Radopholus similis (burrowing nematode)
- 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
- Pathway Vectors
- 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
- Radopholus similis (Cobb, 1893) Thorne, 1949
Preferred Common Name
- burrowing nematode
Other Scientific Names
- Anguillulina acutocaudatus (Zimmermann, 1898) Goodey, 1932
- Anguillulina biformis (Cobb,1909) Goodey, 1932
- Anguillulina granulosa (Cobb, 1893) Goodey, 1932
- Anguillulina similis
- Radopholus acutocaudatus (Zimmermann, 1898) Siddiqi, 1986
- Radopholus biformis (Cobb, 1909) Siddiqi, 1986
- Radopholus citrophilus Huettel, Dickson & Kaplan, 1984
- Radopholus granulosus (Cobb, 1893) Siddiqi, 1986
- Radopholus similis citrophilus Huettel, Dickson & Kaplan, 1984
- Rotylenchus similis
- Tetylenchus granulosus (Cobb, 1893) Filipjev, 1936
- Tylenchorhynchus acutocaudatus (Zimmermann, 1898) Filipjev, 1934
- Tylenchus biformis Cobb, 1909
- Tylenchus granulosus
- Tylenchus similis
International Common Names
- English: banana burrowing nematode; black head disease of banana; citrus burrowing nematode; nematode root rot; pepper yellows nematode; slow wilt nematode; spreading decline of citrus
- Spanish: declinación propagante de los cítricos; nematodo coco; nematodo del banano (Argentina); nematodo del plátano (Mexico)
- French: anguillule mineuse du bananier
Local Common Names
- Brazil: nematoide cavernicola
- Sri Lanka: mid-country species of nematode
- RADOCI (Radopholus citrophilus)
- RADOSI (Radopholus similis)
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Nematoda
- Family: Pratylenchidae
- Genus: Radopholus
- Species: Radopholus similis
Notes on Taxonomy and NomenclatureTop of page Radopholus citrophilus, previously regarded, at least in some quarters, as a separate species from Radopholus similis is now accepted as being synonymous (Valette et al., 1998a; Elbadri et al., 1999; Kornobis, 1999).
DescriptionTop of page
Description (after Orton Williams and Siddiqi, 1973)
Body straight to slightly arcuate ventrally; cuticle distinctly annulated. Lateral field with 4 incisures, not areolated except towards extremities, arising from near median oesophageal bulb and ending near tail terminus; inner incisures coalescing near middle of tail. Lip region hemispherical, sometimes offset, usually with 3-4 annules; sclerotization strong; dorsal and ventral arms of framework not wider than submedians; lips 6, equal. Anterior cephalids just posterior to labial sclerotization. Spear about 18 µm long, with well developed round basal knobs which are usually indented anteriorly; dorsal knob sometimes appearing larger than subventrals. Median oesophageal bulb well developed, round to oval, valvular apparatus prominent. Oesophageal glands 3, in separate lobes, overlapping intestine dorsally and dorso-laterally; dorsal gland anterior. Hemizonid 3 annules long, just anterior to excretory pore which is at or just behind the level of the oesophago-intestinal valve. Vulva prominent, just postequatorial. Reproductive organs paired, opposed, outstretched. Spermathecae spherical, usually packed with small rod-shaped sperms. Ovaries generally with a single row of oocytes. Intestine filled with spherical granules, indistinctly overlapping rectum. Tail somewhat elongate-conoid with a narrow rounded or indented terminus.
Oesophagus and spear degenerate; median bulb and valvular apparatus indistinct, spear without distinct knobs. Lip region elevated, 4-lobed, with lateral lips considerably reduced, not strongly sclerotized, with 3-5 annules posteriorly. Hemizonid just anterior to excretory pore which is usually 2-3 body widths behind median oesophageal bulb. Single testis, outstretched anteriorly; spermatocytes in 3 rows followed by 5; spermatozoa rod-like. Bursa coarsely crenate, enveloping about two thirds of tail. Spicules strongly cephalated, 18-22 µm long, with pointed distal ends. Gubernaculum rod-like, protrusible, with distinct sharp claw-like titillae at distal end.
Note: Cobb (1893) published the descriptions of Tylenchus granulosus n. sp. and Tylenchus similis n. sp. from diseased banana plant material sent to him in New South Wales from Fiji in July, 1891. T. granulosus is the female and T. similis the male of R. similis, T. granulosus having page priority over T. similis. To preserve the well known name similis for this widely distributed economic pest, Sher (1968) proposed its retention, regarding T. granulosus as a senior synonym.
SEM studies of populations of R. similis collected from different countries showed differences in morphological characteristics, especially in the number of anterior hypoptygmata in the males and annules terminating the vulva of the females. Many of the Indonesian populations were found to have a forked tail end (Elbadri et al., 1999).
(Topotypes, after Sher, 1968)
L = 520-880 (690) µm; a = 22-30 (27); b = 4.7-7.4 (6.5); b' = 3.5-5.2 (4.5); c = 8-13 (10.6); c' = 2.9-4.0 (3.4); V = 55-61 (56); spear = 17-20 (19) µm; o = 12-20 (18).
L = 590-670 (630) µm; a = 31-44 (35); b = 6.1-6.6 (6.4); b' = 4.1-4.9 (4.8); c = 8-10 (9); c' = 5.1-6.7 (5.7); spear = 12-17 (14) µm; spicules = 19-22 (20) µm; gubernaculum = 8-12 (9) µm.
(Topotypes, after Taylor, 1969)
20 females (young):
L = 540-660 (605.2) µm; V = 53-58 (55.9); tail length = 55-77 (66.9) µm; spear = 18 µm; phasmids from tail terminus = 44-61 (53) µm.
6 females (gravid):
L = 610-745 (685.8) µm; V = 52-57 (55.3); tail length = 52-74 (59.8) µm; spear = 18 µm; eggs = 50-68 (56.1) µm x 19-30 (23.3) µm.
L = 535-650 (585) µm; tail length = 64-86 (72.8) µm; spear = 12 µm; spicules = 18-19 (18.2) µm; gubernaculum = 10-11 (10.3) µm; phasmids from tail terminus = 46-58 (53.7) µm.
DistributionTop of page R. similis was detected in imported Anthurium plants in a glasshouse in Israel in June 1999 (EPPO, 1999). Phytosanitary measures were imposed and CABI was informed in January 2006 that the status was 'Absent, intercepted only' by the Plant Protection and Inspection Services (PPIS), Israel.
