Rotylenchulus reniformis




Rev 02/22/2022

Reniform Nematode Classification Hosts
Morphology and Anatomy Life Cycle
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           Rotylenchulus reniformis Linford and Oliveira, 1940

Reniform nematode

Linford and Oliviera established the genus Rotylenchulus in 1940 with R. reniformis as the type species. The generic name was given by Linford and Oliviera because they thought that the nematode species was similar to the genus Rotylenchus, having features of that genus and other Hoplolaimidae. The species name was coined because of the kidney shape of the mature female.

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Morphology and Anatomy:

Drawing by Charles S. Papp, CDFA

Immature Female: 

  • Body vermiform, slender and spiral to C-shaped when heat-killed.

  • Length about 0.4 mm.

  • Stylet knobs are rounded and slope posteriorly.

  • DEGO  distant from base of stylet knobs, perhaps one stylet length or more.

  • The median bulb of the esophagus has a distinct valve and the basal glands of esophagus overlap the intestine laterally and ventrally. 

  • The vulva is not prominent; it is located at about 70% of the body length. 

  • Ovaries paired and opposed with double flexure. . 

  • Tail tapers to a narrow rounded terminus.

Mature Female: 

  • Body swollen, kidney-shaped, with an irregularly neck, 0.38-0.52 mm long. 

  • The vulva has raised lips. 

  • The body beyond the anus is hemispherical, with a slender terminal portion 5-9 µm long. 

  • Well-developed stylet.

  • Cuticle thick.  

  • Ovaries very long, convoluted; vulva post-equatorial. 

  • Eggs deposited in a gelatinous matrix. 

Rotylenchulus reniformis: immature and mature female


  • Vermiform.  Anterior end reduced; stylet reduced. 

  • The esophagus is degenerate with reduced median bulb and valve. 

  • Males do not feed.

  • The spicules are elongate-slender, ventrally curved.

  • Caudal alae present but difficult to see, not quite reaching tail end. 

Juveniles and males remain in soil.


  Reported median body size for this species (Length mm; width micrometers; weight micrograms) - Click:


Rotylenchulus reniformis:
female and male head; male tail

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Rotylenchulus reniformis is widely distributed in many tropical and subtropical regions of the world. It has been reported in tropical and sub-tropical West and Central Africa; Central and South America; Southeast Asia, the Carribean, Mexico, Japan, the Middle East, South Pacific, Italy, Spain, China and the Far East. 

Within the United States the reniform nematode is known to be established in Alabama, Arkansas, Florida, Georgia, Hawaii, Louisiana, Mississippi, North Carolina, South Carolina, and Texas. Its reported pattern of distribution suggests that it is likely to be present in southwestern Tennessee and possibly Oklahoma.

In Hawaii on cowpea; also in the Southeastern U.S., and in Texas.

In California, R. reniformis infested Phoenix roeselenii and Cycas sp. plants were detected in San Diego in 1960, having entered the state in a quarantine shipment. The plants had been established in a residential property before a confirmed diagnosis of the pest had been completed. Subsequently, the plants were removed from the infested site and fumigated with methyl bromide. The planting site was also fumigated with methyl bromide.

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Economic Importance:

 A-rated pest in California Nematode Pest Rating System.

Crop stunting due to Rotylenchulus reniformis damage.  Photo by Charles Overstreet.


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Although not closely related (taxonomically) to citrus nematode (Tylenchulus semipenetrans), R. reniformis does have some similarities to it in terms of feeding habits.

Nurse cells form near pericycle - 100-200 per female in soybean.  Nurse cell system is stimulated by feeding which causes hypertrophy of pericycle and endodermis cells, increased cytoplasm density, but cells remain uninucleate with large nucleolus.  Walls may rupture to form a syncytium.  Syncytium about 2 cells deep may extend half way around root in soybeans.  Syncytia are stimulated primarily in pericycle tissues (phase 1: cell wall lysis; phase 2: anabolic phase - increase in organelles of affected cells).


The nematode also feeds on cortical cells of cowpea and phloem of cotton.

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Many species of cultivated plants and fruit trees, cotton, cowpea, tea, and soybean; pineapple in Hawaii.

The reniform nematode attacks over 140 species of more than 115 plant genera in 46 families (Jatala, 1991). 

Some of the economically important host plants are: banana, cabbage, cantaloupe, cassava, citrus, kale, lettuce, mango, okra, pigeon pea, pineapple, pumpkin, coconut, cotton, radish, cowpea, soybean, sweet potato, crimson clover, tobacco, eggplant, tomato and guava.

