Meloidogyne enterolobii


  • Use of names of chemicals or equipment in these EPPO Standards implies no approval of them to the exclusion of others that may also be suitable.

Abstract

Specific scope

This standard describes a diagnostic protocol for Meloidogyne enterolobii1.

Specific approval and amendment

Approved in 2011-09.

Introduction

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).

Identity

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.

Detection

Symptoms

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.

Figure 1.

 Cucumber root system damaged by Meloidogyne enterolobii (Courtesy: Agroscope Changins-Wädenswil, CH).

Extraction procedure

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).

Figure 2.

Meloidogyne enterolobii. Female: (A) Anterior end. (B–F) Entire. (G–I) Perineal pattern (Yang & Eisenback, 1983).

Figure 3.

Meloidogyne enterolobii. Male: (A) Pharyngeal region. (B–C) Lip region. (D) Lateral field near mid-body. (E) Tail (Yang & Eisenback, 1983).

Figure 4.

Meloidogyne enterolobii. Second-stage juveniles: (A) Pharyngeal region. (B) Lip region (lateral). (C) Lip region (ventral). (D) Lateral field near mid-body. (E–F) Tail (lateral). (G) Tail (ventral) (Yang & Eisenback, 1983).

Identification

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).

Morphological characteristics

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).

Morphological characteristics of Meloidogyne spp.

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.

Morphological characteristics of the species M. enterolobii(after Yang & Eisenback, 1983).

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.

Possible confusion with similar species

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).

Figure 5.

 Drawings of perineal patterns of Meloidogyne enterolobii, M. incognita, M. arenaria, M. javanica and M. hapla (after 1Yang & Eisenback, 1983; 2Williams, 1973; 3Williams, 1975; 4Williams, 1972; 5Williams, 1974), different drawings illustrate the variability.

Figure 6.

 Drawings of male lip regions of Meloidogyne enterolobii, M. incognita, M. arenaria, M. javanica and M. hapla (after 1Yang & Eisenback, 1983; 2Williams, 1973; 3Williams, 1975; 4Williams, 1972; 5Williams, 1974), different drawings illustrate the variability.

Table 1.   Morphological and morphometric variations between Meloidogyne enterolobii, M. incognita, M. arenaria, M. javanica and M. hapla (length in μm)
 M. enterolobii1M. incognita2M. arenaria3M. javanica4M. hapla5
  1. 1Yang & Eisenback (1983); 2Williams (1973); 3Williams (1975); 4Williams (1972); 5Williams (1974).

♀ stylet13.2–18.0 (15.1)13–16 (14)14.4–15.8 (15.5)14–18 (15)10–13 (11)
♂ stylet21.2–25.5 (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 body405.0–472.9 (436.6)337–403 (371)450–490387–459 (417)312–355 (337)
J2 tail41.5–63.4 (56.4)38–55 (46)52.2–59.9 (55.8)36–56 (49)33–48 (43)
J2 hyaline part of tail5–156.3–13.5 (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).

Figure 7.

 Drawings of the tail of secondary-stage juveniles of Meloidogyne enterolobii, M. incognita, M. arenaria, M. javanica and M. hapla (after 1Yang & Eisenback, 1983; 2Williams, 1973; 3Williams, 1975; 4Williams, 1972; 5Williams, 1974), different drawings illustrate the variability.

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.

Isozyme electrophoresis

A useful method for identification of females of several Meloidogyne species by isozyme electrophoresis has been developed (Esbenshade & Triantaphyllou, 1985). Esterase (EC 3.1.1.1) and malate dehydrogenase (EC 1.1.1.37) 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.

Molecular methods

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

Reference material can be obtained at the Plant Protection Service, Wageningen (NL).

Reporting and documentation

Guidelines on reporting and documentation are given in EPPO Standard PM7/77 (1) ‘Documentation and reporting on a diagnosis

Further information

Further information on this organism can be obtained from G. Karssen, PPS Wageningen, NL; S. Kiewnick, ACW Wädenswil, CH.

Footnotes

  • 2

    Recent developments combining a classification based on morphological data and molecular analysis refer to ‘Tylenchomorpha’ (De Ley & Blaxter, 2004).

  • 3

    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.

Acknowledgements

This protocol was originally drafted by: S. Kiewnick (Agroscope Changins-Wädenswil, CH) and J. Hallmann (Julius Kühn-Institut, Braunschweig, DE).

Feedback on this Diagnostic Protocol

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 diagnostics@eppo.fr.

Appendices

Appendix 1 – Extraction of motile stages of Meloidogyne enterolobii from roots and soil3

Extraction from roots

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.

Extraction from soil

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).

