Meloidogyne ethiopica
(Root-knot nematode)

Identity

Preferred Scientific Name

• Meloidogyne ethiopica Whitehead

Preferred Common Name

• Root-knot nematode

International Common Names

• Spanish: nematodo agallador
• French: nematode à galles

Summary of Invasiveness

M. ethiopica is a polyphagous pest and classed as a tropical and temperate root-knot nematode (RKN) species (Strajnar et al., 2011). M. ethiopica is considered a damaging species as it can multiply on many different types of plants (including both dicotyledons and monocotyledons). Originally considered a tropical species, it has been shown to survive outdoors in temperate areas also. The distribution in Africa is unknown.

Only in Chile is it considered an invasive nematode. In Chile it occurs over a range of ca 1000 km and has been detected from the Copiapó valley, ca 800 km north of Santiago, to Talca, ca 250 km south of Santiago and it is found on grapevine (Vitis vinifera), kiwi (Actinida deliciosa C.) and potatoes in 80% of samples (Carneiro et al., 2007) collected in this area. It was introduced in Brazil from Chile on kiwi seedlings, and despite not being invasive in Brazil it has caused serious economic problems to grapevine in Chile (Carneiro et al., 2007). Recently, this species has also been recorded on asparagus (Asparagus officinalis L.) in Peru (Murga-Gutierrezet al., 2012). In Europe it has been detected only in Slovenia, on greenhouse tomatoes (Solanum lycopersicum L.) (Širca et al., 2004). It was added to the EPPO Alert List in 2011.

Taxonomic Tree

• Domain: Eukaryota
•     Kingdom: Metazoa
•         Phylum: Nematoda
•             Family: Meloidogynidae
•                 Genus: Meloidogyne
•                     Species: Meloidogyne ethiopica

Notes on Taxonomy and Nomenclature

Meloidogyne ethiopica Whitehead, 1968 was described from a single egg mass culture on tomato (type host) from the Mlalo region, Lushoto District, Tanga Province, Tanzania (type locality); cowpea was also given as a host. At the same time, Whitehead (1968) studied specimens of this species from a single egg mass culture on tomato sent from Zimbabwe and slides from South Africa; both of these populations had been previously identified as M. arenaria (Neal, 1889) Chitwood, 1949. In the original description the perineal patterns of M. ethiopica were characterized as varying from M. arenaria to M. incognita (Kofoid and White, 1919) Chitwood, 1949, even with specimens from single egg mass cultures. It seems possible that the nature of the perineal pattern of M. ethiopica has made its accurate identification difficult or impossible, especially in the absence of other highly distinctive features of the female (Golden, 1992).

M. ethiopica was collected and reported again by Whitehead (1969) from the Mlalo region of Tanzania on bean (Vicia faba), black wattle (Acacia mearnsii), cabbage, pepper (Capsicum frutescens), potato, pumpkin and tobacco. O’ Bannon (1975) found M. ethiopica in two locations in Ethiopia; on Awasa Road, one of the locations, specimens were collected from lettuce, soybean, sisal and three weeds (Ageratum conyzoides, Datura stramonium, Solanum nigrum). Jepson (1987) discussed this species but had no further information on its morphology and distribution and hosts. Recently, it was re-described based on its detection in Brazil and Chile parasitizing kiwi (Actinidia deliciosa (Chevalier) Liang and Ferguson) and grapevine (Vitis vinifera L.) plants, respectively (Carneiro et al., 2003; 2004).

Description

The redescription of M. ethiopica Whitehead, 1968 (Carneiro et al., 2004) was made by comparing the populations from Brazil with the populations from Chile and Kenya identified as M. ethiopica (G Karssen, pers. comm.). Unfortunately, the type material was not in good condition, so only the perineal patterns allowed good observation. The measurements of the population from Brazil are presented in Table 1. Major quantitative and qualitative characters varied only slightly among the three studied populations of M. ethiopica, but distinguishing features were not observed.

Female

Body translucent–white, variable in size, elongated, ovoid to pear-shaped. Neck sometimes prominent, posterior end rounded, without distinct protuberance. Body cuticle distinctly annulated, annuli smaller in anterior neck region. Head region set off, usually marked by annulations. Head cap distinct, labial disc squarish and elevated. Cephalic framework weakly sclerotized. Under SEM stoma slit-like, located in ovoid prestomatal cavity, surrounded by pit-like openings of six inner labial sensilla. Raised labial disc, separated from lateral and medial lips. Lateral lips triangular, separated from the head region. Stylet robust, cone generally slightly curved dorsally and gradually increases in width posteriorly. Likewise, the shaft gradually widening posteriorly to near the junction with the stylet knobs, which are rounded shaped and gradually tapering onto the shaft. Excretory pore located between dorsal pharyngeal gland orifice and metacorpus.Perineal patterns oval to squarish, striae coarse, widely separated, usually continuous, smooth to wavy. Tail terminus sometimes distinct, with or without very fine, broken striation. Fold over anus present. Vulva slit-like without striae near lateral. Phasmids distinct. Dorsal arch moderately high to high, rounded to squarish, never forming "shoulders". Distinct lateral lines absent. Sometimes lateral lines with short, vertical striae near phasmid area. Ventral part region rounded.

