Ditylenchus africanus (peanut pod 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
- Ditylenchus africanus Wendt et al., 1995
Preferred Common Name
- peanut pod nematode
Other Scientific Names
- Ditylenchus destructor Thorne, 1945
Local Common Names
- South Africa: grondboonpeulaalwurm; groundnut pod nematode
- DITYAF (Ditylenchus africanus)
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Nematoda
- Class: Secernentea
- Order: Tylenchida
- Family: Anguinidae
- Genus: Ditylenchus
- Species: Ditylenchus africanus
Notes on Taxonomy and NomenclatureTop of page The peanut pod nematode was first discovered in 1987 (De Waele et al., 1989) when it was soaked from hulls and seeds showing symptoms resembling those caused by the fungus Chalara elegans. It was first identified as Ditylenchus destructor (De Waele et al., 1989). Since it does not damage potatoes (De Waele et al., 1991) and thrives at high temperatures around 28 to 30°C (De Waele and Wilken, 1990) it was considered a distinct race from the populations found in Europe and the USA.
However, a molecular study of comparative taxonomy of some populations of Ditylenchus by Wendt (1992) threw doubt on this classification. A subsequent study based on characteristics of morphology and restriction fragment length polymorphisms (RFLPs) of ribosomal DNA, described the South African population of D. destructor as Ditylenchus africanus sp. n.
DescriptionTop of page The morphometrics of the South African population agreed with those reported for D. destructor: 6 to 11 lateral incisures, a rounded tail tip, and a long post-uterine sac relative to the vulva-anus distance (De Waele et al., 1989).
De Waele et al. (1989) and Wendt et al. (1995), give detailed descriptions of the adult females and males (larvae not described). The following description is given by Wendt et al. (1995):
Female: Head flattened, about 1.3 µm high and 6.4 - 7.3 µm wide, not offset from, but narrower than rest of body. SEM shows labial area with pore-like stoma opening surrounded by six outer labial sense organs and two large, medial lips, each with a pair of cephalic sensillae. Outline of labial area and head region hexagonal. Amphidial aperture elliptical, directed towards stomal opening. First head annule discontinuous, caused by position of amphidial apertures. Apart from labial disc, four lip annuli in lip region. Stylet delicate, knobs distinct, separated, sloping backwards; shaft about 60% of total stylet length. Median bulb with crescentic valves. Basal bulb overlapping intestine. Postvulval uterine sac 50 -143 (79.2+-21) µm long, comprising about 8% of total body length or 37 - 85% of vulva-anus distance and equal to 1.5 - 3.7 times vulval body diameter. Egg measurements: 45 - 60 µm x 20.5 - 33.5 µm. Tail elongate-conoid, tapering in posterior one-third to a finely rounded terminus.
Male: Bursa 33 - 60 (47+-8.6) µm long, leptoderan, covering 48-66% of tail length. Spicule arcuate ventrad, slightly cephalated.
DistributionTop of page Groundnut pods with symptoms very similar to those caused by D. africanus have been collected from various countries in southern Africa, but since it is difficult to retrieve live nematodes from dried seed, these results have not been verified (C Venter, ARC-Grain Crops Institute, Potchefstroom 2520, South Africa, personal communication, 1997. Present address: Green Circle Crop Protection, 14 Krom Street, Potchefstroom 2531, SA).
See also CABI/EPPO (1998, No. 159).
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.
Risk of IntroductionTop of page RISK CRITERIA CATEGORY
ECONOMIC IMPORTANCE Moderate
DISTRIBUTION South Africa
SEEDBORNE INCIDENCE Moderate
SEED TRANSMITTED Yes
SEED TREATMENT None
OVERALL RISK High
Notes on phytosanitary risk
D. africanus is widespread within South Africa (De Waele et al., 1989). There are therefore no quarantine regions within South Africa. The limited distribution of this economically important pathogen would warrant phytosanitary regulations to protect against its introduction by infected groundnut seeds.
Hosts/Species AffectedTop of page The earliest report of D. africanus on groundnuts was made by De Waele et al. (1989).
