Meloidogyne chitwoodi and Meloidogyne fallax
2004; Wiley; Volume: 34; Issue: 2 Linguagem: Galês
10.1111/j.1365-2338.2004.00735.x
ISSN1365-2338
Tópico(s)Legume Nitrogen Fixing Symbiosis
ResumoEPPO BulletinVolume 34, Issue 2 p. 315-320 Diagnostic protocols for regulated pests†Free Access Meloidogyne chitwoodi and Meloidogyne fallax First published: 10 September 2004 https://doi.org/10.1111/j.1365-2338.2004.00735.xCitations: 6 European and Mediterranean Plant Protection Organization PM 7/41(1) Organisation Européenne et Méditerranéenne pour la Protection des Plantes AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat Specific scope This standard describes a diagnostic protocol for Meloidogyne chitwoodi and Meloidogyne fallax. Specific approval and amendment This Standard was developed under the EU DIAGPRO Project (SMT 4-CT98-2252) by partnership of contractor laboratories and intercomparison laboratories in European countries. Approved as an EPPO Standard in 2003-09. Introduction At present more than 90 species of root-knot nematode have been described. All members are obligate endoparasitic pathogens on plant roots and they are detected worldwide. About 10 species are agricultural pests, while four are major pests and distributed worldwide in agricultural areas. The root-knot nematodes Meloidogyne chitwoodi and Meloidogyne fallax parasitize monocotyledons and dicotyledons, including several crop plants such as potatoes, carrots and tomatoes (Santo et al., 1980; O’Bannon et al., 1982; Brinkman et al., 1996; Karssen, 2002). The juveniles of the second stage are attracted to the roots and penetrate the roots closely behind the root tip, where they enter the vascular cylinder. The parasites finally start feeding on cells, which are rapidly turned into multinucleated 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 nematodes are able to move only a few metres annually on their own, but they can be spread readily through the transport of infested plants and plant products, in soil, adhering to farm implements and in irrigation water. Root-knot nematodes affect root growth, yield and quality of their hosts. The above-ground symptoms are not readily apparent, but they may consist of various degrees of stunting, lack of vigour, and wilting under moisture stress. Hosts may be heavily infected without showing external symptoms on the harvested products, for example symptomless potato tubers. M. chitwoodi was described from the Pacific North-west region of the USA in 1980. At present this species has been recorded from Argentina, Belgium, Germany, Netherlands, Portugal, USA, Mexico and South Africa. M. fallax was detected for the first time in 1992 in a field plot experiment one mile north of Baexem (NL) and initially remarked as a deviating M. chitwoodi population (Karssen, 1994). After the first report, it was recorded on potato at several locations in the southern and south-eastern part of the Netherlands (Karssen, 1996a), close to the German and Belgium borderland. It was also found in a plastic house in France (Daher et al., 1996). It has so far been detected outside Europe in New Zealand, Australia and South Africa (Marshall et al., 2001; Nobbs et al., 2001; Fourie et al., 2001). Identity Name: Meloidogyne chitwoodi Golden et al. 1980 Taxonomic position: Nematoda: Heteroderidae: Meloidogyninae Bayer computer code: MELGCH Phytosanitary categorization: EPPO A2 list n°227, EU Annex designation I/A2 Name: Meloidogyne fallax Karssen 1996 Taxonomic position: Nematoda: Heteroderidae: Meloidogyninae Bayer computer code: MELGFA Phytosanitary categorization: EPPO A2 list n°295, EU Annex designation I/A2 Detection Above-ground symptoms of heavily infested plants include stunting and yellowing, while below ground galling is typical (Web Fig. 1). The root galls produced by M. chitwoodi and M. fallax are comparable to those produced by several other root-knot species, i.e. relatively small galls in general without secondary roots emerging from them (as in M. hapla). On potato tubers, M. chitwoodi and M. fallax cause numerous small pimple-like raised areas on the surface (in M. hapla these swellings are not evident). Some potato cultivars, although heavily infected, may be free from visible external symptoms, while the internal potato tissue is necrotic and brownish, just below the peel. Identification In order to identify nematodes that may be present on a commodity, it is necessary to extract specimens from the roots, potato