See also CABI/EPPO (1998, Nos 166 & 167).
Records under the name R. citrophilus are included in the distribution (CABI/EPPO, 1999).
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|
|Brunei Darussalam||Present, few occurrences||Bridge, 1993; CABI/EPPO, 1999; EPPO, 2014|
|China||Eradicated||1988||CABI/EPPO, 1999; EPPO, 2014|
|-Fujian||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|India||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|-Andhra Pradesh||Present||Sundararaju, 2006|
|-Arunachal Pradesh||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Assam||Present||Khan, 1999; EPPO, 2014|
|-Bihar||Present||Khan, 1999; EPPO, 2014|
|-Goa||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Jammu and Kashmir||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Karnataka||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Kerala||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Madhya Pradesh||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Maharashtra||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Manipur||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Nagaland||Present||Khan, 1999; EPPO, 2014|
|-Odisha||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Tamil Nadu||Widespread||CABI/EPPO, 1999; EPPO, 2014|
|-Uttar Pradesh||Present||Khan, 1999; EPPO, 2014|
|-West Bengal||Present||Khan, 1999; EPPO, 2014|
|Indonesia||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Sumatra||Present||CABI/EPPO, 1999; EPPO, 2014|
|Israel||Absent, intercepted only||1999||CABI/EPPO, 1999; EPPO, 2014|
|Japan||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|Lebanon||Present||CABI/EPPO, 1999; EPPO, 2014|
|Malaysia||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|-Peninsular Malaysia||Present||CABI/EPPO, 1999; EPPO, 2014|
|Oman||Present||Waller and Bridge, 1978; CABI/EPPO, 1999; EPPO, 2014|
|Pakistan||Present||Shahina and Maqbool, 1992; CABI/EPPO, 1999; EPPO, 2014|
|Philippines||Present||Timm, 1965; CABI/EPPO, 1999; EPPO, 2014|
|Sri Lanka||Present||Sivapalan, 1968; Gnanapragasam et al., 1991; CABI/EPPO, 1999; EPPO, 2014|
|Taiwan||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|Thailand||Present||Timm, 1965; CABI/EPPO, 1999; EPPO, 2014|
|Turkey||Absent, confirmed by survey||EPPO, 2014|
|Yemen||Present, few occurrences||CABI/EPPO, 1999; EPPO, 2014|
|Burkina Faso||Present||EPPO, 2014|
|Burundi||Present||Bridge, 1988a; CABI/EPPO, 1999; EPPO, 2014|
|Cameroon||Present||Bridge et al., 1995; CABI/EPPO, 1999; EPPO, 2014|
|Central African Republic||Present||CABI/EPPO, 1999; EPPO, 2014|
|Congo||Present||Luc et al., 1964; CABI/EPPO, 1999; EPPO, 2014|
|Congo Democratic Republic||Present||Elmiligy and Geraert, 1971; CABI/EPPO, 1999; EPPO, 2014|
|Côte d'Ivoire||Widespread||Adiko, 1988; CABI/EPPO, 1999; EPPO, 2014|
|East Africa||Present||Gaidashova et al., 2009|
|Egypt||Present||CABI/EPPO, 1999; EPPO, 2014|
|Ethiopia||Present||O'Bannon, 1975; CABI/EPPO, 1999; EPPO, 2014|
|Gabon||Present||O'Bannon, 1977; CABI/EPPO, 1999; EPPO, 2014|
|Gambia||Present||Bridge, 1993; CABI/EPPO, 1999; EPPO, 2014|
|Ghana||Present||Addoh, 1971; CABI/EPPO, 1999; EPPO, 2014|
|Guinea||Present||Luc, 1968; CABI/EPPO, 1999; EPPO, 2014|
|Guinea-Bissau||Present||CABI/EPPO, 1999; EPPO, 2014|
|Kenya||Present||Ngundo and Taylor, 1973; CABI/EPPO, 1999; EPPO, 2014|
|Madagascar||Present||Luc, 1968; CABI/EPPO, 1999; EPPO, 2014|
|Malawi||Present||Saka and Siddiqi, 1979; CABI/EPPO, 1999; EPPO, 2014|
|Mauritius||Present||CABI/EPPO, 1999; EPPO, 2014|
|Morocco||Present||Sarah, 1989; CABI/EPPO, 1999; EPPO, 2014|
|Mozambique||Present||Evaristo, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Nigeria||Present||Caveness, 1965; CABI/EPPO, 1999; EPPO, 2014|
|Réunion||Present||Vilardebo and Guerout, 1976; CABI/EPPO, 1999; EPPO, 2014|
|Rwanda||Present||Gaidashova et al., 2009|
|Senegal||Present||Luc, 1968; CABI/EPPO, 1999; EPPO, 2014|
|Seychelles||Present||CABI/EPPO, 1999; EPPO, 2014|
|Somalia||Present||Beccari and Scavazzon, 1966; CABI/EPPO, 1999; EPPO, 2014|
|South Africa||Restricted distribution||Jones and Milne, 1982; CABI/EPPO, 1999; EPPO, 2014|
|Sudan||Present||Decker et al., 1980; CABI/EPPO, 1999; EPPO, 2014|
|Tanzania||Restricted distribution||Ngundo and Taylor, 1973; CABI/EPPO, 1999; EPPO, 2014|
|-Zanzibar||Present||Sebasigari and Stover, 1987|
|Uganda||Present||Ngundo and Taylor, 1973; CABI/EPPO, 1999; EPPO, 2014|
|Zambia||Present||Raemaekers and Patel, 1973; CABI/EPPO, 1999; EPPO, 2014|
|Zimbabwe||Present||Martin, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Canada||Present, few occurrences||CABI/EPPO, 1999; EPPO, 2014|
|-British Columbia||Present, few occurrences||CABI/EPPO, 1999; EPPO, 2014|
|Mexico||Present||Taboada and Caballero, 1968; CABI/EPPO, 1999; EPPO, 2014|
|USA||Restricted distribution||1953||CABI/EPPO, 1999; EPPO, 2014||R. similis is present in Florida, Hawaii, Louisiana and Texas. Establishment of the nematode outdoors in other states of the USA is unlikely due to unsuitable climatic conditions.|
|-Arizona||Absent, confirmed by survey||EPPO, 2014|
|-California||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|-Florida||Present||Suit and Ducharme, 1953; CABI/EPPO, 1999; EPPO, 2014|
|-Hawaii||Present||Sher, 1954; CABI/EPPO, 1999; EPPO, 2014|
|-Louisiana||Present||Suit and Ducharme, 1953; CABI/EPPO, 1999; EPPO, 2014|
|-Texas||Present, few occurrences||CABI/EPPO, 1999; EPPO, 2014|
Central America and Caribbean