The crops (Basson et al., 1990) and weeds (De Waele et al., 1990) listed were hosts under glasshouse conditions. The fungal hosts Chalara spp., Penicillium spp., Phytophthora spp., Aspergillus spp. and Fusarium spp. were found under laboratory conditions (C Venter, ARC-Grain Crops Institute, Potchefstroom 2520, South Africa, personal communication, 1997. Present address: Green Circle Crop Protection, 14 Krom Street, Potchefstroom 2531, SA).
Host Plants and Other Plants AffectedTop of page
|Arachis hypogaea (groundnut)||Fabaceae||Main|
|Chenopodium album (fat hen)||Chenopodiaceae||Other|
|Datura stramonium (jimsonweed)||Solanaceae||Other|
|Eleusine indica (goose grass)||Poaceae||Other|
|Glycine max (soyabean)||Fabaceae||Other|
|Gossypium hirsutum (Bourbon cotton)||Malvaceae||Other|
|Helianthus annuus (sunflower)||Asteraceae||Other|
|Lupinus albus (white lupine)||Fabaceae||Other|
|Medicago sativa (lucerne)||Fabaceae||Other|
|Nicotiana tabacum (tobacco)||Solanaceae||Other|
|Phaseolus vulgaris (common bean)||Fabaceae||Other|
|Pisum sativum (pea)||Fabaceae||Other|
|Solanum tuberosum (potato)||Solanaceae||Other|
|Sorghum bicolor (sorghum)||Poaceae||Other|
|Tagetes minuta (stinking Roger)||Asteraceae||Other|
|Triticum aestivum (wheat)||Poaceae||Other|
|Vigna unguiculata (cowpea)||Fabaceae||Other|
|Xanthium strumarium (common cocklebur)||Asteraceae||Other|
|Zea mays (maize)||Poaceae||Other|
Growth StagesTop of page Fruiting stage
SymptomsTop of page D. africanus does not cause visible lesions on the roots of groundnuts, but affects the appearance, and sometimes the weight and untimely germination, of the pods and seeds (Venter et al., 1991). Infected hulls also show greyish-black to brown necrotic tissue at the point of connection with the peg, and in broad bands along the longitudinal veins. Infected seeds are usually shrunken with dark brown to black micropyles and yellow to dark flaccid testae with dark vascular strands. The embryos may also become darkly discoloured (Jones and De Waele, 1990).
No symptoms are visible on the roots of alternate hosts (Basson et al., 1990) or the tubers of potatoes (De Waele et al., 1991).
List of Symptoms/SignsTop of page
|Seeds / lesions on seeds|
Biology and EcologyTop of page The biology of D. africanus is very closely related to that of the groundnut plant. The nematode apparently survives in the soil, on fungi and the roots of groundnut and alternate hosts and weeds, in very low numbers until the groundnut pegs appear in the soil (around 8 weeks after sowing). Hereafter the nematode penetrates the immature pod at its connection with the peg, entering the exocarp and moving either longitudinally in this cell layer towards the beak-end of the pod (hence the pod discolouration), or through the mesocarp into the endocarp of the hull (shell) (Venter et al., 1995). It then migrates to the seed micropyle from where it invades the seed testa and embryo (giving rise to the seed symptoms). The nematode has not been found within the cotyledons of the seed (Jones and De Waele, 1990).
The nematode reproduces within the pod and seed, and at 28°C the life-cycle from adult to adult is 6 to 7 days (De Waele and Wilken, 1990).
As physiological maturity approaches (from 17 to 21 weeks after sowing, varying between cultivars) the relative number of eggs and anhydrobiotes in the pod and seed tissues increases (Venter et al., 1995).
Post harvest, the nematode is able to survive in planting seed, which may be symptomless, and in the field soil in the absence of host plants, or in hulls buried in the soil, for at least 28 to 32 weeks (Basson et al., 1993). This is long enough to survive the dry winter season in South Africa.
With the first spring rains the eggs hatch and anhydrobiotes rehydrate. Basson et al. (1993) showed that rehydrated soil populations of D. africanus are able to invade the next crop of groundnut and cause damage. Although relatively few nematodes survive in whole stored seed at 10°C, the surviving nematode population is also able to build up to levels capable of causing damage to the next crop (Basson et al., 1993).