Sugarcane was recommended as rotation crop based on field observations indicating that the nematode is not present.  However, the rotation was rarely used when effective nematicides were readily available.  With reduction in nematicide use, there have been reports of damage to pineapple following sugarcane.  

Rotylenchulus reniformis is a pest of sugarcane in West Africa and of cotton in Louisiana.

For an extensive host range list for this species, click

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Life Cycle:

Ecophysiological Parameters:

For Ecophysiological Parameters for this species, click If species level data are not available, click for genus level parameters


Rotylenchulus reniformis is a sedentary, semi-endoparasite in the mature female stage. The reniform nematode reproduces sexually. It may also reproduce parthenogenetically.

Reniform nematode females and egg masses on a cotton root.  Photo by Charles Overstreet. Life cycle of Rotylenchulus reniformis

Graphic by Charles Overstreet.


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In Louisiana, R. reniformis causes a 40-60% reduction in cotton yield, with a comcomitant increase in Fusarium wilt.  

In the presence of this nematode, Fusarium wilt-resistant varieties of cotton also become susceptible.

Above ground symptoms on host plants include dwarfing, shedding of leaves, formation of malformed fruit and seeds, and general symptoms of an impaired root system. Below ground, roots are discolored and necrotic (dead) with areas of decay. Plant mortality is possible in heavy infestations.

In banaba and plantain (Musa spp.) symptoms and damage attributed to R. reniformis include necrosis and reduction of secondary root development, stunting, chlorosis of aboveground vegetation, and restricted development and reduced yield of banana and plantain. Significant yield losses of between 25 and 60% have been recorded with population levels of 0.1 to 10 R. reniformis cm3 of soil (Riascos-Ortiz et al., 2019).

In experiments in Hawaii, pineapple fruit weight increased with increasing population densities up to 300–310 nematodes/250 cm3 soil but decreased at 1020–1360 nematodes/250 cm3.  Preplant populations of R. reniformis below 300 nematodes/250 cm3 soil damage pineapple but are not the major factor limiting yield. Yield losses at these population levels may be offset by managing soil fertility and physical soil factors. R. reniformis becomes the major yield-limiting factor at population densities above 1000 nematodes/250 cm3 soil (Sipes and Schmitt, 2000).

Rotylenchulus reniformis damage to pineapple roots (left) in pot tests.
Root necrosis around the head region of R. reniformis females in banana roots.



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Rotylenchulus reniformis is a Class A pest in California.  

The CDFA Nematode Study Committee recommended the adoption of a reniform nematode quarantine  independent of any other nematode quarantine measures. California Department of Food and Agriculture's Reniform Nematode Exterior Quarantine Program was established in 1997 in order to continue to prevent the introduction of this nematode species through infested plant and associated materials in out-of-state shipments to California. Similar to the burrowing nematode quarantine program, a secondary screening mechanism exists in the nursery certification program.

A. Exclusion

The CDFA Nematology Laboratory made 13 detections in 1989, 9 in 1990, 6 in 1991, 2 in 1992, 5 in 1993, 2 in 1994, 4 in 1995, 8 in 1996, 6 in 1997 and 9 in 1998.

B.  Eradication

Infested Phoenix roeselenii and Cycas sp. plants detected in San Diego in 1960 were established in a residential property before a confirmed diagnosis of the pest had been completed. Subsequently, the plants were removed from the infested site and fumigated with methyl bromide. The planting site was also fumigated with methyl bromide.

Infestations of R. reniformis on established Yucca gloriosa plants were first detected in 13 residential properties in Highland, San Bernardino County during a residential grid survey in 1967. The infestation was traced to yuccas brought into California from Harlingen, Texas and planted in the subdivision. The infested areas were treated with Nemagon (DBCP). In 1971, the nematode was detected again in the same locality. Despite a second treatment of Nemagon, new infestations of the nematode appeared in 1973 and 1974. Subsequent herbicide and fumigation trials were conducted, and on December 31,1978, R. reniformis was officially declared eradicated from the infested areas in San Bernardino County. 

In 1980, the nematode was detected again from areas found free of the nematode in the 1970's. The current status of the San Bernardino infestation is not known (Chitambar, 1997).

C.  Management


1,3-Dichloropropene (1,3-D) (Telone) (8 gal/acre)

 Temik (6 lb/acre) on cotton and pineapple.