|Barbados||Present||CABI/EPPO, 1999; EPPO, 2014|
|Belize||Present||Pinochet and Ventura, 1977; CABI/EPPO, 1999; EPPO, 2014|
|Costa Rica||Widespread||CABI/EPPO, 1999; EPPO, 2014|
|Cuba||Present||Stoyanov, 1967; CABI/EPPO, 1999; EPPO, 2014|
|Dominica||Present||Edmunds, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Dominican Republic||Present||CABI/EPPO, 1999; EPPO, 2014|
|El Salvador||Present||Wehunt and Edwards, 1968; CABI/EPPO, 1999; EPPO, 2014|
|French West Indies||Present||Risède et al., 2009|
|Grenada||Widespread||Edmunds, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Guadeloupe||Present||Scotto, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Guatemala||Present||Loos, 1961; CABI/EPPO, 1999; EPPO, 2014|
|Honduras||Present||Loos, 1961; CABI/EPPO, 1999; EPPO, 2014|
|Jamaica||Present||Cobb, 1915; CABI/EPPO, 1999; EPPO, 2014|
|Martinique||Widespread||Scotto, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Nicaragua||Present||Wehunt and Edwards, 1968; CABI/EPPO, 1999; EPPO, 2014|
|Panama||Present||CABI/EPPO, 1999; EPPO, 2014|
|Puerto Rico||Widespread||Roman et al., 1974; CABI/EPPO, 1999; EPPO, 2014|
|Saint Kitts and Nevis||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|Saint Lucia||Present||Introduced||Invasive||Edmunds, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Saint Vincent and the Grenadines||Widespread||Edmunds, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Trinidad and Tobago||Restricted distribution||Scotto, 1969; CABI/EPPO, 1999; EPPO, 2014|
|United States Virgin Islands||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|Windward Islands||Present||Williams et al., 2004|
|Argentina||Absent, formerly present||CABI/EPPO, 1999; EPPO, 2014|
|Bolivia||Present||Bridge et al., 1982; CABI/EPPO, 1999; EPPO, 2014|
|Brazil||Present||Zem and Lordello, 1983; CABI/EPPO, 1999; EPPO, 2014|
|-Alagoas||Present||Andrade et al., 2009|
|-Bahia||Present||Zem and Lordello, 1983; CABI/EPPO, 1999; EPPO, 2014|
|-Ceara||Present||Zem and Lordello, 1983; CABI/EPPO, 1999; EPPO, 2014|
|-Espirito Santo||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Minas Gerais||Present||CABI/EPPO, 1999; EPPO, 2014|
|-Pernambuco||Present||Costa et al., 2008|
|-Rio de Janeiro||Present||Zem and Lordello, 1983; CABI/EPPO, 1999; EPPO, 2014|
|-Santa Catarina||Present||Costa et al., 2008|
|-Sao Paulo||Present||Zem and Lordello, 1983; CABI/EPPO, 1999; EPPO, 2014|
|Colombia||Present||Loos, 1961; CABI/EPPO, 1999; EPPO, 2014|
|Ecuador||Restricted distribution||Bridge, 1976; CABI/EPPO, 1999; EPPO, 2014|
|French Guiana||Present||CABI/EPPO, 1999; EPPO, 2014|
|Guyana||Present||CABI/EPPO, 1999; EPPO, 2014|
|Peru||Present||Sasser et al., 1962; CABI/EPPO, 1999; EPPO, 2014|
|Suriname||Present||Maas, 1969; CABI/EPPO, 1999; EPPO, 2014|
|Venezuela||Present||Haddad et al., 1973; CABI/EPPO, 1999; EPPO, 2014|
|Austria||Absent, no pest record||EPPO, 2014|
|Belgium||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|Croatia||Absent, confirmed by survey||EPPO, 2014|
|Denmark||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|France||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|Germany||Absent, intercepted only||1970||CABI/EPPO, 1999; EPPO, 2014|
|Italy||Restricted distribution||1978||CABI/EPPO, 1999; EPPO, 2014|
|Netherlands||Restricted distribution||NPPO of the Netherlands, 2013; CABI/EPPO, 1999; EPPO, 2014|
|Poland||Absent, formerly present||CABI/EPPO, 1999; EPPO, 2014|
|Portugal||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|-Madeira||Eradicated||CABI/EPPO, 1999; EPPO, 2014|
|Slovenia||Restricted distribution||CABI/EPPO, 1999; EPPO, 2014|
|Sweden||Eradicated||198*||CABI/EPPO, 1999; EPPO, 2014|
|Switzerland||Absent, intercepted only||CABI/EPPO, 1999; EPPO, 2014|
|UK||Absent, intercepted only||CABI/EPPO, 1999; EPPO, 2014|
|American Samoa||Present||CABI/EPPO, 1999; EPPO, 2014|
|Australia||Restricted distribution||Blake, 1972; CABI/EPPO, 1999; EPPO, 2014|
|-Australian Northern Territory||Present||CABI/EPPO, 1999; EPPO, 2014|
|-New South Wales||Present||Blake, 1963; CABI/EPPO, 1999; EPPO, 2014|
|-Queensland||Widespread||Blake, 1963; CABI/EPPO, 1999; EPPO, 2014|
|-South Australia||Present, few occurrences||CABI/EPPO, 1999; EPPO, 2014|
|-Western Australia||Present||CABI/EPPO, 1999; EPPO, 2014|
|Cook Islands||Present||Grandison, 1990; CABI/EPPO, 1999; EPPO, 2014|
|Fiji||Present||Cobb, 1915; CABI/EPPO, 1999; EPPO, 2014|
|French Polynesia||Present||CABI/EPPO, 1999; EPPO, 2014|
|Guam||Present||Bridge, 1988b; CABI/EPPO, 1999; EPPO, 2014|
|Micronesia, Federated states of||Present||Bridge, 1988b; CABI/EPPO, 1999; EPPO, 2014|
|New Caledonia||Present||Grandison et al., 2009; EPPO, 2014|
|Niue||Present||Orton, 1980; CABI/EPPO, 1999; EPPO, 2014|
|Norfolk Island||Present||Khair, 1982; CABI/EPPO, 1999; EPPO, 2014|
|Palau||Restricted distribution||Bridge, 1988b; CABI/EPPO, 1999; EPPO, 2014|
|Papua New Guinea||Present||Bridge and Page, 1984; CABI/EPPO, 1999; EPPO, 2014|
|Samoa||Present||Orton, 1980; Grandison, 1996; CABI/EPPO, 1999; EPPO, 2014|
|Solomon Islands||Present||Bridge, 1988b; CABI/EPPO, 1999; EPPO, 2014|
|Tonga||Present||Kirby et al., 1980; CABI/EPPO, 1999; EPPO, 2014|
Risk of IntroductionTop of page R. similis is spread on infested vegetative planting material such as rootstocks, corms and tubers. It is a tropical nematode and can become a pest of any of the susceptible host crops in subtropical and tropical climates. Crops in temperate climates are not at risk.
