrounding cortex. As the plant becomes completely infected, wilting and senescence occur as a result of water stress induced by the combined effect of mycelial growth in the xylem, fungal toxins and vascular occlusion by the host in an attempt to prevent systemic spread of the fungus. As the plant dies and tissues become moribund, the pathogen produces chlamydospores and the disease cycle is completed when host residues decay and spores are returned to the soil.
Cotton plants are predisposed to infection by vascular-colonizing F. oxysporum f.sp. vasinfectum isolates when the roots are invaded by nematodes (Smith and Dick, 1960; Jorgenson et al., 1978). The main predisposing nematodes are Meloidogyne spp. (Hillocks and Bridge, 1992), Belonolaimus longicaudatus (Cooper and Brodie, 1963) and Rotylenchulus spp. (Khadr et al., 1972). The predisposing effect is due to root wounding and nutrient accumulation at the nematode feeding site. In the case of Meloidogyne, the nematode affects the physiology of the host, interfering with resistance mechanisms against systemic infection by the wilt fungus, resulting in increased susceptibility (Hillocks, 1985). Root-rotting isolates of F. oxysporum f.sp. vasinfectum do not require nematode injury to infect roots and produce symptoms.
Seedborne AspectsTop of page
Infection of cotton seed by F. oxysporum f.sp. vasinfectum, at an incidence of 5%, was first demonstrated by Elliot (1923). It has also been detected on cotton seeds in East Africa (Perry, 1962), India (Kulkarni, 1934), the former Soviet Union (Gubanov and Sabirov, 1972) and West Africa (Lagiére, 1952). In Tanzania, most of the infected seed was derived from plants which developed wilt symptoms late in the growing season. Seed cotton harvested from these plants had infection levels of up to 21%. Susceptible varieties produced more infected seeds than resistant ones (Hillocks, 1983). Infection of seed with race 4 at less than 0.1% incidence was confirmed in California fields (Bennett et al., 2008).
F. oxysporum f.sp. vasinfectum is also seedborne in okra (Gangopadhyay and Kapoor, 1977).
F. oxysporum f.sp. vasinfectum can be seedborne and seed certification is recommended to avoid movement of the pathogen to non-infested areas (Robbs et al., 1972). In the absence of formal certification or regulatory programmes, growers should be aware and cautious if seed is originating from areas where the disease occurs. Hillocks (1981) showed that spread of the pathogen in Tanzania is reduced by issuing seed produced in Fusarium wilt-free areas. However, no quantitative estimates are available as to the proportion of infected seeds which give rise to infected plants.
According to Hillocks and Kibani (2002), the above phytosanitary measures instituted at the cotton ginneries to prevent the distribution, for planting, of seed infected with the wilt fungus have become difficult to apply since economic liberalization and the entry of the private sector into cotton ginning and lint marketing. Surveys of cotton fields, ginneries and cotton-buying posts were conducted in 1997 to determine the factors affecting disease incidence and spread. In affected fields, disease incidence was generally less than 5%. Where it was greater than this, wilt symptoms were associated with root damage caused by the root-knot nematode (Meloidogyne incognita). At a number of ginneries, herdsmen were allowed to remove seed husks that accumulate at the ginneries as a by-product of oil extraction. The husks are used as cattle feed and this was identified as a potential source of disease spread. At the buying posts visited, there was no system for separating cotton varieties or for identifying seed cotton purchased from villages infected with fusarium wilt. As a result, seed subsequently distributed for planting is likely to be a source of infection for the spread of this disease.
Soil is the primary inoculum source for F. oxysporum f.sp. vasinfectum. The pathogen is a soil-invading, weak saprophyte that can remain dormant in the soil for long periods in the form of chlamydospores (Garrett, 1956). The pathogen can also infect non-host plants without causing symptoms and perpetuate indefinitely.
The fungus could be associated with gin trash, as well as the manure from livestock fed contaminated cottonseed or gin trash.
Biological or chemical seed treatments have not been sufficiently effective or economical. Previous research evaluated chemical seed treatment with carbendazim, thiophanate-methyl and ethylene thiosulphonate (Shen, 1985; Sharma and Sandhu, 1989) and biological treatments with Trichoderma spp. (Charati et al., 1998).
Thermotherapy with hot water can substantially reduce seed contamination of Fusarium oxysporum f.sp. vasinfectum, but it also reduces seed viability and vigour (Bennett and Colyer, 2010; Doan and Davis, 2015).
Seed Health Tests
Culture plate (Hillocks, 1983)
- Cotton seed must be acid-delinted in concentrated sulphuric acid and surface sterilized to remove surface contamination.
- The seeds are placed on water agar or PDA in a Petri dish and incubated at 25°C for 5-7 days.