Seedborne AspectsTop of page Incidence
A 1987 survey of damaged seeds obtained from 877 farms representative of the major groundnut production areas, showed that 73% of the samples were infected with D. africanus (De Waele et al., 1989). The average number of nematodes per seed was 160, with up to 3338 per seed in the south western Transvaal, the area in which the pest was first found. D. africanus is found within the seed and seed embryo, but not within the cotyledons (Jones and De Waele, 1990). In an extensive histological study, Venter (1995) showed that D. africanus entered the immature pegs and pods of groundnut (Arachis hypogaea cv. Sellie) at the peg-connection and subsequently invaded the parenchymatous regions of the hull exocarp and endocarp, and eventually the seed testa. The nematode caused malformations of the cells of infected tissues, cell wall breakage, and cell collapse. The damage appeared to be due to enzymatic activity. In some testae the entire parenchyma region, which aids in protection of the seed, was destroyed. In immature pods, the nematodes moved across the fibrous region of the mesocarp into the hull endocarp. In mature pods, however, the fibrous mesocarp of the hull was lignified and apparently was a barrier to penetration of the inner pod tissues. In late-harvested pods, increased numbers of eggs and anhydrobiotes were found in the hull tissues, and eggs in the seed testa, suggesting the onset of winter survival mechanisms of the nematode.
Effect On Seed Quality
D. africanus causes destruction of the parenchymous middle layer of the seed testa (Venter et al., 1995). This results in the flaccid yellow to dark brown testa. With heavy infections, seed mass may be decreased, and in some cultivars, especially if pre-harvest rains occur, the damaged hulls split and the seeds germinate into seedlings around the mother plant (Venter et al., 1991). Many infected seeds, however, may have no symptoms (Bolton et al., 1990).
The nematode reproduces within the pod and seed, and at 28°C the life-cycle from adult to adult is 6 to 7 days (De Waele and Wilken, 1990). As physiological maturity approaches (from 17 to 21 weeks after sowing, varying between cultivars), the relative number of eggs and anhydrobiotes in the pod and seed tissues increases (Venter et al., 1995). Post harvest, the nematode is able to survive in planting seed, which may be symptomless, and in the field soil in the absence of host plants, or in hulls buried in the soil, for at least 28 to 32 weeks (Basson et al., 1993). This is long enough to survive the dry winter season in South Africa. With the first spring rains the eggs hatch and anhydrobiotes rehydrate. Basson et al. (1993) showed that rehydrated soil populations of D. africanus are able to invade the next crop of groundnut and cause damage. Although relatively few nematodes survive in whole stored seed at 10°C, the surviving nematode population is also able to build up to levels capable of causing damage to the next crop (Basson et al., 1993). The relative importance of the seed and soil inoculum, in the infection of the next crop, has not been determined.
In South Africa, there are a number of nematicides registered for soil application. However, there are no nematicides registered as seed treatments against D. africanus. Attempts to develop nematicide seed treatments against D. africanus have shown that dosages (and microwave treatments) sufficient to kill the nematodes, also destroy the germination potential of groundnut seed.
Seed Health Tests
Soaking Method (Bolton et al., 1990)
1. A subsample of groundnut seeds is taken and the seed cut open.
2. The cut seeds are soaked in tap water for 24 hours at approximately 22°C.
3. The nematodes in the water are poured off and counted.
Notes on testing methods
The efficiency of the soaking method was significantly higher (2 x for hulls; 3 x for seeds) and more consistent (as expressed by the coefficient of variation) than the efficiency of the centrifugal flotation method. The soaking method is also an inexpensive and rapid method involving few steps. The absence of any sieving during the soaking method may have reduced the loss of larvae compared with the centrifugal-flotation method. The recovery of immobile adults by the soaking method indicates that the nematodes not only actively moved out of the tissues but that they were passively released by swelling and bursting of the tissues (Bolton et al., 1990). The total number of nematodes recovered by soaking after 14 days appears to be a good estimation of the total number of all life stages, including eggs, present inside the tissues at the beginning of the soaking period. Soaking for 24 hours (x) gives a reliable estimate of this number (y): y = 37,415 + 1132 x (r = 0.911; P = 0.05) for hulls, and y = 48,663 + 1411 x (r = 0.827; P = 0.05) for seeds (Bolton et al., 1990).