R. similis now occurs in most tropical and subtropical areas of the world. It has been found to be widespread in almost all the banana- and plantain-growing regions of the world except Israel, the Canary Islands, Cape Verde Islands, Cyprus, Crete, Mauritius and Taiwan. It also appears to be absent from some of the important areas of production in the highlands of Eastern Africa.
Its worldwide distribution is relatively recent (beginning of the 19th century) and is due to the transfer of infected plant material, particularly banana sets, from country to country for commercial purposes. The wide distribution of R. similis is correlated with the areas where banana sets of the sub-group Cavendish (AAA) were imported. Adaption may cause the development of a wider host range as it spreads on different AAA, AAB and ABB clones in Africa, and on ornamental plants which increasingly are being exported to regions outside the tropics.
0ne of the most important means of spread of R. similis in tea areas is the dissemination of infested plants to fields from contaminated nurseries. The spread of nematodes could also occur through infested soil and water, poor soil conservation measures adopted in infested areas, use of contaminated irrigation water, use of infested planting material for intercropping tea areas, and the presence of alternative hosts in the vicinity of tea areas. Uprooting old tea fields from the bottom of the slope upwards could also cause a threat by exposing the newly planted, young tea to re-infestation from the infested soil above (Gnanapragasam, 1989a).
Hosts/Species AffectedTop of page Radopholus similis (sensu lato) is very polyphagous, attacking hundreds of plant species notably those belonging to the Rustaceae (Citrus and related genera) but also many other families including the Arecaceae, Musaceae, Poaceae, Brassicaceae, Rubiaceae and Solanaceae to name but a few.
It is a serious pest on commercial citrus in Florida and on banana, plantain, black pepper, ginger, coffee, tea, coconut, arecanut and other such crops in tropical and subtropical areas worldwide, with only a few exceptions (see Phytosanitary Risk).
For more detail on hosts of R. similis, see Christie (1959), Colbran (1964), Edwards and Wehunt (1971), O'Bannon (1977), Bridge (1988b), Gowen and Queneherve 1990), Koshy and Bridge (1990) and Gnanapragasam et al. (1991).
Host Plants and Other Plants AffectedTop of page
|Ananas comosus (pineapple)||Bromeliaceae||Main|
|Arachis hypogaea (groundnut)||Fabaceae||Main|
|Areca catechu (betelnut palm)||Arecaceae||Main|
|Camellia sinensis (tea)||Theaceae||Main|
|Chrysalidocarpus lutescens (butterfly palm)||Arecaceae||Habitat/association|
|Citrus aurantiifolia (lime)||Rutaceae||Other|
|Citrus reticulata (mandarin)||Rutaceae||Other|
|Citrus sinensis (navel orange)||Rutaceae||Other|
|Citrus x paradisi (grapefruit)||Rutaceae||Other|
|Cocos nucifera (coconut)||Arecaceae||Main|
|Coffea arabica (arabica coffee)||Rubiaceae||Main|
|Coffea canephora (robusta coffee)||Rubiaceae||Main|
|Curcuma longa (turmeric)||Zingiberaceae||Main|
|Daucus carota (carrot)||Apiaceae||Main|
|Illicium verum (star anise)||Illiciaceae||Other|
|Musa textilis (manila hemp)||Musaceae||Main|
|Musa x paradisiaca (plantain)||Musaceae||Main|
|Persea americana (avocado)||Lauraceae||Main|
|Pinus kesiya (khasya pine)||Pinaceae||Habitat/association|
|Piper betle (betel pepper)||Piperaceae||Main|
|Piper nigrum (black pepper)||Piperaceae||Main|
|Saccharum officinarum (sugarcane)||Poaceae||Main|
|Solanum lycopersicum (tomato)||Solanaceae||Main|
|Solanum nigrum (black nightshade)||Solanaceae||Wild host|
|Zea mays (maize)||Poaceae||Main|
|Zingiber officinale (ginger)||Zingiberaceae||Main|
Growth StagesTop of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage
SymptomsTop of page Bananas
The most obvious symptom of attack of R. similis on banana is the toppling over or uprooting of plants, especially those bearing fruit, but there is a range in damage severity, from the lengthening of the vegetative cycle to the drastic reduction in bunch weight. This reveals two types of damage that can occur in banana plantations; that affecting the anchorage of the plant, and less apparent, the effect on the plant's ability to take up water and nutrients and the subsequent effect on yield.
Macroscopically, several dark red lesions appear on the outer part of the root, penetrating throughout the cortex but not into the stele; adjacent lesions may coalesce and the cortical root tissue atrophies and later turns black. In heavy infestations the lesion girdles the roots. Nematodes migrate from infected roots into the corm causing black lesions which may then spread around the corm. Roots emerging become infected as they grow out of the corm. Uprooting occurs commonly in windstorms or if heavy rains loosen the soil (Gowen and Quénéhervé, 1990).
R. similis on black pepper is associated with pepper yellows (slow-wilt) disease, which appears as pale yellow or whitish-yellow drooping leaves on the vines. The number of such leaves increases gradually until large numbers of leaves, or even the entire foliage, become yellow. Yellowing is followed by shedding leaves, cessation of growth and dieback symptoms. The symptoms are well pronounced when soil moisture is depleted. Within 3-5 years of initiation of yellowing, all the leaves are shed and death of the vine takes place; hence the name slow-wilt disease.
