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Datasheet

Rhadinaphelenchus cocophilus
(red ring nematode)

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Datasheet

Rhadinaphelenchus cocophilus (red ring nematode)

Summary

  • Last modified
  • 15 July 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Natural Enemy
  • Preferred Scientific Name
  • Rhadinaphelenchus cocophilus
  • Preferred Common Name
  • red ring nematode
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Nematoda
  •       Order: Aphelenchida
  •         Family: Aphelenchoididae

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Pictures

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PictureTitleCaptionCopyright
Left: Palm tree affected by red ring disease. Right: Palm trunk showing characteristic red ring.
TitleSymptoms
CaptionLeft: Palm tree affected by red ring disease. Right: Palm trunk showing characteristic red ring.
Copyright©Sanchez
Left: Palm tree affected by red ring disease. Right: Palm trunk showing characteristic red ring.
SymptomsLeft: Palm tree affected by red ring disease. Right: Palm trunk showing characteristic red ring.©Sanchez

Identity

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Preferred Scientific Name

  • Rhadinaphelenchus cocophilus (Cobb, 1919) Goodey, 1960

Preferred Common Name

  • red ring nematode

Other Scientific Names

  • Aphelenchoides cocophilus (Cobb, 1919) Goodey, 1933
  • Aphelenchus cocophilus Cobb, 1919
  • Bursaphelenchus cocophilus (Cobb, 1919) Baujard, 1989
  • Chitinoaphelenchus cocophilus (Cobb, 1919) Chitwood in Corbett, 1959

International Common Names

  • Spanish: anillo rojo; nematodo del anillo rojo

Local Common Names

  • Brazil: anel vermelho

EPPO code

  • RHAACO (Bursaphelenchus cocophilus)

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Nematoda
  •             Order: Aphelenchida
  •                 Family: Aphelenchoididae
  •                     Genus: Rhadinaphelenchus
  •                         Species: Rhadinaphelenchus cocophilus

Notes on Taxonomy and Nomenclature

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The generic placement of this species requires confirmation as some authors regard it as belonging to the monotypic genus Rhadinaphelenchus and others to the genus Bursaphelenchus. Although cocophilus is undeniably close to other species in the genus Bursaphelenchus (and may well prove to belong to that genus), there remains some doubt as to which genus it should belong to. Accordingly, being a species of economic importance with an established literature, it is perhaps more prudent to leave it in Rhadinaphelenchus until a wider consensus is achieved.

Description

Top of page The following morphological description is after CIH (1975).

Female

Body about 1 mm long and very slender (a=60-96), arcuate to nearly straight when relaxed; cuticle thin, marked with transverse striae 0.6-1 µm apart; lateral fields with four incisures and a faint median line suggesting a fifth incisure, occupying 0.25 of body-width (Goodey, 1960); deirids and phasmids absent. Lip region smooth, high, anteriorly flattened with rather straight sides, slightly narrower than body; head frame-work prominent, sclerotized. Spear 11-13 µm long, attenuated knobbed at base but knobs may be obscure, especially on immature specimens (Thorne, 1961); anterior part less than half spear length and sharply pointed; protrudor muscles of spear prominent, attached to basal plate of labial frame-work. Procorpus elongate-cylindrical; metacorpus or median bulb oval, usually about twice as long as wide, with prominent valve-plates just posterior to centre; dorsal oesophageal gland orifice midway between anterior margin of bulb and valve plates. Oespohageal glands overlapping intestine dorsally, usually obscure. Nerve ring a wide band surrounding isthmus about 0.5-1 bulb-length behind the bulb; excretory pore a little behind nerve ring and anterior to hemizonid which is about 3 annules long. Intestine with small granules and indistinct lumen. Vulva slit-like appearing as an open C in ventral view, slightly over-hung by a wide, thick dorsal lip; posterior lip is also thick and heavily sclerotized. Esser says in litt. that a vulval flap as illustrated by Goodey (1960) may or not be present. Of 12 females he examined, one possibly possessed a vulval flap. Vagina thick-walled, slightly curved as it leads inwards to a distance of about 0.5 of body-width. Anterior gonad well developed, outstretched; oocytes in a row. Postvulval uterine sac elongate, extending to about 0.75 of vulva-anus distance, often with a few large spheroid sperms. Rectum about 1.5 anal body-widths long; anus distinct. Tail elongate-subcylindrical with a rounded, unstriated terminus, 10-17 anal body-widths long.