Male

Body vermiform, tapering anteriorly, bluntly rounded posteriorly, length variable. Head cap high, rounded, continuous with body contour. In LM cephalic framework strongly developed, vestibule and extension distinct. Stylet robust, large, cone straight, pointed as long as shaft, gradually increasing in diameter, posteriorly, shaft cylindrical widening slightly near junction with knobs. Knobs smooth, rounded–pear shaped and back-wardly sloping. Head region slightly set off and marked sometimes by incomplete annulation. Head cap with distinct labial disc, amphidial opening elongated slits. In SEM (face view), high labial disk almost circular to hexagonal, distinctly separated from fused medial lips. Medial lips crescent shaped with distinctly lateral indentation at junction with labial disc. Diameter of medial lips smaller than labial disc. Cephalic sensilla obscure. Lateral lips absent. Stoma opening slit-like, situated below large, hexagonal prestoma. Inner labial sensilla indistinct, opening into prestomatal cavity. Lateral field with four incisures, aerolated. Procorpus well defined. Metacorpus oval shaped, with large valve. Pharingo-intestinal junction obscure at level of nerve ring. Gland-lobe variable in length, with two nuclei. Caecum extending to level of metacorpus. Excretory pore position variable, terminal excretory duct long. Hemizonid two to four annules anterior to excretory pore. Most males sex reversed with two testes, sometimes one testis in normal developed males. Testis outstretched or distally reflexed. Spicules are thicker walled, with strongly ridged shaft, arcuate, head cylindrical, set off. Blade tip slightly curved ventrally, with two distal pores. Gubernaculum distinct. Tail short, phasmids at level of cloaca.

Second–Stage Juvenile (J2)

Body vermiform, tapering more posteriorly than anteriorly, tail region distinctly narrowing. Body annules distinct, increasing in size and becoming irregular in posterior tail region. Head region truncate without annulations, not set off from body. Head cap low, narrower than head region. Under SEM, stoma slit-like located in ovoid prestomatal cavity, surrounded by pit-like openings of six inner labial sensilla. Labial disc rounded, distinctly raised. Outer margins of medial lips crescent shaped. Medial lips and labial disc dumbbell shaped. Lateral lips fused at right angle with medial lips, lower than medial lips, margins rounded to slightly triangular, fusing or not with head region. Cephalic sensilla indistinct in LM cephalic framework weak, hexaradiate. Vestibule and vestibule extension more distinct than the rest of framework. Stylet cone increasing in width gradually, shaft cylindrical to tapering posteriorly; knobs rounded and set off from shaft. Orifice of oesophagal gland branched into three channels; ampulla indistinct. Procorpus faintly outlined, metacorpus ovoid with prominent valve; isthmus not clearly outlined; triradiate lining strongly sclerotized; subventral gland orifices branched, located immediately posterior to enlarged lumen of median bulb. Pharyngo–intestinal junction indistinct, at level of nerve ring. Gland lobe of variable length with three nuclei. Excretory pore distinct; hemizonid two to four annules anterior to excretory pore. Tail slender, ending in rounded tip or in a pointed thin tip with annules of sometimes irregular size in the posterior region; hyaline tail terminus distinct; rectal dilation large. Lateral fields with four incisures extending the entire length of the nematode and narrowing in the tail region in three areolated incisures. Areolation not necessarily involving the entire length of lateral fields. Phasmids in ventral incisure of lateral field, always posterior to anus.

Distribution Table

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.

History of Introduction and Spread

In 1989 kiwi plants (Actinidia deliciosa (Chevalier) Liang and Ferguson) from Chile (Curicó) were introduced to Lagoa Vermelha (Serra Gaúcha region), in the State of Rio Grande do Sul, Brazil. Ten years later, poor development of the plants was associated with high populations of root-knot nematode (RKN).

These populations expressed an atypical enzymatic profile, named Ki3, which had never been found in other RKN populations from Brazil (Carneiro et al., 1996; 2000). To confirm the hypothesis that this nematode was introduced from Chile (JC Magunacelaya., pers. comm.), samples containing grapevine (Vitis vinifera L.) roots, infected by Meloidogyne sp., collected in Casa Blanca, Chile, were analyzed using the esterase phenotype. The same esterase profile Ki3 was observed, proving that the nematode was the same species as that in Brazilian kiwi plants. So far in Chile, the same esterase phenotype Ki3 has been detected on kiwi fruit in a survey conducted in Central Valley, although this population was incorrectly considered as M. javanica (Treub, 1885) Chitwood, 1949 (Philippi et al., 1996).

Based on morphological, biological and molecular differences with other species of Meloidogyne,and based on a comparison with the type material and onepopulation of M. ethiopica from Kenya, these nematode isolates from Brazil and Chile are identified, characterized and illustrated as M. ethiopica Whitehead,1968 (Carneiro et al, 2004). In Chile, it is suspected that movements of contaminated grapevine nursery stock have probably resulted in serious infestations in various vineyards. Some surveys made in Brazil showed that M. ethiopica is not a widespread species in kiwi plants in Rio Grande do Sul state (Somavila et al., 2011), or on soybean (Castro et al., 2003), vegetables (Carneiro et al., 2008) or on infested bulbs of Polymnia sonchifolia (yacón or Peruvian ground apple) in Distrito Federal (Carneiro et al., 2005). Although M. ethiopica occurs frequently in vineyards in Chile, it has not been found on grapevines in Brazil yet (Somavila, 2011).

In 2004, Meloidogyne ethiopica was found for the first time in a tomato greenhouse in Slovenia (Širca, 2004). This was also the first record for Europe.