Appendix 2 – Isozyme electrophoresis for identification of Meloidogyne enterolobii

1. General information

  • 1.1 Several reliable isozyme electrophoresis methods are available for the identification of single young egg-laying Meloidogyne females. The following method was originally developed by Esbenshade & Triantaphyllou (1985) and modified and adapted for PhastSystem, i.e. an automated electrophorectic apparatus, by Karssen et al. 1995. It is possible to run 2 gels a time with a total of 24 females per electrophoresis run. The applied method uses native gradient polyacrylamide gel electrophoresis in a discontinuous buffer system.
  • 1.2 The PhastSystem apparatus, prefabricated gels and the sample well stamps are manufactured by Amersham Electrophoresis and sold by GE Healthcare.

2. Samples

  • 2.1 Root-knot nematode infested roots are placed in 0.9% NaCl solution; young egg-laying females are isolated under a dissecting microscope and placed in 0.9% NaCl solution on ice or stored in a freezer at −20°C.
  • 2.2 Before electrophoresis, the females are transferred from the NaCl solution to reagent-grade water on ice for a few minutes for desalting.

3. Sample preparation

  • 3.1 After desalting a sample well stamp (on ice) with 12 wells is filled with 1 female per well.
  • 3.2 The 2 middle wells are each filled with a reference material (preferably Meloidogyne javanica) female.
  • 3.3 To each well 0.6 μL extraction buffer is added (20% sucrose, 2% Triton X-100 and 0.01% Bromophenol Blue).
  • 3.4 The females are carefully macerated with a small glass rod and loaded on two 12/03 sample applicators (0.3 μL per well).
  • 3.5 Both applicators are inserted at the cathode slot into the left and right applicator arm.

4. Electrophoresis

  • 4.1 Before electrophoresis 2 PhastGel gradient gels (8–25) are placed on the gel-bed and pre-cooled at 10°C.
  • 4.2 The following adapted program is used for electrophoresis:
  • 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.

  • 4.3 After adding the sample applicators the programme can be started.
  • 4.4 The gels are placed after electrophoresis in a Petri dish for staining.

5. Staining

  • 5.1 One gel is stained for esterase (EST, EC 3.1.1.1) activity, the other for malate dehydrogenase (MDH, EC 1.1.1.37).
  • 5.2 Staining solutions are prepared according to Table 2.
  • 5.3 Staining solution is added to each Petri dish with gel and placed in an incubator at 37°C.
  • 5.4 The total staining times for EST and MDH are 60 min and 5 min, respectively.
Table 2.   Esterase (EST) and malate dehydrogenase (MDH) staining solutions
  1. *10.6 g Na2CO3 + 1.34 g L-Malic acid in 100 mL water.

EST
0.1 M Phosphate buffer, pH 7.3100 mL
Fast Blue RR salt0.06 g
EDTA0.03 g
Alpha-Naphthyl acetate (dissolved in 2 mL acetone)0.04 g
MDH
Beta-NAD0.05 g
Nitro Blue Tetrazolium0.03 g
Phenazine Methosulfate0.002 g
0.5 M Tris, pH 7.15 mL
Stock*7.5 mL
Reagent-grade water70 mL

6. Results

Figure 8.

 Esterase (EST) phenotypes of Meloidogyne species. (Redrawn after Carneiro et al., 2000) Esterase types: J3 = M. javanica (Rm 1.0, 1.25, 1.4); I1 = M. incognita races 2 and 3 (Rm 1.0); I2 = M. incognita races 1 and 4 (Rm 1.0, 1.1); A3, A2 = M. arenaria race 2 (Rm 1.1, 1.2, 1.3 and Rm 1.2, 1.3); M2 = M. enterolobii (Rm 0.7, 0.9; occasionally 0.75, 0.95); H1 = M. hapla (Rm 1.1).

Figure 9.

 Phenotypes of malate dehydrogenase (MDH) in Meloidogyne species. (Redrawn after Carneiro et al., 2000). N1 = M. javanica, M. incognita and M. arenaria (Rm 1.0); H1 = M. hapla (Rm 1.9); N1a = M. enterolobii (Rm 1.4).

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.