Male

Body ventrally arcuate, more strongly curved in tail region. Head, spear and oespohagus as in female. Testis single, anteriorly outstretched, with spermatogonia in a row. Spicules paired, small; dorsal limb 9-11 µm long with an elongated rounded apex and ending distally before the ventral limb whose distal end appears to recurve to join the dorsal limb so that the entire spicule appears notched distally; the ventral element has a distinct rostrum proximally and appears to be connected to the dorsal limb through a transverse bar with a central hole for passage of nerves. No gubernaculum, but dorsal wall of spicule pouch is thickened to form an apophysis. Tail strongly curved ventrally (may form 1.5 circles), subcylindrical in anterior half, then conoid to a pointed terminus. Bursa (or caudal alae) terminal, prominent in dorsal or ventral view (not easily detectable in lateral view as it does not project beyond tail contour) with finely striated margins, enveloping distal 0.4-0.5 of tail. There are two pairs of distinct ventro-submedian papillae near base of bursa and a pre-anal pair, about 0.5 of the spicule length anterior to cloaca; a fourth pair of ventro-lateral papillae behind the cloaca has been reported by Thorne (1961) and Nickle (1970) but these are very obscure and often not detectable.

Larvae

Larvae have high, dome-shaped heads, not offset from body. Tails of second- and third-stage larvae have conoid or sharply mucronate tips, and those of fourth-stage larvae have dimorphic tips: in female larvae they are rounded as in the female, and in male larvae sharply drawn out.

Distribution

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At present, R. cocophilus has a restricted distribution and has only been reported from the West Indies and from Latin America. R. cocophilus does not occur in the northern Caribbean islands, Florida, Cuba or other parts of the world outside the Western Hemisphere (Dean, 1979).

Distribution Table

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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/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

North America

MexicoWidespreadAlcocer, 1955; CABI and EPPO, 1999; EPPO, 2014

Central America and Caribbean

BahamasAbsent, unreliable recordCABI and EPPO, 1999; EPPO, 2014
BarbadosAbsent, reported but not confirmedCABI and EPPO, 1999
BelizePresentCABI/Anon., 1954; EPPO, 2014
Costa RicaWidespreadTidman, 1951; CABI and EPPO, 1999; EPPO, 2014
Dominican RepublicAbsent, unreliable recordCABI and EPPO, 1999; EPPO, 2014
El SalvadorWidespreadBrathwaite and Siddiqi, 1975; CABI and EPPO, 1999; EPPO, 2014
GrenadaWidespreadCABI/Nowell, 1919; Singh, 1972; EPPO, 2014
GuatemalaWidespreadDean, 1979; CABI and EPPO, 1999; EPPO, 2014
HaitiAbsent, unreliable recordCABI and EPPO, 1999; EPPO, 2014
HondurasWidespreadBrathwaite and Siddiqi, 1975; CABI and EPPO, 1999; EPPO, 2014
JamaicaAbsent, reported but not confirmedCABI and EPPO, 1999
NicaraguaWidespreadDean, 1979; CABI and EPPO, 1999; EPPO, 2014
PanamaWidespreadBriton-Jones, 1940; CABI and EPPO, 1999; EPPO, 2014
Puerto RicoAbsent, unreliable recordCABI and EPPO, 1999; EPPO, 2014
Saint Vincent and the GrenadinesPresentCABI/Brathwaite & Siddiqi, 1975; EPPO, 2014
Trinidad and TobagoPresentCABI/Briton-Jones, 1940; EPPO, 2014