Introductions

Risk of Introduction

As a root-knot nematode species, M. ethiopica can easily be transmitted with soil and plant root material. Infested soil and growing media, plants for planting, bulbs and tubers from countries where M. ethiopica occurs are the most probable pathways of introduction into different regions. Soil attached to machinery, tools, footwear or plant products is also another possible pathway. M. ethiopica is a polyphagous species and many of its host plants are of economic importance as they are cultivated as arable, vegetable, ornamental or fruit crops. The recent detection of this pest in Brazil, Chile and Slovenia clearly demonstrated that it has the potential to enter in different regions. Recent studies have showed that, despite its tropical origin, M. ethiopica has the potential to survive outdoors in a continental climate (hot summers and cold winters), even in areas where soil temperatures fall below zero during winter, as well as in sub-Mediterranean climate (Strajnar et al., 2011). This indicates that M. ethiopica could establish and spread in different regions of different countries. In addition, M. ethiopica could survive under glasshouse conditions across regions with a sub-Mediterranean or a continental climate. Once root-knot nematodes have been introduced, it is generally difficult to control or eradicate them. Only in EPPO regions has this nematode been listed as a quarantine pest (EPPO, 2011).

Habitat

M. ethiopica is a tropical and temperate root-knot nematode species detected essentially on cultivated crops. It is a new pest in South America and Europe (Carneiro et al, 2003; 2007; Sirca et al., 2004). In Copiapo and Talca (Central Valley, Chile), where M. ethiopica is present, the weather is hot during the summer and cold during the winter. In Brazil, M. ethiopica occurrs in subtropical (Rio Grande do Sul, Santa Catarina and Paraná States) and tropical areas (Distrito Federal and São Paulo States).

Strajnar et al. (2009) examined its ability to survive in open fields located in regions with sub-Mediterranean and continental European climates. The outdoor field experiment consisted of two locations and extended over three growing seasons and two winter seasons. The results demonstrated that M. ethiopica was able to survive at both locations and also that it retained its infection ability despite several instances of temperatures falling below freezing.

Habitat List

Hosts/Species Affected

M. ethiopica is considered as a damaging species as it can multiply on many different types of plants, incuding both dicotyledons and monocotyledons (Sirca et al., 2004).

M. ethiopica Whitehead, 1968 was described from a single egg mass culture on tomato (type host), Tanzania (type locality); cowpea was also given as a host. M. ethiopica was collected and reported again by Whitehead (1969) from the Mlalo region of Tanzania on bean (Vicia faba), black wattle (Acacia mearnsii), cabbage, pepper (Capsicum frutescens), potato, pumpkin and tobacco. O’ Bannon (1975) collected specimens from lettuce, soybean, sisal and three weeds (Ageratum conyzoides, Datura stramonium, Solanum nigrum) from Awasa Road, Ethiopa.

Glasshouse tests with agronomic crops important to Brazil revealed that rice cv. BR 410, soybean cv. Cristalina, peach cv. Capdebosq and grapevine (Vitis labrusca) cv. Niágara Rosa are good hosts, whereas wheat cv. BR4, apple rootstocks cvs Maruba and M7, pear rootstock Calleryana, strawberry cvs Dover and Vila Nova, raspberry cv. Tupi, mulberry cv. Batu, blueberry cv. Powderblue and grapevine (Vitis rupestris) cv. Rupestris du Lot are non-hosts (Carneiro et al., 2003). In another test, tomato cv. Rutgers, tobacco cv. NC95, pepper cv. California Wonder and watermelon cv. Charleston Gray were all found to be good hosts, whilst cotton cv. Deltapine 61 and peanut cv. Florunner were non-hosts (Carneiro et al., 2004). Recent surveys conducted in Brazil showed that M. ethiopica presented a restricted distribution in Rio Grande do Sul on kiwi (Somavila et al., 2011) and in Distrito Federal States on vegetables (Carneiro et al., 2008).

In 2003, M. ethiopica was reported for the first time in a tomato greenhouse in Slovenia. This was also the first record for Europe (Sirca et al., 2004).

Host Plants and Other Plants Affected

Growth Stages

List of Symptoms/Signs

Biology and Ecology

Biochemical Study

Species-specific esterase phenotypes (E3) with three bands (Rm: 0.9, 1.05, 1.20) were observed in the three isolates from Brazil, Chile and Kenya, as well as a malate dehydogenase phenotype N1 (Rm 1,0), which is shared with several species of Meloidogyne (Carneiro et al., 2000).

Genetics

The reproduction of M. ethiopica is by mitotic parthenogesis and the chromosome number is 36-38 (Carneiro et al., 2004).

Reproductive biology

The following is taken from Moens et al. (2009):

Females lay eggs into gelatinous masses composed of a glycoprotein matrix, which is produced by rectal glands in the female. The glycoprotein matrix it keeps the eggs together and protects them against environmental extremes and predation. The egg masses are usually found on the surface of galled roots, although they may also be embedded within the gall tissue. The egg mass is initially soft, sticky and hyaline but becomes firmer and dark brown with age. Surprisingly, there has been only limited analysis of the glycoproteins (Sharon and Spiegel, 1993) or other components of the gelatinous matrix, despite its obvious importance. Within the egg, embryogenesis proceeds to the first-stage juvenile, which moults to the infective second-stage juvenile (J2). Hatch of the J2 is primarily dependent on temperature and sufficient moisture, although other factors, including root diffusate and generation, modify the hatching response so that the J2 hatch when conditions are favourable for movement and host location; according Somavilla (2011), the temperatures 25°C and 30°C are most favorable to M. ethiopica  J2 hatching. The ability of M. ethiopica to survive is enhanced by several physiological and biochemical adaptations, including delayed embryogenesis, quiescence and diapause, and lipid reserves that prolong viability until the J2 reaches and invades a host. In the soil, the J2 is vulnerable and needs to locate a host as rapidly as possible. J2 are attracted to roots, and there is evidence that when both resistant and susceptible plant roots are present the susceptible ones are more attractive.