Appendix 3 – PCR method based on mitochondrial DNA differences for identification of Meloidogyne enterolobii

1. General information

  •  1.1 Protocol developed by Blok et al. (2002).
  •  1.2 Nematodes (second-stage juveniles, females, eggs, males) are extracted from the roots or from soil as described in Appendix 1. Specimens are collected from the extract and placed in 50 μL sterile water.
  •  1.3 The targeted region is a 63 bp repeat in the mitochondrial DNA.
  •  1.4 The amplicon size is 301 bp (including primer sequence)
  •  1.5 Oligonucleotides are used at a final concentration of 20 μM each.
  •  1.6 Forward primer 63VNL, 5′-GAAATTGCTTTATTGTTA CTAAG-3′, Reverse primer 63VTH, 5′-TAGCCACAG CAAAATAGTTTTC-3′.
  •  1.7 One unit Taq DNA Polymerase (Roche) used per reaction (50 μL).
  •  1.8 Nucleotide concentration is 0.2 mM each.
  •  1.9 Reaction buffer: 10 mM Tris–HCL (pH 8.3), 1.5 mM MgCl2, 50 mM KCl.
  • 1.10 Molecular grade water (MGW).
  • 1.11 Perkin-Elmer 480 thermal cycler (Perkin Elmer Applied Biosystems).

2. Methods

  • 2.1 Nucleic acid extraction and purification
    • 2.1.1 DNA is extracted from either 50 μL packed-cell volume followed by standard phenol: chloroform extraction (Blok et al., 1997) or from females ground in 30 μL MGW water.
    • 2.1.2 DNA is stored in MGW at −20°C.
    • 2.1.3 Alternatively, single second-stage juveniles, females or males are transferred to 0.2 mL polymerase chain reaction (PCR) tube containing 25 μL of sterile water. An equal volume of lysis buffer containing 0.2 M NaCl, 0.2 M Tris-HCl (pH 8.0), 1% (vol/vol) β-mercaptoethanol, and 800 μg mL−1 of proteinase K is added. Lysis takes place in a Thermomixer (Eppendorf, Hamburg, Germany) at 65°C at 750 rpm for 2 h followed by a 5 min incubation at 100°C (Holterman et al., 2006). This protocol is routinely used in Switzerland.
    • 2.1.4 Lysate can be used immediately or stored at −20°C.
  • 2.2 Polymerase Chain Reaction – PCR.
    • 2.2.1 Total reaction volume of a single PCR reaction is 50 μL.
    • 2.2.2 1× PCR buffer.
    • 2.2.3 2 nM dNTPs.
    • 2.2.4 One unit of Taq polymerase.
    • 2.2.5 20 μM forward primer.
    • 2.2.6 20 μM reverse primer.
    • 2.2.7 10 ng DNA.
    • 2.2.8 PCR grade water is added to 50 μL.
    • 2.2.9 PCR cycling parameters 35 cycles of 94°C for 40 s, 50°C for 30 s and 72°C for 90 s followed by 72°C for 5 min.

3. Essential procedural information

  • 3.1 DNA fragments are separated by electrophoresis on agarose gel and visualized under UV light according to standard procedures (e.g. Sambrook et al., 1989).
  • 3.2 For a reliable test result to be obtained, the following (external) controls should be included for each series of nucleic acid isolation and amplification of the target organism and target nucleic acid, respectively:
    •  Positive isolation control (PIC) to ensure that nucleic acid of sufficient quantity and quality is isolated: nucleic acid extraction and subsequent amplification of the target organism or a sample that contains the target organism.
    •  Negative amplification control (NAC) to rule out false positives due to contamination during the preparation of the reaction mix: amplification of PCR grade water that was used to prepare the reaction mix.
    •  >Positive amplification control (PAC) to monitor the efficiency of the amplification: amplification of nucleic acid of the target organism. This can include nucleic acid extracted from the target organism, total nucleic acid extracted from infected host tissue, whole genome amplified DNA or a synthetic control (e.g. cloned PCR product).

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.

  • 3.3 Interpretation of results: in order to assign results from a PCR-based test the following criteria should be followed:
    •  A PCR test will be considered positive if it produces amplicons of 301 bp and provided that the contamination controls (NAC) are negative.
    •  A PCR test will be considered negative, if it produces no band or no band of a similar size and provided that the test and extraction inhibition controls (PIC and PAC) are positive.
    •  Tests should be repeated if any contradictory or unclear results are obtained.

4. Performance criteria (based on experience in Agroscope Changins-Wädenswil, CH)

  • 4.1 Sensitivity is 1 individual (female, juvenile, male).
  • 4.2 Specificity is high. No cross reactions were found with the closely related species M. incognita, M. javanica, M. arenaria and M. hapla.
  • 4.3 Repeatability is high (100%).
  • 4.4 Reproducibility is high (100%).

Appendix – 4

Wishart et al. (2002) Ribosomal Intergenic Spacer: a polymerase chain reaction diagnostic for Meloidogyne chitwoodi, M. fallax, and M. hapla.