South America

BrazilWidespreadCABI and EPPO, 1999; EPPO, 2014
-AmazonasPresentCABI and EPPO, 1999; EPPO, 2014
-BahiaPresentChaves Batista, 1948; CABI and EPPO, 1999; EPPO, 2014
-CearaPresentPonte et al., 1971; CABI and EPPO, 1999; EPPO, 2014
-ParaPresentChaves Batista, 1948; CABI and EPPO, 1999; EPPO, 2014
-SergipePresentSharma & Loof, 1982; CABI and EPPO, 1999; EPPO, 2014
ColombiaWidespreadCabrera, 1965; CABI and EPPO, 1999; EPPO, 2014
EcuadorPresentBrathwaite and Siddiqi, 1975; CABI and EPPO, 1999; EPPO, 2014
French GuianaPresentCABI and EPPO, 1999; EPPO, 2014
GuyanaPresentCABI/Briton-Jones, 1940; EPPO, 2014
PeruWidespreadLiceras, 1967; CABI and EPPO, 1999; EPPO, 2014
SurinamePresentCABI/Maas, 1969; EPPO, 2014
VenezuelaWidespreadFenwick, 1959; CABI and EPPO, 1999; EPPO, 2014

Risk of Introduction

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R. cocophilus is a potential phytosanitary risk on coconuts in all tropical countries, especially where the palm weevil, Rhynchophorus palmarum, is known to occur. Local dissemination occurs by the weevil vector but wider movement can only occur by the transport of infested coconut and other palm tissues.

Habitat

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R. cocophilus is found in tropical environments, closely associated with palms. It is found mainly in the stems of palm trees, but also in leaf petioles and occasionally in roots. The palm weevil, Rhynchophorus palmarum, is the vector of the nematode and life stages of R. cocophilus can be found in the gut, body cavity and the region of the ovipositor of the weevil.

Hosts/Species Affected

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R. cocophilus is a parasite of palms only.

Growth Stages

Top of page Flowering stage, Fruiting stage, Vegetative growing stage

Symptoms

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Young coconut palms easily succumb to R. cocophilus attack. There is no record of any tree, once affected, having recovered. The disease occurs more commonly in trees 2.5-10 years old, with greatest incidence in those 4-7 years old. Occasionally, a palm as young as 1.5 or as old as 20 years or more may be attacked.

The symptoms described are those for palms of the tall cultivar of coconuts or 'typica' which grow in the West Indian islands. These symptoms differ somewhat in the dwarf variety 'nana' and also some panama talls. Chlorosis first appears at the tips of the oldest leaves and spreads towards their bases but, occasionally, the younger leaves may be affected first. The brown lower leaves may break across the petiole of the lower part of the rachis, or they may become partly dislodged at the base and hang down. Nuts are shed prematurely, either simultaneously with the development of leaf symptoms, or slightly before. The crown often topples over, about 4-6 weeks after symptoms first appear, due to associated severe damage caused internally by the larvae of the palm weevil. However, the trunk remains standing in the field for several months until it decays. At the onset of symptoms, the chlorotic yellow appearance of the leaves around the stem is sometimes indistinguishable from those of trees growing under conditions of poor drainage or during intense drought.

The most characteristic symptoms are the internal lesions. In a cross-section of the stem, they appear as an orange to brick-red coloured ring, 2-4 cm wide, and at a distance of 3-5 cm from the periphery. In longitudinal section, the reddened tissue may appear as two united bands joined in the bole forming a 'U'-shape. Lesions at the upper end of the stem in the vicinity of the crown are discrete, appearing first as streaks and then as dots. The meristematic tissue in the bud remains white and apparently healthy. There is no putrefaction of the bud associated with R. cocophilus attack. In the roots, the normally white soft cortex becomes orange to faint red in colour, and has a dry and flaky texture when diseased. In the leaves, a solid core of mottled tissue, dull red to brown in colour, extends from the leaf-base up to 75 cm in the petioles.

The disease is not recognizable externally in its very early stages. The roots, stems and leaf petioles are already infested and there is full development of internal symptoms before the first external symptoms become visible. In the dwarf cultivars, the red colour gives way to shades of brown. Thus, instead of a red ring internally, there is a brown band. The discrete spots are also brown and the yellow discoloration of the leaves is not often apparent. Generally, the leaves become dried and brown, beginning at the tips of the leaflets and progressing downwards. The yellow dwarf cultivars respond in the same way as the green and the crosses between talls and dwarfs, or between Panama tall and any dwarf. They show a browning instead of a characteristic reddening of the leaves and stem tissue.