The invasive J2 commences feeding after it has invaded the root, usually behind the root tip, and moved through the root to initiate and develop a permanent feeding site. The feeding of the J2 on protoxylem and protophloem cells induces these cells to differentiate into specialized nurse cells, which are called giant cells. Once a giant cell is initiated, the nematode becomes sedentary and enlarges greatly to assume a ‘sausage’ shape. Under favorable conditions, the J2 stage moults to the third-stage juvenile (J3) after ca14 days, then to the fourth-stage juvenile (J4), and finally to the adult stage. The combined time for the J3 and J4 stages is much shorter than for the J2 or the adult, and typically takes 4–6 days. J3 and J4 lack a functional stylet and do not feed. Males, when present, are vermiform and there is no evidence that they feed. Males may be found in parthenogenetic species when conditions are unfavorable for female development, such as when population densities are very high and presumably there is a limitation of food supply.

The initiation, development and maintenance of the giant cell is the subject of continuing investigation, facilitated by molecular techniques with the impetus of developing novel control strategies based on preventing giant cell formation or, more likely, development. These specialized feeding sites are remarkable for their complexity. They are greatly enlarged from typical phloem and xylem parenchyma, or cortical cells, with final cell volumes nearly 100-fold greater than normal root cells. The giant cells are functionally similar to syncytia induced by other plant-parasitic nematodes that have sedentary adult females.

Longevity

M. ethiopica required 67, 48 and 36 days to complete one reproduction cycle at mean daily temperatures of 18.3, 22.7 and 26.3ºC, respectively, in a study by Strajnar et al. (2011). Reproduction did not occur at 13.9ºC. On grape susceptible rootstock  M. ethiopica completed its life cycle in 27 days at 25ºC (Somavilla, 2011).

Activity Patterns

Strajnar et al. (2011) examined the survival ability of M. ethiopica in continental climate conditions that are characterised as mild, with hot summers and cold winters, as well as in a sub-Mediterranean climate characterised by hot summers and mild winters. M. ethiopica survived and maintained infection ability in open fields in European climate conditions, despite the fact that the winter temperatures fell below zero degrees several times. The survival of the nematode population was observed for two winters and three growing seasons. Root galling, an indicator of nematode infection, increased in the second growing season.

Population Size and Density

The population density is elevated on susceptible tomato plants, similar to other important root-knot nematode species like M. incognita, M. javanica and M. arenaria.

Nutrition

M. ethiopica is an obligatory plant parasite (hosts are listed Hosts/Species Affected).

Associations

The associations with wild plants are relevant to the survival or invasiveness in field conditions; however, this has not been very well studied.

Environmental Requirements

Strajnar et al. (2011) carried out four experiments in a growing chamber using a range of mean daily temperatures (13.9, 18.3, 22.7 and 26.3ºC). M. ethiopica required 67, 48 and 36 days to complete one reproduction cycle at mean daily temperatures of 18.3, 22.7 and 26.3ºC, respectively. Reproduction did not occur at the lowest temperature (13.9ºC). In a histological study conducted by Somavilla (2011) in root sections of grape susceptible rootstock inoculated with M. ethiopica, it was verified that the nematode could to penetrate, develope and complete its life cycle with 27 days of inoculation at 25ºC.

Climate

Means of Movement and Dispersal

The wide distribution of M. ethiopica in Chile was probably caused by contaminated grapevine seedlings. In Chile, the prohibition on producing seedlings in areas with root-knot nematodes, and on the movement of infested seedlings into new growing areas, seem not to be effective, considering that many nursery plants across the country are severely infected with M. ethiopica.

This nematode was probably introduced into Rio Grande do Sul State, Brazil, accidentally from Chile (Curico locality) in 1989 on kiwi seedlings.

Pathway Causes

Pathway Vectors

Plant Trade

Impact Summary

Impact

Data is lacking on the extent of damage and the environmental and economic impact this nematode may cause on its different host plants. The use of toxic nematicides can contaminate soils and water and impact human and animal health.

Risk and Impact Factors

• Invasive in its native range
• Abundant in its native range
• Highly adaptable to different environments
• Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
• Capable of securing and ingesting a wide range of food
• Benefits from human association (i.e. it is a human commensal)
• Long lived
• Fast growing
• Has high reproductive potential
• Gregarious
• Reproduces asexually

• Negatively impacts agriculture
• Negatively impacts cultural/traditional practices
• Damages animal/plant products

• Induces hypersensitivity
• Interaction with other invasive species
• Parasitism (incl. parasitoid)
• Pathogenic
• Rapid growth
• Rooting

• Highly likely to be transported internationally accidentally
• Highly likely to be transported internationally illegally
• Difficult to identify/detect as a commodity contaminant
• Difficult to identify/detect in the field
• Difficult/costly to control

Diagnosis

A characteristic esterase isozyme patterns E3 (Rm: 0.9, 1.05,1.20) can be used to provide a more rapid and reliable identification of M. ethiopica (Carneiro et al., 2003; 2004).

Detection and Inspection

Roots and tubers should be inspected for small and large multiple galls. Females can be collected from inside the roots.