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||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Fruits (inc. pods)||adults; eggs; juveniles||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Roots||adults; eggs; juveniles||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|True seeds (inc. grain)||adults; eggs; juveniles||Yes||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|
|Growing medium accompanying plants|
|Stems (above ground)/Shoots/Trunks/Branches|
ImpactTop of page Although D. africanus is suspected of being present in many southern African countries, it has only been reported to cause damage in South Africa. In this country it is found in all the major groundnut production areas, on about 75% of all fields (De Waele et al., 1989).
It is a pest of the pods of groundnuts on which it can multiply to heavy infections (100,000 nematodes/pod), causing 100% losses in some fields (C Venter, ARC-Grain Crops Institute, Potchefstroom 2520, South Africa, personal communication, 1997. Present address: Green Circle Crop Protection, 14 Krom Street, Potchefstroom 2531, SA). It is found only in low numbers on the roots of groundnut or alternate crops (Basson et al., 1990), and causes no damage to the tubers of potatoes (De Waele et al., 1991), which is the host of the closely related D. destructor.
In heavy infections (final density in excess of 700 nematodes/seed) the seed mass may be reduced by up to 50%, and untimely germination of seedlings may reduce the number of harvestable seeds by up to 25% (Venter et al., 1991).
The greatest economic damage is, however, the increase in the percentage of seeds which are blemished (discoloured) and/or unsound (germinating seeds within closed pods; hypocotyl 1-2 mm) (Venter et al., 1991). This is a major factor in determining the grade of the yield. South African grading regulations require that consignments of groundnut seed containing more than 10 and 20% damaged seed be downgraded from choice edible to standard edible and to crushing grade, respectively, resulting in price decreases of 15 and 65%, respectively. In the glasshouse, a population of 50 D. africanus in the rhizosphere of a groundnut seedling at the beginning of the season, or 20 D. africanus per harvested seed, were able to cause this damage (Venter et al., 1991). In the field, the average number of nematodes per harvested seed in infected samples, from all major groundnut production areas, was 160 (De Waele et al., 1989).
DiagnosisTop of page Although PCR was also used by Wendt et al. (1993) to distinguish Ditylenchus dipsaci from D. destructor and Ditylenchus myceliophagus, it is only one of several tools required for identification of these species. A description of morphology and preferably also of host plants and ecology would be required to complete the species identification.
Nematode DNA was extracted from D. africanus, D. destructor from the UK and the USA (Wisconsin), D. myceliophagus and D. dipsaci, and samples were used for PCR amplifications of the ITS (internal transcribed spacer) region of ribosomal DNA. DNA fragments were size fractionated by electrophoresis, and fragment sizes and coefficients of dissimilarity were calculated.
The ITS region of D. africanus was 1.0 Kb, while the Wisconsin and UK isolates of D. destructor showed an ITS fragment of 1.2 Kb. The same ITS fragment was 0.9 Kb in both D. myceliophagus and D. dipsaci. The number and size of DNA restriction fragments generated by restriction of the ITS region of the Wisconsin and UK isolates of D. destructor were identical using seven restriction enzymes, but different from those of D. africanus. The number and size of restriction bands of D. myceliophagus, D. dipsaci and D. africanus were also different. The proportion of bands shared by the species were small, and the coefficients of dissimilarity were quite high (Wendt et al., 1995).
The size (and probably structure) of the ITS region, amplified between the primer's sequences in the PCR reaction, was different for the three plant parasitic species examined (D. dipsaci, D. destructor and D. africanus), but the two populations of D. destructor had an ITS region of the same size. Almost all restriction sites in the ITS of D. africanus were unique and were not found in the other species examined (Wendt et al., 1995).
Although the sample examined was small, the RFLP data presented leaves no doubt that D. africanus is well separated from D. myceliophagus, D. dipsaci and D. destructor, and that it should not be considered a race or a sibling of any of the species (Wendt et al., 1995).
Detection and InspectionTop of page D. africanus is extremely difficult to extract from soil, or from the roots of any crops.
The quickest detection, in the field, is by inspection of the mature pods for the characteristic grey discolouration which begins at the peg connection and develops in broad bands down one or both longitudinal veins, until the entire pod surface is discoloured (Venter et al., 1991). The inside of the shell is also discoloured and the seed testae flaccid and darkly discoloured.
