This standard describes a diagnostic protocol for Meloidogyne enterolobii1.
This standard describes a diagnostic protocol for Meloidogyne enterolobii1.
Approved in 2011-09.
Currently, close to 100 species of root-knot nematodes have been described (Hunt & Handoo, 2009). All members are obligate endoparasites on plant roots and they occur worldwide. About 10 species are significant agricultural pests, while four are major pests and are distributed worldwide in agricultural areas, i.e. Meloidogyne incognita, M. javanica, M. arenaria and M. hapla. The root-knot nematode M. enterolobii is polyphagous and has many host plants including cultivated plants and weeds. It attacks woody as well as herbaceous plants. The species M. enterolobii is of great importance as it displays virulence against several sources of root-knot nematode-resistance genes and is considered particularly aggressive. Furthermore, a higher pathogenicity and reproductive potential was found for M. enterolobii when compared to other root-knot nematodes species such as M. incognita or M. arenaria (Kiewnick et al., 2009).
Similar to other Meloidogyne species, the second stage juveniles (J2) are attracted to the roots of a suitable host, and once they have invaded the root (usually behind the root tip) they move through the root to initiate a permanent feeding site. The feeding of the J2 on root cells induce them to differentiate into multinucleate nursing cells, called giant cells. At the same time as the giant cells are formed, the cells of the neighbouring pericycle start to divide, giving rise to a typical gall or root-knot. The root-knot juveniles can only move a few metres annually on their own, but can be spread readily through the transport of infested plants and plant products, in soil, adhering to farm equipment or by irrigation water.
Root-knot nematodes affect growth, yield, lifespan and tolerance to environmental stresses of infested plants. Typical symptoms include stunted growth, wilting, leaf yellowing and deformation of plant organs. Crop damage due to root-knot nematodes may consist of reduced quantity and quality of yield.
Meloidogyne enterolobii was first described from Hainan Island, China, in 1983. At present, this species has been recorded from Africa (Burkina Faso, Ivory Coast, Malawi, Senegal, South Africa, Togo), Asia (China, Vietnam), North America USA (State of Florida), Central America and Caribbean (Cuba, Guatemala, Martinique, Guadeloupe, Puerto Rico, Trinidad and Tobago), and South America (Brazil, Venezuela). For Europe, M. enterolobii was first recorded in France (South Brittany; Blok et al., 2002), but is currently under eradication. It has also been reported from 2 greenhouses in Switzerland (Kiewnick et al., 2008).
Updated information on geographical distribution of M. enterolobii can be viewed in the EPPO Plant Quarantine data Retrieval system (PQR) (EPPO, 2011).
Name:Meloidogyne enterolobii (Yang & Eisenback, 1983).
Synonyms:M. mayaguensis, Karssen et al. 2011, in preparation.
Taxonomic position: Nematoda: Tylenchida2 Meloidogynidae.
EPPO code: MELGMY.
Phytosanitary categorization: A2 list no.361.
Above-ground symptoms of heavily infested plants include stunting and yellowing, while below-ground typical root galls are found (Fig. 1). The root galls (knots) produced by M. enterolobii are comparable to those produced by other tropical root-knot nematode species. Particularly on tomato root-stock carrying the Meloidogyne Mi-1 resistance gene, extremely large galls can be found.
In order to identify nematodes that may be present on a commodity, it is necessary to extract specimens from the roots, soil or growing medium. If galls are found on roots, all stages of the nematode should be obtained, particularly mature swollen females (Fig. 2), males (Fig. 3) and second-stage juveniles (Fig. 4). If root-galls are not found but motile second-stage juveniles and/or males are obtained from soil, these should be distinguished from all other soil-inhabiting nematodes. Mature females can be observed within the roots by means of a dissecting microscope using transmitted light and should be extracted from roots by dissecting the tissue. They should be transferred to a 0.9% NaCl solution in order to avoid possible osmotic disruption in plain water. Alternatively, enzymatic digestion of roots and tubers with cellulase and pectinase can be used for the recovery of females and eggs (Araya & Caswell-Chen, 1993). Other stages, i.e. males and second-stage juveniles of the species, should be obtained from plant tissues or soil by suitable extraction techniques (Appendix 1).