The correct identification of M. ethiopica requires a detailed study of the distinctive morphological features of females, males and second-stage juveniles (J2) (see Description). By comparing the populations from Brazil and Chile and the population sent as M. ethiopica from Kenya, it is possible to see that they present very similar morphological and morphometrical features, as well as the following common characters, as described for the population from Tanzania (Whitehead, 1968): the perineal patterns of females are highly variable, going from M. arenaria (Neal, 1889) Chitwood, 1949 to M. incognita (Kofoidand White, 1919) Chitwood, 1949, but are very similar to the examined types. Phasmids are large and distinct, with a conspicuous phasmidial canal, and are also observed in the three populations, a good diagnostic character for differentiating this species from M. arenaria and M. incognita (Golden, 1992). Head region of females set off with elevated squarish labial disc, usually marked by two or three annulations. Female stylet robust, curved dorsally with small stylet knobs and the shaft gradually widening posteriorly to near the junction with the stylet knobs. Head region of males set off and sometimes marked by annules on the three populations. Male stylet robust, knobs small, rounded, usually sloping backwards. Lateral fields with four incisures, areolated, the spicules more thickly walled with strongly ridged shafts. Head region of second-stage juveniles(J2) truncate, not set off from the body, without annulations. The tail is slender with the terminus rounded in a pointed thin tip. In combination, these characters are useful to differentiate M. ethiopica from other known Meloidogyne spp.

Similarities to Other Species/Conditions

The population of M. ethiopica described by Whitehead (1968) from Tanzania, and the three other populations studied by Carneiro et al. (2004), differ in some morphological and morphometrical features. The male head region is either marked by two annules behind the head region (Tanzania) or sometimes marked by one or two annules (Kenya, Chile and Brazil). Other differences include: the length of male stylets (14.4–24.1 [21.4] µm in Tanzania vs 22.8-26.8 [24.8] µm in Kenya, Chile and Brazil); the stylet length of second-stage juveniles (J2) (9.1-10.9 [10.0] µm vs 11.2-13.5 [12.2] µm); the tail length of J2 (41.0-52.0 [47.0] µm vs 52.2-72.4 [61.7] µm), and the hyaline tail terminus length ([8.4] µm 12.0-15.5 [13.5] µm) (Jepson, 1987). However, many of the morphological characters used in the past to distinguish species, such as the number of annulations on the head region and many body measurements, have been found to be variable and unreliable for Meloidogyne species characterization (Esser et al., 1976; Franklin,1979; Jepson, 1983).

Based on morphological characteristics, M. ethiopica can be confused with M. incognita, M.arenaria and M. inornata. However, characteristic esterase isozyme patterns, known as E3, have been described for M. ethiopica to provide more rapid and reliable identification (Carneiro et al., 2003; 2004).

Prevention and Control

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.

The use of toxic nematicides can contaminate soils and water and can impact human and animal health.

In Chile, the prohibition on producing seedlings in areas with root-knot nematodes, and on the movement of infested seedlings into new growing areas, seem not to be effective, considering that many nursery plants across the country are severely infected with M. ethiopica.

Preventing crop infection in the first instance is perhaps the single most important strategy to avoid or limit crop losses in terms of quality and yield. This is particularly true since the treatment of nematode-infected crops, or a ‘therapeutic’ approach, is essentially more complicated and costly for producers. The use of clean, healthy, nematode-free planting material is a prerequisite for good crop production and cannot be overemphasized. Within an overall cropping system, the physical removal or destruction of plant material infected with root-knot nematodes, particularly roots or tubers, should be considered.

Planting crops that are poor or non-hosts of root-knot nematodes in rotation in infested soil reduces nematode populations. For the management of M. ethiopica in infested soil Lima et al. (2009) recommend planting successions of different summer and winter non-host plants:

‘A. Mucuna deeringiana, Avena strigosa, Arachis hypogaea; B. Crotalaria spectabilis, Lolium multiflorum, Cajanus cajan ‘PPI 832’; C. Vigna unguiculata, Secale cereale ‘IPR 69’, Cotralaria apioclice; D. Ricinus communis ‘IAC 80’, Raphanus sativus ‘IPR 116’, Crotalaria grantiana; E. Crotalaria grantiana, Secale cereale ‘IPR 69’, Cajanus cajan ‘PPI 832’; F. Crotalaria grantiana, Lolium multiflorum, Mucuna deeringiana; G. Vigna unguiculata, Avena strigosa, Arachis hypogaea. These plant sequences should be adapted to different regions and areas where M. ethiopica is a major agricultural problem’ (Lima et al., 2009). Ricinus communis ‘BRS Energia’, ‘CPACT 40’, ‘AL Guarani’, ‘Sara’, ‘Lyra and ‘Nordestina’ can also be used (Santos and Gomes, 2011).

In case of perennial plants like grapevine, use resistant rootstocks: SO4, IAC 313, P1103 88d, Salt Creck, K5 BB Kober’s and Harmony (Somavilla et al., 2008). Other fruit perennial species as Eugenia involucrata, E. guabiju, E. uvalha, E. uniflora, Campomanesia xanthocarpa, Myciaria cauliflora, Morus nigra ‘Xavante’ and ‘Guarani’, Psidium cattleianum and Citrus sunki ‘Maravilha’ and ‘Tropical’ are also poor hosts of M. ehtiopica (Somavilla et al., 2009).

Gaps in Knowledge/Research Needs

Informatio is lacking on the extent of the damage and the environmental and economic impacts this nematode may cause on its different host plants.

The associations with wild plants are relevant to the survival or invasiveness of M. ethiopica in field conditions, but have not been very well studied.

The development of a molecular marker to identify M. ethiopica would be beneficial, as would a study of the resistance of different crops like potato, soybean and kiwi.