The heaviest losses due to R. cocophilus occur at the end of the wet season and in the first 2-3 months of the dry season (December to March) in Trinidad.

List of Symptoms/Signs

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SignLife StagesType
Fruit / premature drop
Leaves / abnormal colours
Leaves / abnormal leaf fall
Leaves / wilting
Leaves / yellowed or dead
Roots / cortex with lesions
Stems / internal red necrosis
Whole plant / plant dead; dieback

Biology and Ecology

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The vector of R. cocophilus is the palm weevil, Rhynchophorus palmarum, and the biology and life cycle of R. cocophilus is intimately associated with the weevil. Weevil size was said to be an indicator of vector status, but recent evidence suggests that this is not the case (Gerber and Giblin-Davis, 1990). Experimentally, it has also been shown that red ring disease can be caused by the nematodes entering the plant through the root system.

Larvae of R. palmarum feed by burrowing through coconut stems and, when this occurs in trees which are infected with R. cocophilus, the larvae can become inoculated with the nematode. Adult weevils emerging from trees infected with R. cocophilus carry the nematode to new sites. R. cocophilus enters the haemocoel of weevil larvae via the gut tract; in adult weevils, R. cocophilus can be found in the gut, body cavity and the region of the ovipositor.

In coconut tissues, R. cocophilus invades parenchyma tissue in the roots, stems and leaves, and also artificially infested nuts. At first, the nematodes occur as intercellular parasites in newly invaded tissue but later they can be found both intercellularly and intracellularly. In many cases, lysigenous cavities are formed in which large numbers of nematodes are found. One gram of such tissue can contain as many of 10 000 nematodes. The heaviest losses due to R. cocophilus occur at the end of the wet season and in the first 2-3 months of the dry season (December to March) in Trinidad. R. cocophilus inoculated into the mesocarp of coconuts have a life cycle, from egg to egg, of 9-10 days.

R. cocophilus infestation occurs more commonly in trees 2.5-10 years old, with greatest incidence in those 4-7 years old. Occasionally, a palm as young as 1.5 or more than 20 years old may be attacked.

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Bark 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
Leaves 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
Wood 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
Bulbs/Tubers/Corms/Rhizomes
Flowers/Inflorescences/Cones/Calyx
Fruits (inc. pods)
True seeds (inc. grain)

Vectors and Intermediate Hosts

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VectorSourceReferenceGroupDistribution
Rhynchophorus palmarumInsect

Impact

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R. cocophilus causes major crop loss of coconut plantations in its restricted area of distribution. It can also seriously damage oil palms. The percentage loss can vary from a few percent to complete destruction of young coconuts. Young coconut palms easily succumb to R. cocophilus attack. There is no record of any tree, once affected, having recovered. The heaviest losses due to R. cocophilus occur at the end of the wet season and in the first 2-3 months of the dry season (December to March) in Trinidad.

Diagnosis

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Recovery of R. cocophilus from coconut tissue

The methods used for recovering nematodes from the palm differ according to the degree of activity of the nematodes in the tissue, and also the density of nematodes per gram of infested tissue. In the method originally used by Fenwick (Fenwick and Maharaj, 1963), diseased coconut tissue is chopped into fine pieces about 1 cm thick, placed in a large funnel of water, whose stem is closed at one end with a tube and clip, and whose neck has a light plug of cotton acting as a filter separating the tissue from the 10-20 ml of clear water in the stem. This can be modified by actually macerating the diseased tissue in a blender in order to liberate more lethargic nematodes. The following modification was devised by Schuilling and Van Dinther (1981). 15 g of chopped tissue, suspended in 250 ml of water, are blended in an electric mixer for 30 seconds. The resultant suspension is made up to 1 litre in a bottle and allowed to stand for 30 minutes. The contents of the bottle are then sedimented over another container filled with water. After 30 minutes the contents of the lower bottle are discarded. The contents of the top bottle are sieved four times through a 60 µm sieve.