As morphological characters of M. enterolobii are similar to other Meloidogyne species, identification to species level is based on a combination of morphological/morphometrical characters and biochemical or molecular methods (isozymes or PCR).
Differential interference phase contrast is recommended to identify specimens mounted in fixative on microscope slides. No complete key has been published on the genus Meloidogyne since Jepson (1987). This protocol presents the main morphological and morphometrical charateristics to help discrimination between similar species but as noted above identification to species level should be confirmed by biochemical or molecular methods (isozymes or PCR).
Sedentary females are annulated, pearly white and globular to pear shaped, 400–1300 μm long, 300–700 μm wide and have lateral fields each with 4 incisures. The stylet is dorsally curved, 10–25 μm long, with rounded to ovoid stylet knobs, set off to sloping posteriorly.
The males are vermiform, annulated, slightly tapering anteriorly, bluntly rounded posteriorly, 700–2000 μm long and 25–45 μm wide. The stylet is 13–30 μm long, with stylet knobs, variable in shape.
The second-stage juveniles are vermiform, annulated, tapering at both ends, 250–700 μm long, 12–18 μm wide, tail length 15–100 μm and hyaline tail part 5–30 μm in length.
The perineal pattern is round to ovoid; the arch is moderately high to high and usually rounded (Fig. 2). Phasmids are large. The stylet knobs in females are divided longitudinally by a groove so that each knob appears as two (=indented). The mean distance of the excretory pore to the anterior end in the female is 62.9 (42.3–80.6) μm. Males have a large, rounded labial disc that fuses with the medial lips to form a dorso-ventrally elongate lip region (Fig. 3). The labial disc is slightly elevated, and the medial lips are crescent shaped. The lip region is high and rounded, and only slightly set off from body. Stylet knobs of some individuals are indented but not as pronounced as in females. The distance of the pharyngeal gland orifice to stylet base is long (3.7–5.3 μm). The hemizonid is located 2–4 annules anterior to the excretory pore. Tails are short and rounded. Phasmids are small, pore-like at level of anus. The second-stage juvenile mean body length is 436.6 (405.0–472.9) μm, the hemizonid is located 1–2 annules anterior to the excretory pore and lateral lips are large and triangular in face view (Fig. 4). The anterior end is truncate. Stylet knobs are large and rounded. The tail is very thin with a broad, bluntly pointed tip. Hyaline tail terminus is clearly defined.
M. enterolobii can be separated from other described species of the genus by perineal pattern shape, male and female stylet morphology; morphology of the male; body length and morphology of the lip region, as well as tail and hyaline tail part in second-stage juveniles.
M. enterolobii differs from M. incognita by the following morphological characteristics. In the female, the stylet knobs are rounded, slightly sloping backwards and divided longitudinally by distinct grooves so that each knob appears as two. The distance of the dorsal gland orifice (DGO) to the stylet base is longer in M. enterolobii (3.7–6.2 μm) than in M. incognita (2–4 μm). The perineal pattern is usually oval shaped with the dorsal arch being moderately high to high and often rounded (Fig. 5). In males, the distance of the DGO to the stylet base is longer in M. enterolobii (3.7–5.3 μm) than in M. incognita (1.7–3.5 μm) (Fig. 6). Second-stage juveniles can be separated from M. incognita by their body length (Table 1).
|M. enterolobii1||M. incognita2||M. arenaria3||M. javanica4||M. hapla5|
|♀ stylet||13.2–18.0 (15.1)||13–16 (14)||14.4–15.8 (15.5)||14–18 (15)||10–13 (11)|
|♂ stylet||21.2–25.5 (23.4)||23.0–32.7 (25.0)||20.7–23.4 (21.6)||20.0–23.0 (21.2)||17.3–22.7 (20.0)|
|J2 body||405.0–472.9 (436.6)||337–403 (371)||450–490||387–459 (417)||312–355 (337)|
|J2 tail||41.5–63.4 (56.4)||38–55 (46)||52.2–59.9 (55.8)||36–56 (49)||33–48 (43)|
|J2 hyaline part of tail||5–15||6.3–13.5 (8.9)||10.8–19.8 (14.8)||9–18 (13.7)||11.7–18.9 (15.7)|
M. enterolobii differs from M. arenaria by juvenile body length and from M. javanica and M. hapla by male stylet length and juvenile body length (Table 1). Second-stage juveniles of M. enterolobii can be separated from M. incognita and other Meloidogyne species by the very thin and relatively long tail with its clearly defined hyaline tail terminus, the posterior part of the hyaline part is running straight and parallel (Fig. 7); however, demarcation of the hyaline tail part is not as clear as for M. chitwoodi and M. fallax (see PM 7/41 for pictures for M. chitwoodi and M. fallax).