If in doubt, the shells and seeds can be soaked in tap water for 24 hours (Bolton et al., 1990) and the water inspected for the presence of D. africanus.
Stored seeds release far fewer nematodes when soaked. These are most often dead, and their diagnostic features are in a deteriorated condition (C Venter, ARC-Grain Crops Institute, Potchefstroom 2520, South Africa, personal communication, 1997. Present address: Green Circle Crop Protection, 14 Krom Street, Potchefstroom 2531, SA).
Similarities to Other Species/ConditionsTop of page The histopathology of D. africanus resembles that of Aphelenchoides arachidis, the groundnut testa nematode, which has been found in groundnut seeds in Nigeria (Bos, 1977; Bridge et al., 1977).
In South Africa, groundnut producers often confuse the symptoms of D. africanus with those of the fungus Chalara elegans. Symptoms are distinguished by the pattern of development of pod discolouration. That of D. africanus begins at the peg connection, develops in broad bands down one or both lateral veins, before covering the entire pod surface (Venter et al., 1991). It causes a grey discolouration of the mesocarp, which cannot be removed by scratching off the exocarp of the pod, and is visible on the inside of the shell. That of C. elegans begins at any and various points on the pod, is dark black, and can be scratched off with the exocarp. The seeds of pods infected by D. africanus have yellow to brown flaccid testae, often with darkened veins, and discoloured embryos; while those infected by C. elegans are not discoloured by the fungus.
D. africanus is very similar to Ditylenchus myceliophagous with respect to the number of lines in the lateral field, shape of the tail terminus, c', c, stylet length, length of the post-uterine sac expressed in vulval body diameters, V-value, and spicule and bursa length. However, it differs significantly from D. myceliophagous in its molecular character and host specificity (Wendt et al., 1995).
D. africanus is also very similar to D. destructor (Wendt et al., 1995). De Waele et al. (1989) first considered it to be a race of D. destructor with a limited host range.
These three species can only be clearly differentiated using sensitive RFLP analysis (Wendt et al., 1995).
Prevention and ControlTop of page
Cultural Control and Sanitary Methods
A range of weeds (De Waele et al., 1990), volunteer plants of groundnut and a range of rotation crops (Basson et al., 1990) and fungi, host the peanut pod nematode. General weed, volunteer plant and fungal control is therefore important in controlling the pre-plant build-up of the nematode. Glasshouse trials have shown that a pre-plant population of only 50 nematodes per seedling can lead to a downgraded yield (Venter et al., 1991).
The survival mechanisms of D. africanus are initiated with ripening of the pods. Increased numbers of eggs and anhydrobiotes are found with delayed harvest (Venter et al., 1995). Timely harvesting will allow drying of the hulls and seeds before eggs are laid en mass, and such hull stubble and seed for planting will therefore contain lower numbers of survival stages of the nematode (Venter, 1994).
Many producers delay harvesting to allow maximum seed fill, particularly where symptoms of nematode infection are not prominent (Venter et al., 1992a). The danger then exists that the hull stubble will be a source of re-infestation of the soil. This waste should be removed from the field, where possible.
One of the key strategies in the control of seedborne disease must be the production or selection of high quality seed for planting. Trials in a glasshouse have shown that planting seed with only 20 nematodes per seed can give a downgraded yield (Venter et al., 1991).
The cultivar Sellie and the other cultivars available when the peanut pod nematode was discovered, are very susceptible to the nematode (Venter et al., 1993). Since then, the partially tolerant Kwarts has been released for multiplication and commercial use. Other groundnut lines showing greater promise of resistance (Van der Merwe and Joubert, 1994) are currently being developed for release.
Producers in the Northern Cape Province of South Africa, who have grown groundnuts in monoculture for 10 to 30 years, claim that the peanut pod nematode has spontaneously disappeared and is no longer a problem in these fields (Venter, 1994). With the hope that these soils had become suppressive as a result of the build up of nematophagous fungi, a survey was carried out to isolate these fungi from these and other soils.