Obtaining samples of nematodes from living trees

A stainless steel tube, sharpened at one end, is driven at an angle of 45° at the point selected for sampling. The extracted core is placed in a blender with 50 ml of water and processed for 2 minutes. The contents of the blender are then poured into a dish and left for 20 minutes for the nematodes to emerge. The nematodes are then recovered by sieving. Generally, advantage is taken of the level of activity of the nematode during extraction methods. In coconut and the palmiste palms the nematodes are most active in the stem tissue except in the very necrotic regions. The core tissue generally shows a red cylinder of necrotic tissue.

Detection and Inspection

Top of page The stem of the coconut needs to be examined either by cutting through, or by taking a sample from the living tree.

Similarities to Other Species/Conditions

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No other pest causes the red ring symptoms characteristic of R. cocophilus infestation. The nematodes are similar to species belonging to Bursaphelenchus.

Prevention and Control

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Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Epidemiology

Red ring disease in new groves generally begins by infection of a 4-10 year old palm by the palm weevil, Rhynchophorus palmarum, carrying R. cocophilus. Effective patterns of control may be employed during several phases of the development of the epidemic.

The dispersal rate from the primary infector plant depends upon the development of R. palmarum within the diseased tree. Parasitism by the nematodes may limit the number of developing vectors and reduce the size, fecundity and longevity of the vector adults. Three months after infection, a new tree can be infected by a vector (female) emerging from the infector plant. If the insect is unmated and infertile, no vector will develop from this infection and red ring can die out when the diseased palm dies. This diseased tree, however, forms a source of inoculum as it becomes chemically attractive to all palm weevils including potential vectors. Phytosanitary measures of control are most effective at this time since disease symptoms are apparent before the progeny of the newly invaded insects emerge after 3 months.

Control in Coconut

There are no simple means of controlling R. cocophilus and no effective measures are available for control of the nematode in living palms. Control is based on prevention rather than cure either by the destruction of infested palm material by cutting and burning, or by the injection of herbicides and burning, or by trapping and killing of the weevil vectors before they spread the nematodes.

Many trees show yellowing and browning of leaves which may not be due to R. cocophilus attack. To prevent unnecessary destruction of trees, a core sample of the trunk should be taken with a 2-cm pipe (see Detection section) to determine the presence of R. cocophilus before control measures are employed.

Trapping

Traps or guard baskets are designed to protect plantations from frequent outbreaks of R. cocophilus. They do so by attracting and killing palm weevils which may enter the plantations from nearby diseased trees. Guard baskets are made of 2-cm-mesh wire. They are cylindrical, 1 m high and 0.3 m in diameter. These baskets are filled with chunks of fresh tissue from diseased coconut trees to attract the weevil. If such trees are not available, chunks of palmiste or 'gru-gru' trees may be used. The guard baskets are sprayed with pesticide and distributed on the ground in the plantations at one basket per acre (2.5 baskets/ha) of young coconut trees. This procedure is especially recommended in the dry season when the weevils are most active in the cool nights. Guard baskets remain for about 2 weeks, after which the tissue and insecticide in the basket should be burnt. Fresh tissue should be placed in the basket and treated as previously described. Several variations are used in practice with different types of tissue, such as pineapple and papaya (Griffith and Koshy, 1990).

References

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Alcocer GL, 1955. Enfermedad de la palma de coco conocida como `anillo rojo'. Fitofilo, Mexico, 8:8-11.

Anon., 1954. Report of the Department of Agriculture for the Year 1954. Belize, British Honduras: Ministry of Agriculture.

Brathwaite CWD, Siddiqi MR, 1975. Rhadinaphelenchus cocophilus. C.I.H. Descriptions of Plant-parasitic Nematodes, Set 5(No. 72):4 pp.

Briton-Jones HR, 1940. The Diseases of the Coconut Palm. London, UK: Bailliere Tindall and Cox.

CABI, EPPO, 1999. Rhadinaphelenchus cocophilus. [Distribution map]. Distribution Maps of Plant Diseases, April (Edition 1). Wallingford, UK: CAB International, Map 786.