As the range of values for each of these characteristics can overlap between species, care is needed. Confirmation with isozyme electrophoresis or molecular techniques is required. Morphometrical characteristics for M. chitwoodi and M. fallax are not presented in Table 1 as these 2 species are usually not associated with M. enterolobii. In case of doubt, refer to PM 7/41 Diagnostic protocol for Meloidogyne chitwoodi and Meloidogyne fallax.
A useful method for identification of females of several Meloidogyne species by isozyme electrophoresis has been developed (Esbenshade & Triantaphyllou, 1985). Esterase (EC 184.108.40.206) and malate dehydrogenase (EC 220.127.116.11) isozyme patterns discriminate M. enterolobii, M. incognita, M. arenaria, M. javanica and M. hapla (Xu et al., 2004). The test is described in Appendix 2.
The following PCR methods can be recommended for molecular identification:
PCR methods can be applied to all developmental stages of nematodes.
A molecular diagnostic key for the identification of single juveniles of 7 common and economically important species of root-knot nematode (Meloidogyne spp.) has been published by Adam et al. (2007).
Reference material can be obtained at the Plant Protection Service, Wageningen (NL).
Guidelines on reporting and documentation are given in EPPO Standard PM7/77 (1) ‘Documentation and reporting on a diagnosis’
Further information on this organism can be obtained from G. Karssen, PPS Wageningen, NL; S. Kiewnick, ACW Wädenswil, CH.
Recent developments combining a classification based on morphological data and molecular analysis refer to ‘Tylenchomorpha’ (De Ley & Blaxter, 2004).
Note: although no validation data are available for these methods they are used in different laboratories and were considered as reliable by the EPPO ad hoc Panel on Nematodes.
This protocol was originally drafted by: S. Kiewnick (Agroscope Changins-Wädenswil, CH) and J. Hallmann (Julius Kühn-Institut, Braunschweig, DE).
If you have any feedback concerning this Diagnostic Protocol, or any of the tests included, or if you can provide additional validation data for tests included in this protocol that you wish to share please contact email@example.com.
From roots of plants that are suspected to be infested, second-stage juveniles of the nematode can be extracted by the incubation method: roots are carefully washed free of adhering soil and cut into small pieces. These root pieces are then transferred to a moist filter paper which is supported by an extraction sieve and placed in a shallow dish with a thin layer of water. After incubation for 7–14 days at 20°C, most of the juveniles will have hatched from the eggs, egress from root tissues and move through the filter paper into the water. To collect the nematodes, the water is poured into a counting dish and examined at 25–35× magnification using a dissecting microscope.
A faster extraction method is the maceration/centrifugal/flotation method (Hooper et al., 2005): the roots are homogenized in an electric macerator at about 12 600 rev min−1 for 30 s. The suspension is poured onto a sieve of 1200 μm pore size, washed and the water passing through the sieve is collected, mixed with 1% kaolin powder and centrifuged for 4 min at 1500 g. The supernatant is poured off and the residue re-suspended in a sucrose, MgSO4 or ZnSO4 solution of specific gravity 1.18 and centrifuged again at 1500 g for 4 min. The supernatant is poured through a sieve of 5 to 20-μm pore size. Nematodes can be transferred to a glass dish for examination under a microscope at 25–35× magnification.