- Any fungus growing from the seeds should be subcultured onto fresh PDA for identification.
- Cultures confirmed as F. oxysporum are then inoculated into cotton seedlings to distinguish the pathogenic strains from the saprophytes.
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)||hyphae; spores||Yes||Yes||Pest or symptoms usually invisible|
ImpactTop of page
F. oxysporum f.sp. vasinfectum causes significant crop losses in several of the main cotton-producing countries. The disease is widespread in the USA, the former Soviet Union (Menlikiev, 1962) and China. In Africa, Tanzania is the worst affected country (Hillocks, 1981). Although losses nationally may not be great, estimated for instance at 0.2% for the whole of the USA in 1989 (Blasingame, 1990), losses are much greater in localized areas and for individual farmers in areas where the disease is endemic. The presence of a pathogen in a field can exclude the planting of a variety with desirable agronomic characteristics (.e.g. yield) if it is susceptible to the pathogen.
DiagnosisTop of page
The plant tissue to be used for isolations depends upon which plant parts are showing symptoms. With a root-rotting F. oxysporum f.sp. vasinfectum strain, there would be root rot or black streaking in the pith and this tissue should be targeted for isolations. With a vascular-colonizing strain, there would be browning of the xylem in stems and this tissue should be targeted for isolations. To isolate the fungus from an infected plant, cut a 1 cm section of tissue and surface sterilize in 70% ethyl alcohol or 1% NaOCl, then rinse in sterile water and place on PDA in a Petri dish. Incubate at 25°C for 3 days, by which time the fungus should be growing from the cut ends of the tissue and can be subcultured to fresh PDA for identification on the basis of cultural pigmentation, microconidia produced on a short conidiophore and the production of chlamydospores as the culture ages. Refer to Leslie and Summerell (2006) for details on morphological identification.
Fungus morphology, together with isolation from a cotton plant with wilt symptoms, should provide a good indication that the disease is Fusarium wilt. However, final confirmation requires the completion of Koch's postulates with the inoculation of the pathogen into a healthy cotton plant to produce symptoms of the disease. This can be done by growing the fungus on PDA and rinsing the surface of the culture with sterile water to bring the conidia into suspension. A spore suspension containing from 100,000 to 10,000,000 conidia/ml will produce symptoms in 7-10 days after inoculation by one of several methods. The most effective inoculation method is to dip the roots of a cotton seedling in the suspension and carefully repot the plant and keep it at a mean temperature above 25°C.
A PCR-based technique was developed which was capable of detecting the pathogen within host tissues, even when symptoms were absent (Moricca et al., 1998). A PCR protocol specific for race 4 was developed by Ortiz et al. (2017) and there is a PCR-based detection kit available specific for race 4 (Doan et al., 2014). These protocols are useful for detection from plants growing in the field but, to date, there are no effective diagnostic methods for analysing soil or seed.
Detection and InspectionTop of page
With root-rotting F. oxysporum f.sp. vasinfectum, bare spots within a field will have seedlings that have been killed. Wilting symptoms develop later in the season and root rot symptoms are easier to find at this time. With vascular-infecting strains, the disease is detectable in the field when foliar symptoms appear, usually 6-10 weeks after planting, and a tentative field diagnosis can be made with the finding of xylem browning in cut stems. Other pathogens or abiotic injuries and nutrient deficiencies can cause symptoms resembling Fusarium wilt, so additional laboratory tests are necessary for confirmation (see Diagnosis).
Similarities to Other Species/ConditionsTop of page
The symptoms of Fusarium wilt on cotton are similar to those of Verticillium wilt (Verticillium dahliae). Seedling death can be caused by other soilborne pathogens, such as Rhizoctonia solani.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
A more detailed review of control approaches can be found in Cianchetta and Davis (2015) and Davis et al. (2006). Control of F. oxysporum f.sp. vasinfectum is necessary in a number of countries, including China, the former Soviet Union, northern Brazil, Israel, Sudan, Egypt, Tanzania and some states in the USA. Once established in the soil, the fungus is practically impossible to eradicate. The most effective and practical means of control is exclusion from non-infested soils and where the pathogen is present and the use of resistant varieties.
Efforts for breeding resistance to F. oxysporum f.sp. vasinfectum are reviewed in Zhang et al. (2015) and include summaries of breeding activities in several countries. There is considerable variability for wilt resistance in the genus Gossypium; major gene resistance is found in some of the Sea Island (G. barbadense) cottons but not in the Upland types (G. hirsutum). Some of the Sea Island varieties are almost immune to the disease and the more resistant Upland cottons may have been derived from crosses or introgression with G. barbadense. The G. barbadense variety, Seabrook Sea Island, is a useful source of resistance (Wilhelm, 1981). However, it has proved difficult to retain high levels of resistance in lines derived from hybridization programmes between G. hirsutum and G. barbadense because resistance seems to be linked to barbadense characters which are undesirable in Upland varieties and are lost during selection for Upland agronomic traits. Furthermore, F. oxysporum f.sp. vasinfectum exists in a number of races (Armstrong and Armstrong, 1958; Armstrong and Armstrong, 1960; Ibrahim, 1966; Armstrong and Armstrong, 1978) and resistance to one race does not necessarily confer resistance to others.