tubers or from soil or growing medium (Fig. 2). Mature females can be observed within the roots by means of a dissecting microscope using transmitted light. Identification to species level is based on a combination of morphological/morphometrical characters and biochemical or molecular methods (isozymes or PCR). For light microscope identification, it is recommended to examine specimens mounted in fixative on microscope slides (Appendix 1). Figure 2Open in figure viewerPowerPoint Decision scheme for the detection and identification of Meloidogyne chitwoodi and M. fallax. Extraction procedure Mature females can be extracted by dissecting apart the tissues but then should be stored in a 0.9% solution of NaCl 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 other developmental stages (Araya & Caswell-Chen, 1993). Other stages, i.e. males (Web Fig. 3) and second-stage juveniles of the species, should be obtained from plant tissues or soil by suitable extraction techniques (Appendix 2). Morphology Sedentary females are thin, annulated, pearly white and globular to pear shaped, 400–1300 µm long and 300–700 µm wide. The stylet is dorsally curved, 10–25 µm long, with rounded to ovoid stylet knobs, set off to sloping posteriorly. The non-sedentary males are vermiform, annulated, slightly tapering anteriorly, bluntly rounded posteriorly, 700– 2000 µm long and 25–45 µm width. The stylet is 13–30 µm long, with stylet knobs, variable in shape. The non-sedentary second-stage juveniles are vermiform, annulated, tapering at both ends, 250–700 µm long, 12–18 µm width, tail length 15–100 µm and hyaline tail part 5–30 µm in length. Morphological and morphometrical differences between M. chitwoodi and M. fallax were noticed when the two species were cultivated on the same host under comparable conditions (Karssen, 1995, 1996a). The most striking differences for males and females are shown in Table 1. With the scanning electron microscope, it was observed that the male head of M. fallax has an elevated labial disk. Secondary differences were recorded in the female perineal pattern (M. fallax: relatively higher dorsal arch and thicker striae) and second-stage juvenile hemizonid position to the excretory pore (same level for M. fallax, anterior adjacent for M. chitwoodi). Based on morphology, the two species are closely related, and this misleading morphological resemblance to M. chitwoodi was the reason for the name given to M. fallax (Petersen & Vrain 1996). Web Figs 4–7 presents some drawings of different stages of M. chitwoodi and M. fallax. However, for identification, molecular techniques should be used in adadtion to morphology & morphometrics (Appendix 3). Table 1. Morphological and morphometrical differences between Meloidogyne chitwoodi and M. fallax (µm) M. fallax M. chitwoodi Female stylet length 13.9–14.5 10.7–13.3 Male stylet length 18.1–20.8 15.8–18.3 Male stylet knob shape prominent and rounded small and irregular J2 body length 368–410 336–385 J2 tail length 46.1–55.6 39.2–44.9 J2 hyaline tail length 12.1–15.8 8.2–12.6 Biochemical techniques Esbenshade & Triantaphyllou (1985) have described a useful method for identification of females of several Meloidogyne species (including M. chitwoodi) by isozyme electrophoresis. Esterase (EC 3.1.1.1) and malate dehydrogenase (EC 1.1.1.37) isozyme patterns discriminated M. fallax and M. chitwoodi females (Karssen et al., 1995; Karssen, 1996b). Additionally, isozymes of glucose 6-phosphate dehydrogenase (EC 1.1.1.49) are useful to differentiate the two species (van der Beek & Karssen, 1997). van der Beek et al. (1997) and Tastet et al. (1999) used two-dimensional gel electrophoresis to study total soluble protein patterns of M. hapla, M. chitwoodi and M. fallax, and confirmed that these species were distinct. Later, Tastet et al. (2001) produced antisera to discriminate M. chitwoodi and M. fallax from other root-knot nematodes, using a combination of two-dimensional gel electrophoresis, internal amino-acid sequencing and serology. Another method for identification of Meloidogyne species is the polymerase chain reaction (PCR). Zijlstra et al. (1995) used PCR amplification and restriction fragment length polymorphisms (RFLPs) of the internal transcribed spacer (ITS) of ribosomal DNA (rDNA), and recorded distinct differences between M. hapla and M. chitwoodi. Three M. chitwoodi isolates (all B-types or M. fallax), could be distinguished from M. chitwoodi. This method is also useful to