In bearing vines, shedding of inflorescences is a major symptom. Large numbers of shed inflorescences are seen at the base of affected vines. In large plantations, affected patches become conspicuous initially as yellowed plants, and later with large numbers of barren standards that have lost the vines, or standards supporting dead vines without any leaves. Young and old plants are affected and the replanted vines normally die within 2 years.
The tender, thin, white feeding roots show typical orange- to purple-coloured lesions. Lesions are not clearly seen on older roots, being brown in colour. The root system exhibits extensive rotting and this results in a lack of fine feeder roots from the main roots. Extensive necrosis of larger lateral roots develops subsequently (Koshy and Bridge, 1990).
Ginger plants infected with R. similis exhibit stunting, reduced vigour and tillering. The topmost leaves become chlorotic with scorched tips. Affected plants tend to mature and dry out faster than unaffected healthy plants. Incipient infections of the rhizomes are evidenced by small, shallow, sunken, water-soaked lesions. The nematodes migrate intracellularly through tissues, producing large infection channels or galleries within the rhizomes.
In coconut, R. similis causes non-specific general decline symptoms such as stunting, yellowing, reduction in number and size of leaves and leaflets, delay in flowering, button shedding and reduced yield. R. similis infestation produces small, elongate, orange-coloured lesions on tender creamy-white roots. Tender roots of coconut seedlings with heavy infestation become spongy in texture. Surface cracks develop on the semi-hard, orange-coloured main roots. Lesions and rotting are confined to the tender portions of the root. Lesions are also not conspicuous on the secondary and tertiary roots as these are narrow and rot quickly on infestation (Griffith and Koshy, 1990).
Plants infested with R. similis induces 'yellow leaf' disease. Infestation produces small, elongate, orange lesions in young, succulent, creamy-white to light-orange portions of the main and lateral roots. The adjoining lesions coalesce and cause extensive rotting (Sundararaju, 1984) .
Tea plants infested with R. similis show similar symptoms to those caused by Pratylenchus loosi, such as stunting with twiggy branches, defoliation, premature flowering and fruiting. The roots of infested plants are sparse and dried compared with the whitish, succulent feeder roots of healthy plants. Although R. similis produces lesions on tea roots, they are very small compared to those formed by P. loosi (Gnanapragasam, 1983).
List of Symptoms/SignsTop of page
|Leaves / abnormal colours|
|Leaves / abnormal leaf fall|
|Leaves / wilting|
|Leaves / yellowed or dead|
|Roots / cortex with lesions|
|Roots / necrotic streaks or lesions|
|Roots / reduced root system|
|Vegetative organs / internal rotting or discoloration|
|Vegetative organs / surface lesions or discoloration|
|Whole plant / dwarfing|
|Whole plant / early senescence|
|Whole plant / uprooted or toppled|
Biology and EcologyTop of page R. similis is a migratory endoparasitic species which completes its life cycle within the root cortex and tissues of corms and tubers.
In bananas, penetration occurs mostly near the root tips, but nematodes can invade along the entire length of the root. Females and all juvenile stages are infective although males, morphologically degenerate (without stylet), are probably not parasitic. After entering the roots of banana, the nematodes occupy an intercellular position in the cortical parenchyma where they feed on the cytoplasm of nearby cells, causing cavities which then coalesce to appear as tunnels. Invasion of the stele is never observed, even in heavily infected roots. The presence of lignified and suberized layers in endodermal cells of endodermal layers limits invasion of the vascular bundle by R. similis. Phenolic compounds play a significant role in the host plant's defence response to the nematode. High levels of lignin, flavanoids, dopamine, cafeic esters and ferulic acids were associated with low levels of penetration in resistant cultivars (Valette et al., 1998b).
It is within infected tissues that females lay their eggs, with an average of four to five eggs per day for 2 weeks. The complete life cycle from egg to egg spans 20-25 days at a temperature range of 24-32°C, the eggs hatch after 8-10 days and the juvenile stages are completed in 10-13 days (Gowen and Quénéhervé, 1990; Loos, 1962).
In absence or reduced densities of competitors such as Helicotylenchus multicinctus, high populations of R. similis colonize the entire set of banana roots. The presence of competitors reduces the density of R. similis in the soil and roots and restricts it to the areas close to the rhizome (Queneherve, 1990).
R. similis forms a disease complex with Fusarium oxysporum f.sp cubense and the damage caused by R.similis was greater in the presence of the fungus. The percentage of root rots caused by the fungus was 6.5% in the presence of R. similis and 4% with the fungus alone (Abdel-Hadi et al., 1987).
In tea, R. similis is attracted to the growing part of the roots. It enters the cortical region and feeds on the cells, destroying them. Being an endoparasite, most of the population is found within the feeder roots in young tea. However, when the parasitized roots are severely damaged or become overparasitized, they move into the soil in search of fresh roots. In mature tea fields, large populations can be encountered within the roots and in the rhizosphere of infested bushes. Field observations and differential host trials indicate that different biological races of R. similis exist in tea areas (Gnanapragasam et al., 1991); this has been confirmed by RAPD analysis (Hahn et al., 1994).
R. similis is sensitive to cold and favours warm temperatures and moist soil conditions. In very wet or dry soil, populations of the nematode are found to decline (Gnanapragasam, 1993).
When P. loosi and R. similis are inoculated together on tea grown at high altitudes, the former rapidly takes over the latter by competitive displacement (Campos et al., 1990). On the other hand, when the tea cultivar is more susceptible to R. similis, populations of this species build up more than P. loosi at warmer temperatures (Gnanapragasam, 1991). In borderline areas, suitable for the build up of both species, it is common to encounter both R. similis and P. loosi. In semi-dry areas, R. similis also occurs with R. reniformis.
Soil type and texture has a significant influence on the reproductive rate and population build up of R. similis. Detailed experiments carried out in a temperature controlled water bath (25±1°C) showed the population to build up most rapidly in sandy soil, followed by gravelly or loamy soil. There was hardly any build up in clay soils. Damage to tea was significantly greater in gravelly soil, followed by sandy, then loamy soil (Gnanapragasam, 1990).
In coconut, R. similis takes about 25 days at 25-28°C to complete its life cycle. Most juveniles and adults, including gravid females, infest healthy, succulent root tips. In the field, the nematode can survive for 6 months in moist soil (27-36°C) and only 1 month in dry soil (29-39°C). Under glasshouse conditions it survives for longer periods: 15 months in moist soil (25.5-28.5°C) and 3 months in dry soil (27-31°C) (Griffith and Koshy, 1990).