References

Aydinli G, Mennan S, Devran Z, Sirca S, Urek G, 2013. First report of the root-knot nematode Meloidogyne ethiopica on tomato and cucumber in Turkey. Plant Disease, 97(9):1262. http://apsjournals.apsnet.org/loi/pdis

Bellé C, Kaspary TE, Schmitt J, Kuhn PR, 2016. Meloidogyne ethiopica and Meloidogyne arenaria parasitizing Oxalis corniculata in Brazil. Australasian Plant Disease Notes, 11(1):24. http://link.springer.com/article/10.1007/s13314-016-0212-7

Bellé, C., Kulczynski, S. M., Kuhn, P. R., Carneiro, R. M. D. G., Lima-Medina, I., Gomes, C. B., 2017. First report of Meloidogyne ethiopica parasitizing sugarcane in Brazil., Plant Disease, 101(4):635 http://apsjournals.apsnet.org/loi/pdis

CABI/EPPO, 2005. Meloidogyne ethiopica. Distribution Maps of Plant Diseases, No. 962. Wallingford, UK: CAB International.

CABI/EPPO, 2013. Meloidogyne ethiopica. [Distribution map]. Distribution Maps of Plant Diseases, No.October. Wallingford, UK: CABI, Map 962 (Edition 2).

Carneiro RMDG, Almeida MRA, 2005. First record of Meloidogyne ethiopica Whitehead on yacon and tomato plants in Brasília, DF, Brazil. Nematologia, 29(2):285-287.

Carneiro RMDG, Almeida MRA, Carneiro RG, 1996. Enzyme phenotypes of Brazilian populations of Meloidogyne spp. Fundamental and Applied Nematology, 19(6):555-560.

Carneiro RMDG, Almeida MRA, Cofcewicz ET, Magunacelaya JC, Aballay E, 2007. Meloidogyne ethiopica, a major root-knot nematode parasitizing Vitis vinifera and other crops in Chile. Nematology, 9(5):635-641. http://www.ingentaconnect.com/content/brill/nemy

Carneiro RMDG, Almeida MRA, Martins I, Souza JF, Pires AQ, Tigano MS, 2008. Occurrence of Meloidogyne spp. and nematophagous fungi on vegetables in the Federal District of Brazil. (Ocorrência de Meloidogyne spp. e fungos nematofagos em hortaliças no Distrito Federal, Brasil.) Nematologia Brasileira, 32(2):135-141.

Carneiro RMDG, Almeida MRA, QuTnThervT P, 2000. Enzyme phenotypes of Meloidogyne spp. populations. Nematology, 2(6):645-654; 20 ref.

Carneiro RMDG, Gomes CB, Almeida MRA, Gomes ACMM, Martins I, 2003. First record of Meloidogyne ethiopica Whitehead, 1968 on kiwi in Brazil and reaction of different plant species. (Primeiro Registro de Meloidogyne ethiopica Whitehead, 1968, em plantas de quivi no Brasil e reação em diferentes plantas cultivadas.) Nematologia Brasileira, 27(2):151-158.

Carneiro RMDG, Randig O, Almeida MRA, Gomes ACMM, 2004. Additional information on Meloidogyne ethiopica Whitehead, 1968 (Tylenchida: Meloidogynidae), a root-knot nematode parasitising kiwi fruit and grape-vine from Brazil and Chile. Nematology, 6(1):109-123. http://www.brill.nl

Castro JMda CE, Lima RDde, Carneiro RMDG, 2003. Isoenzymatic variability in Brazilian populations of Meloidogyne spp. from soybean. (Variabilidade isoenzimática de populações de Meloidogyne spp. provenientes de regiões Brasileiras produtoras de soja.) Nematologia Brasileira, 27(1):1-12.

Conceição IL, Tzortzakakis EA, Gomes P, Abrantes I, Cunha MJda, 2012. Detection of the root-knot nematode Meloidogyne ethiopica in Greece. European Journal of Plant Pathology, 134(3):451-457. http://springerlink.metapress.com/link.asp?id=100265

EPPO, 2011. EPPO Reporting Service. EPPO Reporting Service. Paris, France: EPPO. http://archives.eppo.org/EPPOReporting/Reporting_Archives.htm

EPPO, 2014. EPPO Reporting Service, No. 2014/007. Paris, France: European and Mediterranean Plant Protection Organization.

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

Esser RP, Perry VG, Taylor AL, 1976. A diagnostic compendium of the genus Meloidogyne (Nematoda: Heteroderidae). Proceedings of the Helminthological Society of Washington, 43(2):138-150.

Franklin MT, 1979. Taxonomy of the genus Meloidogyne. Root-knot nematodes (Meloidogyne species); systematics, biology and control. London & New York, USA, UK: Academic Press Inc., 37-52.

Golden AM, 1992. Large phasmids in the female of Meloidogyne ethiopica Whitehead. Fundamental and Applied Nematology, 15(2):189-191.

Gomes CB, Carbonari JJ, Medina IL, Lima DL, 2005. Survey of Meloidogyne ethiopica in kiwi in Rio Grande do Sul State, Brazil, and its association with Nicotiana tabacum and Sida rhombifolia. In: Abstract of a paper presented at the 25th Congresso Brasileiro de Nematologia (Brazilian Congress of Nematology), 29(1). Nematologia Brasileira, 114.

Jepson SB, 1983. Identification of Meloidogyne: a general assessment and comparison of males morphology using light microscopy, with a key to 24 species. Revue de Nematologie, 6:291-309.

Jepson SB, 1987. Identification of root-knot nematodes (Meloidogyne species). (continued). Identification of root-knot nematodes (Meloidogyne species). C.A.B. International., UK x + 265pp.