differentiate mixtures of root-knot nematodes including M. chitwoodi and M. fallax (Zijlstra et al., 1997) without the need for subsequent enzyme digestion (Zijlstra, 1997). Peterson & Vrain (1996) have described a rapid PCR identification method for M. hapla, M. chitwoodi and M. fallax based on amplification of the rDNA intergenic spacer (IGS), without the need for restriction enzyme digestion. Also species-specific primers were developed for M. chitwoodi and M. fallax, based on unique sequences within ribosomal IGS (Peterson et al., 1997) or designed from species-specific RAPD fragments (Zijlstra, 2000). Recently, satellite DNA probes and AFLPs have also been successfully applied to separate M. chitwoodi from M. fallax (Castagnone-Serone et al., 1998, 1999; van der Beek et al., 1998). Castagnone-Serone (2000) reviews molecular tools for the identification of these nematodes. Waeyenberge & Moens (2001) used RAPD to examine the molecular diversity between populations of M. fallax and M. chitwoodi. Possible confusion with similar species M. hapla differs from M. chitwoodi and M. fallax in (female) perineal pattern (rounded with low dorsal arch and lateral lines; punctations present in the tail terminal area); male and female stylet knobs (small rounded and set off from the shaft); male head shape (head region not in contour with the first body annule, but slightly wider); second-stage juvenile tail shape (long, slender tail with a narrow tapering terminus, tip finely rounded to pointed, hyaline terminus not clearly demarcated). Jepson (1987) and Karssen (2002) provide a comparison with other Meloidogyne species and Siddiqi (2000) a comparison with other genera of Tylenchida. Requirements for a positive diagnosis The procedures for detection and identification described in this protocol, and the decision scheme in Fig. 2, should have been followed. If root knots are found on roots, all stages of the nematode should be obtained, particularly mature swollen females, males and 2nd stage juveniles. These should have the characters described above for Meloidogyne chitwoodi or M. fallax. If root-knots are not found but motile nematode stages are obtained from soil (particularly second-stage juveniles), these should be distinguished from all other soil inhabiting nematodes. Report on the diagnosis A report on the execution of the protocol should include: • information and documentation on the origin of the infested material • a description of the disease symptoms • the number of individuals examined drawings or photographs of the following morphological features: tail of second-stage juvenile; male head, including stylet; female perineal • pattern measurements of the morphological features given in the description above • applied molecular and/or isozyme method (Appendix 3) • molecular and/or isozyme results • comments as appropriate on the certainty or uncertainty of the identification. Slides of preserved specimens should be kept. Further information Further information on this organism can be obtained from: L. den Nijs & G. Karssen, Plant Protection Service, Diagnostic Centre, PO Box 9102, 6700 HC Wageningen, the Netherlands. E-mail: L.J.M.F.den.Nijs@pd.agro.nl Footnotes 1 The Figures in this Standard marked ‘Web Fig.’ are published on the EPPO website http://www.eppo.org. 2 L. De Wael (Centrum voor Landbouwkundig Onderzoek, BE); H. Marzin (LNPV Unité de Nématologie Rennes, FR); M. Aalten (CSL, GB); S. Turner (Agriculture and Food Science Centre, Belfast, GB); J. Luimes (NAK, NL); J. Pickup (Scottish Agricultural Science Agency, Edinburgh, GB), SLU (Vaxtvetenskap/Nematologi, SE); D. Heinicke (Pflanzenschutzamt Hannover, DE). Acknowledgements This protocol was originally drafted by: G. Karssen & L. den Nijs, Plant Protection Service, Wageningen (NL). This protocol was ring-tested in different European laboratories22 L. De Wael (Centrum voor Landbouwkundig Onderzoek, BE); H. Marzin (LNPV Unité de Nématologie Rennes, FR); M. Aalten (CSL, GB); S. Turner (Agriculture and Food Science Centre, Belfast, GB); J. Luimes (NAK, NL); J. Pickup (Scottish Agricultural Science Agency, Edinburgh, GB), SLU (Vaxtvetenskap/Nematologi, SE); D. Heinicke (Pflanzenschutzamt Hannover, DE). . Appendices Appendix 1 Preparation of nematode specimens for microscope examination For identification to species level of nematodes recovered from plant material or soil, it is necessary to examine specimens mounted in fixative on microscope slides. A good fixative