The fungi Cylindrocarpon effusum, C. lucid and Cylindrocladium clavatum have been recorded in association with lesions produced by R. similis. When these fungi were inoculated simultaneously with the nematode, the rate of multiplication of R. similis was reduced, as was damage to coconut seedlings (Sosamma and Koshy, 1978, 1983; Koshy and Sosamma, 1987). Inoculation with mycorrhizal fungi also brought about a reduction in the population of R. similis (Koshy et al., 1998; Sosamma et al., 1999).
In black pepper, R. similis penetrates roots within 24 hours of inoculation and the cells around the site of penetration become brown. Nematodes do not enter the stelar portions of the root, but plugging of xylem vessels with a gum-like substance has been reported. The life cycle is completed within 25-30 days, at a temperature range of 21-31°C. In India, the maximum nematode population in roots of pepper occurs during September-October and the minimum population during April-June (Koshy and Bridge, 1990).
R. similis takes about 25-30 days to complete its life cycle on arecanut seedlings at 21-31°C under glasshouse conditions. The population of R. similis in arecanut fluctuates at different times of the year. In India, the maximum populations occur in the roots during 0ctober-November and minimum populations during March-June. Populations are also known to vary depending on the type of roots, palms, groves and soil types during the same period (Koshy and Sosamma, 1978).
The occurrence of R. similis was greater in sandy loam soil (42.3%) followed by laterite (29.6%) and alluvial 28.0%) soils. The maximum number of nematodes recorded was 139 per gram of root (Koshy et al., 1978).
Seedborne AspectsTop of page R. similis is disseminated mainly on banana planting/seed material such as corms and suckers.
Dissemination of the nematode in coconut plantations is mostly through infested seedlings.
Hot-water treatment can be used to eliminate nematodes from vegetative seed/planting material in bananas, plantains, ginger and turmeric.
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|
|Bulbs/Tubers/Corms/Rhizomes||adults; eggs; juveniles||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Growing medium accompanying plants||adults; eggs; juveniles||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Roots||adults; eggs; juveniles||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Seedlings/Micropropagated plants||adults; eggs; juveniles||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Stems (above ground)/Shoots/Trunks/Branches||adults; eggs; juveniles||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|True seeds (inc. grain)|
ImpactTop of page Bananas and Plantains
It is uncommon for bananas to be parasitized by monospecific nematode populations and the relative importance of the different species of other migratory endoparasites in mixed populations and their role in yield loss is not fully known. Most estimates of yield loss come from the use of nematicides that generally decrease populations of all species not only R. similis and can possibly cause other beneficial growth effects.
Where R. similis is the only major nematode present, yield improvement based on gross yield per hectare or weights of harvested bunches following nematicide application can be as high as 267%. Increase in banana and/or plantain yield has been recorded as 86% in Panama and 15% in Honduras (Wehunt and Edwards, 1968), 207-275% in Puerto Rico (over a 3-year period) (Roman et al., 1977), 5-30% in Australia, 71% in Ecuador and 267% in St Vincent (Gowen and Quénéhervé, 1990; Gowen, 1995) and 38% in South Africa (Jones and Milne, 1982). When R. similis is the main nematode species present in association with other nematodes, the yield improvement over untreated plants after nematicide application can range from 16 to 263% in Ivory Coast (Sarah, 1989; Gowen and Quénéhervé, 1990), 20-40% in Cameroon (Melin and Vilardebo, 1973; Vilardebo et al., 1988; Sarah, 1989), 35-40% in Madagascar (Beugnon and Vilardebo, 1974), 29-35% in Martinique (Gowen and Quénéhervé, 1990) and 46% in St Lucia (Gowen, 1975, Gowen and Quénéhervé, 1990).
Black Pepper (Piper nigrum)
R. similis is notorious for being associated with 'yellows disease' of black pepper which caused the loss of 20 million pepper 'trees' on the island of Banka, Indonesia by 1953 (Hubert, 1957). It was commented at the time that "this seems destined to become one of the instances in the history of agriculture where an important industry will be completely wiped out by a nematode" (Christie, 1959). The nematode is also involved in the same or a very similar disease in India known it that country as 'slow-wilt'. Subsequent publications have proven that R. similis is the primary causal agent of both yellows disease (Mustika, 1992) and slow wilt disease (Venkitesan and Setty, 1977; Mohandas and Ramana, 1991).
Pathogenicity tests under simulated field conditions on mature black pepper vines have given significant growth and yield reductions in vines inoculated with 100 or more nematodes. Yield reductions of 29, 50 and 59% are caused by initial R. similis populations of 100, 1000 and 10,000 respectively (Mohandas and Ramana, 1991). Glasshouse experiments on young black pepper plants have shown that all plant growth characteristics are reduced significantly by R. similis alone. After 4 months, initial populations of 1000 nematodes reduce plant heights by 26%, number of nodes by 31% and number of leaves by 39% (Mustika, 1992).
In Fiji, R. similis has been reported on ginger at infection levels in farmers' fields of more than 50% resulting in yield reductions approaching 40% (Vilsoni et al., 1976). In glasshouse experiments, R. similis causes reduction in growth and rhizome weight. Even an initial population of 10 nematodes per plant can cause a 40% reduction in rhizome weight; 100 nematodes results in a 42% reduction and very high populations of 10,000 nematodes will result in a 74% reduction in rhizome weight after 6 months. At this population level the ginger plants are killed due to severe rotting of rhizomes and roots (Sundararaju et al., 1979).
R. similis causes non-specific general decline symptoms on coconut such as stunting, yellowing, reduction in number and size of leaves and leaflets, delay in flowering, button shedding and reduced yield (Griffith and Koshy, 1990; Koshy et al., 1991). In pot experiments, soil population levels of 100 nematodes per seedling cause a 35% reduction in height and a 14% reduction in girth of coconut palms over a five year period (Koshy and Sosamma, 1987). In large field tanks (microplots) in India after seven years, an initial inoculum level of 1000 nematodes per seedling (10 nematodes per 35 640 cm³ of soil) gave reductions of 17, 14 and 35% over uninoculated control in height, number of leaves and girth of stem, respectively (Koshy et al., 1991).