The first wilt-resistant cottons were produced in the USA by mass selection on heavily infested land. Several states and private seed companies in the USA run evaluation programmes for resistance to the Fusarium wilt/Meloidogyne spp. nematode complex and new wilt-resistant varieties are continually being produced (Kappelman, 1980). More emphasis is now placed on selection for resistance to root-knot nematode in material with some resistance to Fusarium wilt as the best way to improve the level of resistance to the disease complex (Kappelman, 1975; Hyer et al., 1979). This has resulted in a series of lines derived from Auburn 56 known as Auburn RNR lines (Shepherd, 1982).
Commercially successful, wilt-resistant varieties have also been produced elsewhere. In Tanzania, UK77 and UK91 are Upland cottons derived from Albar 51 which have good levels of resistance to wilt but become susceptible in the presence of root-knot nematode (Hillocks, 1984; Hillocks and Bridge, 1992). Some of the Giza and Barakat varieties in Egypt are wilt resistant (Abdel-Raheem et al., 1974) and in the Sudan varieties for the areas of the Gezira which are affected by Fusarium wilt have to be selected for resistance to the disease (Yassin et al., 1986). The Egyptian long-staple varieties Ashmouni and Menoufi have been used in wilt resistance breeding programmes in the former Soviet Union (Wilhelm, 1981). 'Pima' cultivars, derived from G. barbadense are grown in parts of the USA, Israel, Peru and elsewhere. Some 'Pima' cultivars are wilt-resistant, such as L-60 DSV-UNP from Peru (Rodríguez-Gálvez and Maldonado, 1998).
Most of these breeding programmes rely on evaluation of wilt resistance using 'wilt-sick plots' of lines selected for other characters in the main breeding programme. Where single plant selection is to be practised then plants can be inoculated by root dip or stem puncture with a conidial suspension of the fungus (Hillocks, 1984; Hillocks, 1992).
As the pathogen cannot be eradicated from the soil once established, it is best to prevent its introduction by planting seed which is certified free of wilt (see Risk of Introduction and Seedborne Aspects). Do not plant seed originating from infested fields and the most cautious approach is to not plant seed originating from areas where Fusarium wilt is widespread.
Cultural Control and Sanitary Methods
F. oxysporum f.sp. vasinfectum can survive in the soil in the absence of its main host, cotton, by remaining dormant in the form of chlamydospores and by localized infection of the roots of a number of non-host plants among crop species and weeds (Wood and Ebbels, 1972; Smith and Snyder, 1975). Crop rotations are often ineffective at reducing the soil inoculum. However, in the former Soviet Union, rotations with barley and other crops reduced the incidence of wilt in cotton planted the following season (Goshaev, 1971). In California, a weed-free, dry, summer fallowing is recommended to reduce damage to future cotton crops.
Soil solarization is effective in decreasing the population of soilborne pathogens and has been shown to decrease the incidence of Fusarium wilt in Israel (Katan et al., 1983) but this approach, along with the chemical fumigation of soil, is not generally employed.
In localities with some infested fields, vehicles and farm implements need to be washed free of adhering soil prior to moving to non-infested fields. The use of detergents improves the effectiveness of washing (Bennett et al., 2011).
When furrow irrigation is used, limit tail water movement from infested fields.
Gin trash from infested fields should not be applied to non-infested fields. The manure of cattle fed cottonseed or gin trash from infested fields should not be applied to non-infested fields, nor should livestock grazing in infested fields be allowed into non-infested fields.
An IPM approach specifically for the control of Fusarium wilt on cotton has not been described but effective control of the disease in cotton requires that the use of resistant varieties is integrated with regulatory measures to prevent the use of infected seed for planting and, where applicable, the use of cropping systems which decrease the incidence of root-knot nematodes.
ReferencesTop of page
Abdel-Raheem A, Haggag MEA, Abou-Daoud MS, 1974. The reaction of some Egyptian cotton varieties and their crosses to Fusarium wilt. Zeitschrift fur Pflanzenkrankheiten und Pflanzenschutz, 81(9):516-521
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06/03/20 Review by:
Thomas Isakeit, Department of Plant Pathology, Texas A&M AgriLife Extension, College Station, Texas, USA.
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