Cabrera CJ, 1965. El anillo rojo del cocotero del Pacifico. Agricultura tropical, Colombia, 21:481-484.

Chaves Batista, A, 1948. O 'anel vermelho' do coqueiro e a fumigacao do solo com D-D. Boletim da Secretaria da Agricultura, Pernambuco, Brazil, 15: 356-387.

Dean CG, 1979. Red ring disease of Cocos nucifera L. caused by Rhadinaphelenchus cocophilus (Cobb, 1919) Goodey, 1960. Technical Communication, Commonwealth Institute of Helminthology, St. Albans, Herts, Uk Commonwealth Agricultural Bureaux. Farnham Royal, Bucks UK, No.47:70 pp

EPPO, 2014. PQR database. Paris, France: European and Mediterranean Plant Protection Organization. http://www.eppo.int/DATABASES/pqr/pqr.htm

Fenwick DW, 1959. Report on a visit to Venezuela. Red Ring Research Scheme, Ministry of Agriculture, Central Experimental Station, Centero, Trinidad & Tobago.

Fenwick DW, Maharaj S, 1963. Recovery of Rhadinaphelenchus cocophilus (Cobb, 1919) Goodey, 1960 from coconut tissues. Journal of Helminthology, 37:11-14.

Gerber K, Giblin-Davis RM, 1990. Association of the red ring nematode and other nematode species with the palm weevil, Rhynchophorus palmarum. Journal of Nematology, 22(2):143-149

Gontalves RD, 1937. A doenta do ` anel vermelho' do coqueiro. O. Biologico, 3:102-103.

Goodey JB, 1960. Rhadinaphelenchus cocophilos (Cobb, 1919) n.comb., the nematode associated with 'red-ring' disease of coconut. Nematologica, 5:98-102.

Griffith R, Koshy PK, 1990. Nematode parasites of coconut and other palms. In: Luc M, Sikora RA, Bridge J, eds. Plant Parasitic Nematodes in Subtropical and Tropical Agriculture. Wallingford, UK: CAB International, 363-386.

Liceras Zarate L, 1967. El nematodo: Rhadinaphelenchus cocophilus (Cobb, 1919) Goodey, 1960, agente causal de la enfermedad del Anillo Rojo del cocotero recientemente detectado en Tumbes. Agricultura y Ganaderia Tropical, Peru, 1:27-29.

Maas PWT, 1969. Two important cases of nematode infestation in Surinam. In: Peachey JE, ed. Nematodes of Tropical Crops. Technical Communication, Commonwealth Bureau of Helminthology, No.40, 149-154.

Nickle WR, 1970. A taxonomic review of the Aphelenchoidea (Fuchs, 1937) Thorne, 1949 (Nemotoda: Tylenchida). J. Parasit., 56:249.

Nowell W, 1919. Red ring disease of coco-nuts. Agricultural News, Trinidad, 18:398.

Ponte, JJ da, Lima, JAA, Brandine, E, 1971. O anel vermelho do coqueiro, no estado do Ceara. Pesq. Agrop. Nord., Recife, 3:85-87.

Schuiling M, Van Dither JBM, 1981. 'Red ring disease' in the Paricatuba oil palm Estate, Para, Brazil. A case study. Zeitschrift fur Angewandte Entomologie, 91:154-169.

Sharma RD, Loof PAA, 1982. Nematodes associated with declining coconut in Sergipe, Brazil. In: : Lordello LGE, ed. Trabalhos apresentados a VI reuniao Brasileira de nematologia, 8-12 fevereiro de 1982, Fortaleza. Publicacao No.6. Sociedade Brasileira de Nematologia Piracicaba, SP Brazil, 79-84

Singh ND, 1972. A survey of red ring disease of coconut palm in Grenada, West Indies. Plant Disease Reporter, 56:339-341.

Thorne G, 1961. Principles of Nematology. London, UK: McGraw Hill.

Tidman DA, 1951. Agricultural and horticultural problems of Brazil. World Crops, 3:341-344.

Distribution Maps

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