1. General information

  •  1.1 Multiplex PCR test developed by Wishart et al., 2002 for identification of M. chitwoodi, M. fallax and M. hapla [See also PM 7/41 (2)]:
  •  1.2 Nematodes (second-stage juveniles, females, eggs, males) are extracted from roots and soil as described in Appendix 1. Nematodes are collected and stored in water.
  •  1.3 The assay is designed to the intergenic spacer (IGS) region of the rDNA sequences of Meloidogyne spp.
  •  1.4 The amplicon size is 615 bp (including primer sequence)
  •  1.5 Oligonucleotides are used with 0.5 per 25 μL reaction of a 10 μM concentration.
  •  1.6 Forward primer JMV1, 5′-GGATGGCGTGCTTTCAAC-3′), Reverse primer JMVtropical, 5′-GCKGGTAATTAA GCTGTCA-3′.
  •  1.7 One unit Taq DNA Polymerase (Promega) used per reaction (25 μL).
  •  1.8 Nucleotide concentration is 2 mM each.
  •  1.9 Reaction buffer.
  • 1.10 Molecular grade water (MGW).
  • 1.11 Applied BiosystemsThermocycler 9700.

2. Methods

  • 2.1 Nucleic acid extraction and purification
    • 2.1.1 DNA can be extracted from single second-stage juveniles, females or males by transferring an individual specimen to 0.2 mL polymerase chain reaction (PCR) tubes containing 25 μL of sterile water. An equal volume of lysis buffer containing 0.2 M NaCl, 0.2 M Tris–HCl (pH 8.0), 1% (vol/vol) β-mercaptoethanol, and 800 μg mL−1 of proteinase K is added. Lysis takes place in a Thermomixer (Eppendorf, Hamburg, Germany) at 65°C at 750 rpm for 2 h followed by a 5 min incubation at 100°C (Holterman et al., 2006).
    • 2.1.2 Lysate can be used immediately or stored at −20°C.
  • 2.2 Polymerase Chain Reaction – PCR.
    • 2.2.1 Total reaction volume of a single PCR reaction is 25 μL.
    • 2.2.2 Reaction buffer: 10 mM Tris–HCL (pH 9.0), 1% Triton X-100, 1.5 mM MgCl2, 5 mM KCl, 2 nM dNTPs.
    • 2.2.3 One unit of Taq polymerase.
    • 2.2.4 0.5 μL of a 10 μM forward primer.
    • 2.2.5 0.5 μL of a 10 μM reverse primer.
    • 2.2.6 0.5 μL DNA extract.
    • 2.2.7 MGW is added to 25 μL.
    • 2.2.8 PCR cycling parameters 40 cycles of 94°C for 30 s, 55°C for 30 s and 72°C for 90 s followed by 72°C for 10 min.

3. Essential procedural information

  • 3.1 DNA fragments are separated by electrophoresis on agarose gel and visualized under UV light according to standard procedures (e.g. Sambrook et al., 1989).
  • 3.2 For a reliable test result to be obtained, the following (external) controls should be included for each series of nucleic acid isolation and amplification of the target organism and target nucleic acid, respectively:
    •  Positive isolation control (PIC) to ensure that nucleic acid of sufficient quantity and quality is isolated: nucleic acid extraction and subsequent amplification of the target organism or a sample that contains the target organism (e.g. naturally infected host tissue or host tissue spiked with the target organism).
    •  Negative amplification control (NAC) to rule out false positives due to contamination during the preparation of the reaction mix: amplification of PCR grade water that was used to prepare the reaction mix.
    •  Positive amplification control (PAC) to monitor the efficiency of the amplification: amplification of nucleic acid of the target organism. This can include nucleic acid extracted from the target organism, total nucleic acid extracted from infected host tissue, whole genome amplified DNA or a synthetic control (e.g. cloned PCR product).

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.

  • 3.3 Interpretation of results: in order to assign results from a PCR-based test the following criteria should be followed:
    •  A PCR test will be considered positive if it produces amplicons of 615 bp and provided that the contamination controls (NAC) are negative.
    •  A PCR test will be considered negative if it produces no band or no band of a similar size and provided that the test and extraction inhibition controls (PIC and PAC) are positive.
    •  Tests should be repeated if any contradictory or unclear results are obtained.

4. Performance criteria [based on a validation performed in the French Plant Health Laboratory (G. Anthoine, pers. comm. 2011)]

  • 4.1  Sensitivity is 1 individual (female, juvenile, male).
  • 4.2 Specificity 100%. No cross reactions were found with the closely related species M. incognita, M. javanica and M. chitwoodi, M. fallax and M. hapla. No species or population with the M. enterolobii profile, except M. enterolobii populations. Tropical populations/species gave specific amplification profile.
  • 4.3 Repeatability: 1 individual 75%, more than 2 individuals 100%.
  • 4.4 Reproducibility: 2 individuals 100% (24 positive repetitions), 1 individual 83% (20 positive repetitions out of 24).

Ancillary