To extract males and second-stage juveniles from small amounts of soil, a thin layer of soil (2–3 mm) is placed on a paper filter laid over a coarse nylon sieve. The sieve is placed in a Baermann funnel (a glass funnel with a piece of rubber tubing attached to the stem and closed with a clamp) filled with water so that the soil is just wetted. After 48 h, nematodes have emerged from the soil and moved to the bottom of the tube. By releasing a small amount of water from the base of the funnel into a glass dish, nematodes can be collected and examined under a dissecting microscope at 25–35× magnification.
For larger amounts of soil, a simple flotation/sieving technique may be used: 100–500 mL of soil is added to a 10 L-bucket of water. The soil particles are suspended in the water by stirring vigorously for 10 s, and then allowed to settle for a further 45 s. The supernatant is poured through a bank of 3 sieves of 50-μm pore size. The soil debris collected on the sieves is washed, collected in a beaker, and poured onto a Baermann funnel (as described above). After 24 h, nematodes can be collected by releasing a small amount of water from the base of the funnel into a glass dish. This can then be examined under a dissecting microscope at 25–35×. This simple method can extract nematodes for identification but is not as efficient, in terms of numbers of nematodes recovered, as more complex methods such as the Oostenbrink and Seinhorst elutriators (Hooper et al., 2005). The maceration/centrifugal/flotation method of Coolen (1979) may be used as described above (first with water and kaolin, then with MgSO4, ZnSO4 or sucrose, solution).
Sample appl. Down at step 3.2 0 Vh.
Sample appl. Up at step 3.3 0 Vh.
Sep. 3.1 400 V, 10 mA, 2.5 W, 10°C, 10 Vh.
Sep. 3.2 400 V, 1 mA, 2.5 W, 10°C, 2 Vh.
Sep. 3.3 400 V, 10 mA, 2.5 W, 10°C, 125 Vh.
|0.1 M Phosphate buffer, pH 7.3||100 mL|
|Fast Blue RR salt||0.06 g|
|Alpha-Naphthyl acetate (dissolved in 2 mL acetone)||0.04 g|
|Nitro Blue Tetrazolium||0.03 g|
|Phenazine Methosulfate||0.002 g|
|0.5 M Tris, pH 7.1||5 mL|
|Reagent-grade water||70 mL|
The species-specific phenotype J3 of M. javanica with Rm values of 1.0, 1.25 and 1.4 (Fig. 8) should be used as a standard control in each gel. Phenotype M2 (=VS1-S1; Esbenshade & Triantaphyllou, 1985) with 2 major bands (Rm 0.7 and 0.9) was described for M. enterolobii with 2 different populations tested. Carneiro et al. (2000) mentioned that occasionally one of these bands resolved in 2 minor bands (Rm: 0.75, 0.95).
The phenotype N1 of M. javanica with the Rm value of 1.0 (Fig. 9) should be used as a standard control in each gel. Phenotype N1a with one major band (Rm 1.4) was described for M. enterolobii with 2 different populations tested (Esbenshade & Triantaphyllou, 1985; Carneiro et al., 2000). Phenotype H1 (Rm 1.9) is species specific for M. hapla with one major band.
As an alternative (or in addition to) to the external positive controls (PIC and PAC), internal positive controls can be used to monitor each individual sample separately. These can include: co-amplification of endogenous nucleic acid, using conserved primers that amplify conserved non-target nucleic acid that is also present in the sample (e.g. plant cytochrome oxidase gene or bacterial 18S rDNA) amplification of samples spiked with exogeneous nucleic acid that has no relation to the target nucleic acid (e.g. synthetic internal amplification controls) or amplification of a duplicate sample spiked with the target nucleic acid.
Wishart et al. (2002) Ribosomal Intergenic Spacer: a polymerase chain reaction diagnostic for Meloidogyne chitwoodi, M. fallax, and M. hapla.
As an alternative (or in addition) to the external positive controls (PIC and PAC), internal positive controls can be used to monitor each individual sample separately. These can include: co-amplification of endogenous nucleic acid, using conserved primers that amplify conserved non-target nucleic acid that is also present in the sample (e.g. plant cytochrome oxidase gene or bacterial 18S rDNA) amplification of samples spiked with exogeneous nucleic acid that has no relation with the target nucleic acid (e.g. synthetic internal amplification controls) or amplification of a duplicate sample spiked with the target nucleic acid.