Plants infested with R. similis significantly reduce the growth and vigour of arecanut palms. Pathogenicity experiments revealed that an initial population of 100 nematodes per seedling or one nematode in 800 g of lateritic soil could cause visible damage. The reduction over the control amounted to 23, 39, 25, 19 and 38% with respect to shoot length, shoot weight, girth at collar region, root length and root weight, respectively (Koshy, 1986).
A severe decline of citrus known as spreading decline, encountered only in the sandy soils of Florida, USA, is caused by R. similis. The nematode was for a period described as a separate species, Radopholus citrophilus, but has now been shown to be a different biological isolate, the citrus race, and not a different species (Kaplan and Opperman, 1997). In infested orchards in Florida, yield losses of 40-70% for oranges and 50-80% for grapefruit have been recorded. On decline trees, the reduction in fruit production varies with the age of the tree, citrus variety, farming practices in the orchards and the duration of the nematode infestation (DuCharme, 1968; Duncan and Cohn, 1990).
In Sri Lanka, decline in tea yields in the mid- and low-elevation tea estates is associated with moderate to high populations of R. similis (Campos et al., 1990). Since R. similis was found associated with P. loosi in many tea areas, it was difficult to study the yield loss under field conditions. However, results from pot experiments carried out at 25±1°C revealed severe damage to tea brought about by a low initial population level of 28 nematodes/100g of soil. When exposed to additional stress conditions in the field such as other pest attack, drought attack and poor soil condition, the damage threshold level could be even lower (Gnanapragasam and Herath, 1989).
Some of the popular clones such as TRI 2025 and TRI 2026, favoured by smallholders and widely planted at mid and lower altitudes are especially susceptible R. similis and have contributed to its spread. When infested, severe damage is encountered in nurseries, new plantings and mature tea areas, resulting in a complete failure to establish. This tea cultivar is now being discouraged from planting in areas prone to damage by R. similis.
DiagnosisTop of page
A diagnostic protocol for Radopholus similis is described in EPPO (2008).
Bananas and Plantains
Samples taken near to the base of the stem of the mother plant will contain roots of different ages and vigour, and consist predominantly of primary roots with relatively smaller quantities of secondary and perhaps no tertiary roots. It is in this region that roots will contain highest populations of R. similis in the root cortex. Generally, the greatest numbers of nematodes occur in the roots of the most actively growing suckers. Samples are best collected within a horizontal distance of up to 30 cm from the base of the plant and down to a depth of 30 cm from the soil surface (Araya et al., 1999).
The techniques used to extract R. similis from bananas may depend on the available laboratory facilities and assistance, and use may be made of non-standard materials purchased locally. This should not prevent or discourage nematologists from adapting a technique which can be used routinely by different operators to give reproducible results.
Whatever extraction procedure is used it is important to obtain a representative root sample which should be chopped in 0.5 cm lengths, mixed thoroughly, and a 25-g subsample taken for processing. A 24-hour period of incubation is sufficient for macerated root samples. Chopped roots should be incubated for 2-4 days and mist extractions may be run for up to 14 days in some laboratories.
It is customary to report nematode populations per 100 g of fresh roots although this quantity is seldom used for extractions (Gowen and Quénéhervé, 1990).
Where nematologists or laboratory facilities are unavailable, R. similis damage is sometimes assessed by recording incidence of uprooting per hectare per month. This may also be correlated with assessments of necrosis on primary roots and on rhizomes taken from randomly selected plants from a plantation. Such techniques can be used by those who are familiar with nematode symptoms but care should be taken not to confuse lesions caused by plant parasitic nematodes with those resulting from other root-infesting pests and pathogens.
The presence of R. similis and its association with yellows or slow wilt disease can be diagnosed by soil sampling at a distance of 25-50 cm from the base of the vine at a depth of 20-30 cm. A soil sample of 200 cubic centimetres and a root sample of 0.5 to 1.0 g thin, tender, feeder roots will yield maximum nematode population (Koshy and Bridge, 1990).
R. similis-infested roots, showing lesions and rotting, may be split longitudinally and cut to a length of 1 to 2 cm. When such roots are submerged in water contained in Petri dishes or shallow pans and incubated at 20-25°C, 50% of nematodes are released in 72 hours. For collecting active nematode populations for culturing and other studies, tease out individual root lesions in water contained in a watch glass under a stereoscopic microscope and quickly transfer the nematodes into fresh water.
Soil and root samples are collected when the nematode population is highest (0ctober-November in India). Maximum populations are found on coconut at a distance of 100 cm from the roots of the palm and at a depth of 50-100 cm.
The tender, semi-hard, orange main roots are peeled and sliced longitudinally into four to eight pieces with lengths of 3-5 cm. The root slices are submerged in water in a Petri dish at 20-25°C. After an incubation period of 24 h, the water is changed and the nematodes are extracted after 72 h (Koshy et al., 1975; Koshy, 1986).
To detect R. similis in arecanut palms, soil and root samples should be collected at a distance of 25-75 cm from the bole, at a depth of 25-75 cm during peak periods (e.g. 0ctober/November in India). The extraction method is similar to that used for coconut (Griffith and Koshy, 1990).
Above-ground symptoms of R. similis attack on tea are often confused with symptoms induced by infestation with P. loosi or other causes of restricted root growth. Positive diagnosis of R. similis requires sampling of both soil and roots from suspected areas in the field.
Soil and root sampling
Tea soils are usually sampled when the soil has been adequately moist for a continuous period of at least 3-4 weeks. Soil is collected at a depth of 15-25 cm and at a distance of 15 cm from the base of the plant. Several samples are generally collected from a given field, with approximately 25-30 samples taken for every 2 hectares. As the build up of nematodes varies in different tea cultivars, composite samples are collected separately from the individual cultivar at a specific location. It is necessary to collect feeder root samples from several random points because most of the nematodes in young plants remain within the roots.
Detection and InspectionTop of page Bananas/Plantains
The presence of R. similis can be detected by inspecting and extracting nematodes from roots and bases of pseudostems (corms) in bananas and plantains; from roots and surrounding soil in crops such as black pepper, coconut, betel vine and sugarcane; and mainly from rhizomes, corms and tubers in ginger, turmeric, taros and yams.
In tea, examination of the roots of infested plants shows a marked reduction in the feeder root system. Infested roots appear dark and dried with small, brown lesions. A positive diagnosis of R. similis can only be achieved by sampling the soil and roots from affected areas and by extracting the nematodes. Extraction from the roots is mostly carried out using a modified Baerman funnel technique (Hutchinson, 1962).