Lima EA, Mattos JK, Moita AW, Carneiro RG, Carneiro RMDG, 2009. Host status of different crops for Meloidogyne ethiopica control. Tropical Plant Pathology, 34(3):152-157. http://www.scielo.br/scielo.php?script=sci_arttext&pid=S1982-56762009000300003&lng=en&nrm=iso&tlng=en

Lima-Medina I, Gomes CB, Nazareno NXR, 2011. Occurrence of Meloidogyne ethiopica in potato in the state of Parana. (Registro da ocorrencia de Meloidogyne ethiopica em batata no estado do Parana. Anais do 44o Congresso Brasileiro de Fitopatologia. 44th Congresso Brasileiro de Fitopatologia, Bento Goncalves-RS, 2011.) In: 44th Brazilian Congress of Plant Pathology, 36 [ed. by Lavras, M. G.]., Brazil: Brazilian Congress of Plant Pathology, 177.

Mandefro W, Dagne K, 2000. Cytogenetic and esterase isozyme variation of root-knot nematode populations from Ethiopia. African Journal of Plant Protection, 10:39-47.

Moens M, Perry RN, Starr JL, 2009. Meloidogyne species - a diverse group of novel and important plant parasites. In: Root-knot nematodes [ed. by Perry, R. N.\Moens, M.\Starr, J. L.]. Wallingford, UK: CABI, 1-17. http://www.cabi.org/CABeBooks/default.aspx?site=107&page=45&LoadModule=PDFHier&BookID=461

Murga-Gutierrez SN, Colagiero M, Rosso LC, Sialer MMF, Ciancio A, 2012. Root-knot nematodes from asparagus and associated biological antagonists in Peru. Nematropica, 42(1):57-62. http://journals.fcla.edu/nematropica/article/view/79582/76900

O'Bannon JH, 1975. Nematode survey in Ethiopia. Addis Ababa, Ethiopia: Institute of Agricultural Research and FAO Rome, 29 pp.

Philippi I, Latorre BA, PTrez GF, Castillo L, 1996. Identification of the root-knot nematodes (Meloidogyne spp.) on kiwifruit by isoenzyme analysis in Chile. Fitopatologi^acute~a, 31(2):96-101; 34 ref.

Santos AV, Gomes CB, 2011. Reaction of castor bean cultivars to Meloidogyne spp. and effect of root exudates on Meloidogyne enterolobii and M. graminicola. (Reação de cultivares de mamona a Meloidogyne spp. e efeito dos exsudatos radiculares sobre Meloidogyne enterolobii e M. graminicola.) Nematologia Brasileira, 35(1/2):1-9. http://docentes.esalq.usp.br/sbn/nbonline/ol%203512/1-9%20co.pdf

Sharon E, Spiegel Y, 1994. Glycoprotein characterization of the gelatinous matrix in the root-knot nematode Meloidogyne javanica. Journal of Nematology, 25(4):585-589.

Sirca S, Urek G, Karssen G, 2004. First report of the root-knot nematode Meloidogyne ethiopica on tomato in Slovenia. Plant Disease, 88(6):680.

Somavilla L, Gomes CB, Antunes LEG, Oliveira RPde, Carneiro RMDG, 2009. Reaction of different fruit crops to Meloidogyne ethiopica. (Reação de diferentes frutíferas a Meloidogyne ethiopica.) Nematologia Brasileira, 33(3):252-255.

Somavilla L, Gomes CB, Carbonari JJ, Carneiro RMDG, 2011. Survey and characterization of root-knot nematode species in kiwi in Rio Grande do Sul State, Brazil. (Levantamento e caracterização de espécies do nematoide das galhas em quivi no Rio Grande do Sul, Brasil.) Tropical Plant Pathology, 36(2):89-94. http://www.scielo.br/scielo.php?script=sci_arttext&pid=S1982-56762011000200004&lng=en&nrm=iso&tlng=pt

Somavilla L, Gomes CB, Naves Lellis RDe, 2008. Reaction of grapevine rootstocks to Meloidogyne ethiopica Whitehead, 1968. (Reacao de porta-enxertos de videira a Meloidogyne ethiopica Whitehead, 1968. 41o Congresso Brasileiro de Fitopatologia, Belo Horizonte-MG, 2008.) In: Proceedings of the 41st Brazilian Congress of Phytopathology., Brazil: Brazilian Congress of Plant Pathology, 261.

Strajnar P, Širca S, Knapic M, Urek G, 2011. Effect of Slovenian climatic conditions on the developmentand survivalof the root-knot nematode Meloidogyne ethiopica. European Journal of Plant Pathology, 129:81-88.

Strajnar P, Sirca S, Stare BG, Urek G, 2009. Characterization of the root-knot nematode, Meloidogyne ethiopica Whitehead, 1968, from Slovenia. Russian Journal of Nematology, 17(2):135-142. HTTP://www.russjnematology.com

Tigano MS, Carneiro RM, Jeyaprakash A, Dickson DW, Adams BJ, 2005. Phylogeny of Meloidogyne spp. based on 18S rDNA and the intergenic region of mitochondrial DNA sequences. Nematology, 7(6):851-862.

Whitehead AG, 1968. Taxonomy of Meloidogyne (Nematodea: Heteroderidae) with descriptions of four new species. Trans . zool. Soc. Lond, 31(3):263-401.

Whitehead AG, 1969. The distribution of root-knot nematodes (Meloidogyne spp.) in tropical Africa. Nematologica, 15(3):315-333.

Distribution References

Aydınlı G, Mennan S, Devran Z, Širca S, Urek G, 2013. First report of the root-knot nematode Meloidogyne ethiopica on tomato and cucumber in Turkey. Plant Disease. 97 (9), 1262. http://apsjournals.apsnet.org/loi/pdis DOI:10.1094/PDIS-01-13-0019-PDN

CABI, Undated. Compendium record. Wallingford, UK: CABI

CABI, Undated a. CABI Compendium: Status as determined by CABI editor. Wallingford, UK: CABI

CABI/EPPO, 2013. Meloidogyne ethiopica. [Distribution map]. In: Distribution Maps of Plant Diseases, Wallingford, UK: CABI. Map 962 (Edition 2).