Studies on the effects of the reniform nematode on yields of various vegetable crops, grown in the Rio Grande Valley in Texas, have shown that soil fumigation prior to planting significantly increased yields in reniform nematode infested fields (Robinson et al., 1987). Soil fumigation with dichloropropene (Telone) type fumigants significantly increased crop yields for cotton, tomato, lettuce, and soybean. In one experiment 1,3-dichloropropene plus aldicarb, when compared with fenamiphos, phorate, terbufos, and aldicarb, provided the greatest protection. In another, aldicarb, carbofuran, and phorate were effective against the reniform nematode infesting Thompson seedless grapevines, with aldicarb providing the maximum yield increase. 

Repeat foliar sprays of oxamyl were the most effective against reniform nematodes infesting tomato and cotton (Rich and Bird, 1973). Methyl bromide will be unavailable as of 2005. The availability of the other pesticides listed will depend upon their registration status when needed.

Soil Solarization - Soil solarization in Egypt controlled the reniform nematode for 60 days after planting; it improved plant growth and increased yields by 25 to 40 percent in broad beans, onions, tomatoes, and clover in various types of soils.

Crop Rotation - Rotation of soybeans with corn, sorghum, and wheat reduced populations of reniform nematode. Plantings of poor host species can reduce the reniform nematode numbers in soil more effectively than fallow alone. 

Reniform nematode is a significant problem on both cotton and soybean in Mississippi.  Resistant varieties of soybean are available, but not of cotton. Soybeans are usually rotated with cotton, but that rotation is a problem when reniform nematode is present.  In that case, nematicides are used prior to the cotton crop.  For soybeans, a 1 yeasr rotation to a non-host crop is effective (Coblentz, 2005).

Some non-hosts reported as good rotational crops for cotton include: sorghum (Heald, 1974), maize (Braithwaite, 1974), two years with reniform resistant soybeans (Gilman et al., 1978), and sugarcane and Pangolagrass (Heald and Thames, 1982).

Sugarcane has been recommended as a rotation crop for pineapple in Puerto Rico and Hawaii, but may be based on poor information.  Rotation not effective with Hawaiian sugarcane (it may be a host, but additional research in needed).

Plant Resistance

 A few reports of plant resistance have been documented in Gossypium spp. (controlled by 2 or more pair of genes) and tomato in Egypt and India (Oteifa and Osman, 1974). The following plants have been reported as showing immunity or resistance to the reniform nematode: barley, hot pepper, barnyard grass, sweet pepper, sweet sorghum, pangola grass, spinach, mustard, sugarcane and oats (Armstrong and Jensen, 1978; Bridge, 1983; Inserra et al., 1983).

Soybean varieties - Peking, Dyer, Custer, Pickett - have a hypersensitive response to nematode, but there is no increase in metabolic activity of cells.

Host Plant Resistance, Non-hosts and Crop Rotation alternatives:

For plants reported to have some level of resistance to this species, click


Break anhydrobiotic survival stage by pre-irrigation 3 months before pineapple - increase decline (Caswell, Apt).

Soil Amendments - Animal manures and cotton seed cakes have been used (Badra et al., 1979). 

Neem, castor leaves, cowpea and okra have proven to be as affective as carbofuran granules in India (Rao et al., 1996). 

Powders of sour orange peel, stored garlic cloves and tobacco leaves gave 84-92% female nematode reduction (Amin and Youssef, 1998). 

Soil application of certain Tagetes spp. plant wastes also controlled the reniform nematode.


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Fortuner, 1987.  Rev. Nematol. 10:219-232

H. Ferris

Linford and Oliveira (1940)

Amin, A.W. and M.M.A. Youssef. 1998. Effect of organic amendments on the parasitism of Meloidogyne javanica and Rotylenchulus reniformis and growth of sunflower. Pakistan Journal of Nematology. 16(1):63-70.

Armstrong J.M. and H.J. Jensen. 1978. Bulletin 639, Indexed bibliography of nematode-resistance in plant. Corvallis, OR: Agricultural Experiment Station.

Badra, T., M.A. Salem and B.A. Oteifa. 1979. Nematicidal activity of some organic fertilizers and soil amendments. Revue de Nematologie. 2:29-36.

Barker, K.S., S.R. Koenning and S.A. Walters. 1994. Effects of soil type on the reproductive potential of Meloidogyne incognita and Rotylenchulus reniformis on cotton and related effects on crop maturity. Journal of Nematology. 26(1): 91-92.