In coconut, R. similis can be detected by examining the creamy-white to orange, semi-hard, main roots and checking for lesions.
Techniques for sampling and methods used for extracting nematodes from plant tissues are described in the Diagnostic Methods section.
Similarities to Other Species/ConditionsTop of page R. similis has a superficial resemblance to the genus Pratylenchus, but can be distinguished by having a median vulva with two genital tracts in the female as opposed to a posterior vulva with one tract. It can be most easily distinguished from other species of Radopholus by the length of the female tail. R. similis is also similar to Hirschmanniella spp. Molecular methodologies are being increasingly employed to investigate the diversity of R. similis populations.
Prevention and ControlTop of page
In Bananas and Plantains
The different practices used for controlling nematodes in bananas are summarized below. In permanent cultivation, the opportunities for control are limited to regular nematicide treatment; however, in subsistence cultivation, the only realistic or economically justifiable techniques for preserving losses from nematodes may be by applying large quantities of mulch to stimulate root growth, and by propping fruit stems. Several of the techniques used for nematode control are also appropriate for controlling the banana borer Cosmopolites sordidus, which is a widespread pest causing damage to banana corms.
The selection of appropriate control techniques will depend largely on the conditions, availability and reliability of workers, and economic considerations. Most control methods depend on the skill and experience of the operators and may be of little value if the work is not supervised.
The following are established practices for decreasing nematode populations in two different banana-growing systems (replanted systems and permanent plantations).
- rotation with alternative crops for 2-3 years
- flooding for 8 weeks after having destroyed previous banana crop
- fallow in absence of banana volunteers for 10-12 months
- selection of disease-free suckers
- use of in vitro-produced plants
- paring diseased tissue from corm
- paring and leaving large corms in sun for 14 days
- immersing corms in hot water
- coating corms with nematicide in mud
- applying nematicide to planting hole and in-fill soil
- regular spot applications with granular or liquid nematicide formulations.
- Planting resistant varieties
- Biological control using endophytic fungi (e.g. Fusarium sp.) (Pocasangre et al., 2000) and mycorrhizal fungi (Koshy et al., 1998).
- regular spot applications with granular or liquid nematicide formulations
- heavy mulches with organic wastes may have beneficial root growth effects
- propping fruiting stems with poles or with string guy ropes may prevent plants uprooting.
- inoculation with mycorrhizal fungi
For further information see Gowen and Queneherve (1990).
In Black Pepper
At present there are no effective control measures for control of slow-wilt or pepper yellows disease. The price of black pepper is known to fluctuate greatly and with the fall in prices, the farmer often loses.
Integrated methods of nematode management that can be suggested are:
- planting of nematode-free rooted cuttings
- uprooting of affected vines and replanting after a period of 9-12 months
- use of non-living supports or standards
- exclusion of R. similis-susceptible trees as standards for trailing black pepper vines
- exclusion of susceptible intercrops such as banana, ginger and turmeric
- application of organic amendments, such as neem oil cake (Azadirachta indica), green foliage, or farm-yard manure
- earthing-up after application of nematicides, NPK fertilizers and organic amendments
For further information see Koshy and Bridge (1990).
The following are suggested control measures in coconut:
- application of cow dung, farm yard manure, oil cakes and green manures to the basins
- applications of chemicals
- avoiding use of banana as a shade crop in coconut nurseries
- use of nematode-free planting material of coconut and other intercrops
- use of tolerant or less susceptible cultivars
Control of R. similis is difficult in arecanut. The use of nematicides is discouraged as it might cause problems of residual toxicity. The common methods of control that have been recommended include:
- use of nematode-free planting material of arecanut and other intercrops - avoiding crops that are susceptible to R. similis (e.g. banana, black pepper) as intercrops
- minimum use of nematicides
- use of any available resistant/tolerant cultivar (Griffith and Koshy, 1990) - treatment with Neem oil cake, either alone or in combination with chemicals (Sudha et al., 1998)
- inoculation with Paecilomyces lilacinus (Sudha et al., 2000)
The main methods for controlling R. similis in ginger are by growing on land where ginger has not been grown in the previous season and has no history of nematode infestation; and by using nematode-free planting material.
Production of nematode-free planting material is by:
- selecting only nematode-free material for planting; all seed rhizomes with external symptoms of nematode infestation should be discarded
- hot-water treatment of ginger seed material at a temperature of 50°C for 10 minutes.
In Sri Lanka, an integrated management technique is adopted for control of R. similis and prevention of its spread in tea areas (Gnanapragasam, 1995). 0nly limited amounts of chemicals are used in suppressing nematodes, with more emphasis on non-chemical means of control. The most important non-chemical means of control is the use of resistant/tolerant clones. Several varieties of tea that are resistant/tolerant to R. similis have been recommended for planting in tea areas of Sri Lanka (Gnanapragasam, 1989c, 1991).
Other strategies include:
- use of nematode free planting material
- growing of non-host plants or resistant varieties of crops in intercropping tea areas
- use of soil amendments
- use of a limited amount of chemicals at the time of planting
- use of botanicals
- use of trap crops
- use of cultural methods
Much work has been done in the past decade on assessing varieties of Musa for resistance to R. similis. Amongst others, Fogain and Gowen (1996), Pinochet et al. (1996), Sarah et al. (1997) and Marin et al. (1998) discussed the possible mechanisms of resistance to nematodes in Musa. Factors such as the presence of polyphenolics and the extent of lignification in the roots may be linked to resistance. Many other studies on the interaction of Musa germplasm and nematodes have been done, including those by Fogain et al. (1996), Sarah et al. (1997), Dinardo-Miranda and Teixeira (1996), Collingborn and Gowen (1997), Collingborn et al. (1998) and Fogain and Gowen (1998). Yangambi Km5 has been suggested as a possible source of resistance to R. similis and Pratylenchus goodeyi.
Sudha et al. (1998) screened 25 coconut cultivars against R. similis, but all proved susceptible with Philippines Ordinary being the least susceptible and Chougat Orange Dwarf the most susceptible.
Tea is intercropped with pepper in middle and lower altitude growing areas in Sri Lanka, especially in smallholdings. Resistant varieties of pepper selected to be recommended for use in these areas (Gnanapragasam, 1989b).
ReferencesTop of page
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