Carneiro R M D G, Almeida M R A, Cofcewicz E T, Magunacelaya J C, Aballay E, 2007. Meloidogyne ethiopica, a major root-knot nematode parasitizing Vitis vinifera and other crops in Chile. Nematology. 9 (5), 635-641. http://www.ingentaconnect.com/content/brill/nemy DOI:10.1163/156854107782024794

Carneiro R M D G, Gomes C B, Almeida M R A, Gomes A C M M, Martins I, 2003. First record of Meloidogyne ethiopica Whitehead, 1968 on kiwi in Brazil and reaction of different plant species. (Primeiro Registro de Meloidogyne ethiopica Whitehead, 1968, em plantas de quivi no Brasil e reação em diferentes plantas cultivadas.). Nematologia Brasileira. 27 (2), 151-158.

Carneiro R M D G, Randig O, Almeida M R A, Gomes A C M M, 2004. Additional information on Meloidogyne ethiopica Whitehead, 1968 (Tylenchida: Meloidogynidae), a root-knot nematode parasitising kiwi fruit and grape-vine from Brazil and Chile. Nematology. 6 (1), 109-123. http://www.brill.nl DOI:10.1163/156854104323072982

Castro J M da C E, Lima R D de, Carneiro R M D G, 2003. Isoenzymatic variability in Brazilian populations of Meloidogyne spp. from soybean. (Variabilidade isoenzimática de populações de Meloidogyne spp. provenientes de regiões Brasileiras produtoras de soja.). Nematologia Brasileira. 27 (1), 1-12.

Conceição I L, Tzortzakakis E A, Gomes P, Abrantes I, Cunha M J da, 2012. Detection of the root-knot nematode Meloidogyne ethiopica in Greece. European Journal of Plant Pathology. 134 (3), 451-457. http://springerlink.metapress.com/link.asp?id=100265 DOI:10.1007/s10658-012-0027-0

EPPO, 2014. PQR database., Paris, France: EPPO. http://www.eppo.int/DATABASES/pqr/pqr.htm

EPPO, 2020. EPPO Global database. In: EPPO Global database, Paris, France: EPPO.

Gomes CB, Carbonari JJ, Medina IL, Lima DL, 2005. Survey of Meloidogyne ethiopica in kiwi in Rio Grande do Sul State, Brazil, and its association with Nicotiana tabacum and Sida rhombifolia. In: Abstract of a paper presented at the 25th Congresso Brasileiro de Nematologia (Brazilian Congress of Nematology), 29 (1) Nematologia Brasileira. 114.

Lima-Medina I, Gomes CB, Nazareno NXR, 2011. Occurrence of Meloidogyne ethiopica in potato in the state of Parana. (Registro da ocorrencia de Meloidogyne ethiopica em batata no estado do Parana. Anais do 44o Congresso Brasileiro de Fitopatologia. 44th Congresso Brasileiro de Fitopatologia, Bento Goncalves-RS, 2011). In: 44th Brazilian Congress of Plant Pathology, 36 [ed. by Lavras MG]. Brazil: Brazilian Congress of Plant Pathology. 177.

Murga-Gutierrez S N, Colagiero M, Rosso L C, Sialer M M F, Ciancio A, 2012. Root-knot nematodes from asparagus and associated biological antagonists in Peru. Nematropica. 42 (1), 57-62. http://journals.fcla.edu/nematropica/article/view/79582/76900

O'Bannon JH, 1975. Nematode survey in Ethiopia., Addis Ababa, Ethiopia: Institute of Agricultural Research and FAO Rome. 29 pp.

Širca S, Urek G, Karssen G, 2004. First report of the root-knot nematode Meloidogyne ethiopica on tomato in Slovenia. Plant Disease. 88 (6), 680. DOI:10.1094/PDIS.2004.88.6.680C

Somavilla L, Gomes C B, Carbonari J J, Carneiro R M D G, 2011. Survey and characterization of root-knot nematode species in kiwi in Rio Grande do Sul State, Brazil. (Levantamento e caracterização de espécies do nematoide das galhas em quivi no Rio Grande do Sul, Brasil.). Tropical Plant Pathology. 36 (2), 89-94. http://www.scielo.br/scielo.php?script=sci_arttext&pid=S1982-56762011000200004&lng=en&nrm=iso&tlng=pt DOI:10.1590/S1982-56762011000200004

Strajnar P, Širca S, Stare B G, Urek G, 2009. Characterization of the root-knot nematode, Meloidogyne ethiopica Whitehead, 1968, from Slovenia. Russian Journal of Nematology. 17 (2), 135-142. HTTP://www.russjnematology.com

Whitehead A G, 1968. Taxonomy of Meloidogyne (Nematodea: Heteroderidae) with descriptions of four new species. Trans . zool. Soc. Lond. 31 (3), 263-401.

Whitehead A G, 1969. The distribution of root-knot nematodes (Meloidogyne spp.) in tropical Africa. Nematologica. 15 (3), 315-333.

Contributors

15/03/12 Original text by:

Regina Maria Dechechi Gomes Carneiro, Embrapa Cenargen, Parque Estação Biológica, Av. W 5 Norte (final), C. Postal 02372, 70770-900 Brasília, Brazil

Marina Dechechi Gomes Carneiro, Embrapa Cenargen, Parque Estação Biológica, Av. W 5 Norte (final), C. Postal 02372, 70770-900 Brasília, Brazil

Distribution Maps