A variety of sixteen fungi were identified (Jones et al., 1996), although they were not specifically related to a suppressive soil. Eight of these species could be cultured, and could therefore be studied for their potential as commercial bio-control agents. Arthrobotrys spp. were the most common, trapping nematodes in two or three dimensional traps. Monacrosporium spp. appeared to be very aggressive, using a three dimensional trapping network, and were also isolated in association with nematodes extracted from within the groundnut hulls, indicating that this fungus is also capable of spreading into the groundnut pod. A procedure for applying the fungi to the soil in pellet form was developed, but trials showed that these fungi were unfortunately not the major factor in causing suppressiveness in soils, and that their use in biological control is not effective and economic (WJ Jooste, ARC-Grain Crops Institute, Potchefstroom 2520, South Africa, personal communication, 1997).
ReferencesTop of page
Basson S; De Waele D; Meyer AJ, 1990. An evaluation of crop plants as hosts for Ditylenchus destructor isolated from peanut. Nematropica, 20:95-101.
Basson S; De Waele D; Meyer AJ, 1993. Survival of Ditylenchus destructor in soil, hulls and seeds of groundnut. Fundamental and Applied Nematology, 16:79-85.
Bolton C; De Waele D; Basson S, 1990. Comparison of two methods for extracting Ditylenchus destructor from hulls and seeds of groundnut. Revue de Nematologie, 13:233-235.
Bridge J; Bos WS; Page LJ; McDonald D, 1977. The biology and possible importance of Aphelenchoides arachidis, a seed-borne endoparasitic nematode of groundnuts from northern Nigeria. Nematologica, 23(2):253-259
De Waele D; Jones BL; Bolton C; Van den Berg E, 1989. Ditylenchus destructor in hulls and seeds of peanut. Journal of Nematology, 21:10-15.
De Waele D; Jordaan EM; Basson S, 1990. Host status of seven weed species and their effects on Ditylenchus destructor infestation on peanut. Journal of Nematology, 22:292-296.
Van der Merwe PJA; Joubert HLN, 1994. Tolerance of groundnut cultivars to the potato-rot nematode (Ditylenchus destructor). In: Proceedings of the 10th South African Maize Breeding Symposium. Technical Communication No. 238. South Africa: Department of Agriculture, 86-89.
Venter C, 1994. Strategies for the control of the peanut pod nematode on groundnuts in South Africa. In: Proceedings of the 11th Regional Groundnut Workshop for Southern Africa, ICRISAT, Mbabane, Swaziland, July 1994.
Venter C; De Waele D; Meyer AJ, 1991. Reproductive and damage potential of Ditylenchus destructor on peanut. Journal of Nematology, 23:12-19.
Venter C; De Waele D; Meyer AJ, 1992a. Minimizing damage by Ditylenchus destructor to peanut seed with early harvest. Journal of Nematology, 24:529-532.
Venter C; De Waele D; Meyer AJ, 1993. Reproductive and damage potential of Ditylenchus destructor on six peanut cultivars. Journal of Nematology, 25:59-62.
Venter C; De Waele D; van Eeden CF, 1992. Plant-parasitic nematodes on field crops in South Africa. 4. Groundnut. Fundamental and Applied Nematology, 15:7-14.
Wendt KR, 1992. Taxonomic differentiation of three species of Ditylenchus (Nematoda) and their races using ribosomal DNA analysis. M. Sc. Thesis. Burnaby, Canada: Simon Fraser University.
Wendt KR; Swart A; Vrain TC; Webster JM, 1995. Ditylenchus africanus sp. n. from South Africa; a morphological and molecular characterization. Fundamental and Applied Nematology, 18(3):241-250; 12 ref.
Wendt KR; Vrain TC; Webster JM, 1994. Separation of three species of Ditylenchus and some host races of D. dipsaci by restriction fragment length polymorphism. Journal of Nematology, 25(4):555-563; 29 ref.
Zhang XueSong; Zhang GuoZhen; Zhang HaiWang; Peng DongWen; Du ZhiJian; Tang ChunYan, 2009. Effects of soil treatments on the quantitative dynamics of rhizospheric nematodes and their control efficacy to root rot of American ginseng caused by Ditylenchus destructor. Acta Phytopathologica Sinica, 39(5):555-560. http://zwblxb.periodicals.net.cn/default.html
Distribution MapsTop of page
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