Birchfield, W, and J. E. Jones. 1961. Distribution of the reniform nematode in relation to crop failure of cotton in Louisiana. Plant Disease Reporter. 45:671-673.

Birchfield, W., and W. J. Martin. 1967. Reniform nematode survival in air-dried soil. Phytopathology. 57:804.

Bridge, J. 1983. Nematodes. Pp 69-84, Pest control in tropical tomatoes. London: Centre for overseas Pest Research, Overseas Development Administration.

Chitambar, J.J. 1997. A brief review of the reniform nematode, Rotylenchulus reniformis. California Plant Pest and Disease Report, CDFA. 16:71-73.

Coblentz, B. 2005. MSU research battles nematodes and weeds.  Mississippi State University Agricultural News, October 13, 2005.

Dasgupta, D. R., and A. R. Seshadri. 1971. Races of the reniform nematode Rotylenchultis reniformis Linford and Oliviera, 1940. Indian Journal of Nematology. 1:21-24.

Fortuner, R. 1987. A reappraisal of Tylenchina (Nemata). The family Hoplolaimidae Filip'ev 1934. Revue de Nematologie. 10:219-232.

Gilman, D.G., J.E. Jones, C. Williams and W. Birchfield. 1978. Cotton-soybean rotation for control of reniform nematodes. Louisiana Agriculture. 21:10-11.

Heald, C.M. and W.H. Thames. 1982. The reniform nematode, Rotylenchulus reniformis. Pp. 139-143, R.D. Riggs et al., Southern Regional Research Committees S-76 and S-154 (eds.), Nematology in the Southern Region of the United States. Southern Cooperative Series Bulletin.

Jatala, P. 1991. Reniform and false root-knot nematodes, Rotylenchulus and Nacobbus spp. Pp. 1035, W.R. Nickle (ed.), Manual of Agricultural Nematology. New York: Marcel Dekker, Inc.

Jensen, H. J. 1972. Nematode pests of vegetable and related crops. Pp. 377-408, J. M. Webster, (ed.), Economic Nematology. New York: Academic Press.

Nakosono, K. 1966. Role of males in reproduction of the reniform nematodes, Rotylenchulus spp. (Tylenchida: Hoplolaimidae). Appl. Ent. Zool. 1:203-205.

Naqvi S.Q.A. and M.M. Alam. 1975. Influence of brinjal mosaic virus in the populations of Tylenchorhynchus brassicae and Rotylenchulus reniformis around eggplant roots. Geobios. 2:120-121.

Oteifa, B.A. and A.A. Osman. 1974. Host-parasite relations of Rotylenchulus reniformis on Solanum lycopersicum. Pp. 78-79, Simposia Internacional (XII) de Nematologia. Sociedad Europe de Nematologos, Septiembre. Granada, Spain.

Rebois, R.V. 1973. Effect of soil temperature on infectivity and development of Rotylenchulus reniformis on resistant and susceptible soybeans, Glycine max. Journal of Nematolology. 5:10-13.

Riascos-Ortiz, E., A.T. Mosquera-Espinosa, F.V. De Agudelo, C.M. Gon�alves de Oliveira and J.E. Mu�oz-Fl�rez. 2019. Morpho-molecular characterization of Colombian and Brazilian populations of Rotylenchulus associated with Musa spp. J. Nematology 51: DOI: 10.21307/jofnem-2019-047

Rich, J.R. and G.W. Bird. 1973. Inhibition of Rotylenchulus reniformis penetration of tomato and cotton roots with foliar applications of oxamyl. Journal of Nematology. 5:221-224.

Robinson, A. F., C. M. Heald, S. L. Flanagan, W. H. Thames and J.Amador. 1987. Geographical distribution of Rotylenchulus reniformis, Meloidogyne incognita, and Tylenchulus semipenetrans in the lower Rio Grande valley as related to soil texture and land use. Annals of Applied Nematology. 1:20-25.

Sasser, J. N. 1972. Nematode diseases of cotton. Pp. 197-214, J. M. Webster (ed.), Economic Nematology. New York: Academic Press.

Singh, R.V. and S. Khera. 1979. Pathogenicity of Rotylenchulus reniformis on brinjal (Solanum melongena L.). Indian Journal of Nematology. 9:117-124.

Sipes, B.S. and Schmitt, D.P. 2000. Rotylenchulus reniformis damage thresholds on pineapple. Acta Hort.  529:239-246


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Copyright © 1999 by Howard Ferris.
Revised: February 22, 2022.