for this purpose is TAF, which is a solution of 7 mL formalin (40% formaldehyde) and 2 mL triethanolamine in 91 mL distilled water. Nematodes are first killed by heating them for a few seconds in a small drop of water on a slide, until they just stop moving (longer heating will damage the specimens). An amount of double-strength TAF equal to the drop of water is immediately added or the nematodes are removed from the water and put into a new drop of warmed TAF. The preserved specimens are mounted on a glass slide with a fixed cover-slip and are examined under high-power light microscopy. To see the necessary diagnostic features, it may be necessary to use the highest power of the microscope (e.g. X100 with oil immersion) or to apply interference contrast microscopy. Appendix 2 Extraction of motile stages of Meloidogyne chitwoodi and M. fallax from roots and soil. Extraction from roots Second-stage juveniles of the nematode can be extracted from the roots of suspect plants by the incubation method: roots are carefully washed free from adhering soil and cut into short lengths (5–10 cm). The root pieces are placed on a moist filter paper on an extraction sieve. The sieve is placed in a shallow dish with a thin layer of water and placed in an incubation chamber for 14 days at 20 °C. During this period, most of the juveniles will hatch from the eggs, leave the root tissues and crawl trough the filter paper into the water. The water, with the nematodes, is poured into a glass dish and examined under a dissecting microscope at 25–35× magnification. A quicker extraction method is the maceration/centrifugal/flotation method of Coolen (1979): the roots are homogenized in an electric macerator at about 12600 rev min−1 for 30 s. The suspension is poured onto a sieve of 1200-µm pore size and washed. The washing water that passes through the sieve is centrifuged with 1% kaolin powder at 1500 g for 4 min. The sediment is re-suspended in sucrose, MgSO4 or ZnSO4 solution (sucrose 484 g L−1; 1.18 specific gravity, others 1.16 sp.gr.) and centrifuged at 1500 g for 4 min. The supernatant is poured through a sieve of 5-µm pore size. Nematodes can be washed off the surface of the sieve and collected in a glass dish for examination under a microscope at 25–35× magnification. Nematodes can be extracted from tubers, even those without symptoms, by peeling the skin, homogenizing it in a blender and examining a sample under the microscope Other suitable methods for extracting motile nematodes from plant tissue may be employed, taking in consideration the time that eggs need to mature before hatching can take place. 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 carefully suspended over 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 (Web Fig. 8). After 48 h, nematodes have emerged from the soil and fallen to the bottom of the funnel. They can be obtained by releasing a small amount of water from the base of the funnel into a glass dish. Any nematodes recovered are examined under a dissecting microscope at 25–35X. For larger amounts of soil, standard methods for extracting motile nematodes 0.3–1.2 mm in length from soil may be employed (Southey, 1986). For example, the following simple flotation/sieving technique may be used: 100 g 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 and collected in a beaker, and poured onto a nylon sieve suspended on a Baermann funnel (as described above, but without the paper filter). 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 25–35X. 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 (Southey, 1986). Appendix 3 – Recommendations for molecular and isozyme detection Molecular and/or isozyme techniques should be used for the identification of root-knot nematodes, besides morphological and morphometrical observations. There is a wide variation in available molecular and isozyme protocols for the identification of Meloidogyne species, and the following protocols are suggested. A reliable electrophoresis method is available for the identification of young adult Meloidogyne females. If young egg-laying females cannot be isolated for electrophoresis, a PCR method may be used for males, second-stage juveniles, eggs or infested root galls. Isozyme electrophoresis The method of Karssen et al. (1995) should be followed. It is preferable to use two isozymes, e.g. malate dehydrogenase (Mdh, EC 1.1.1.37) and esterase (Est, EC 3.1.1.1). A positive control for isozymes should be included, i.e. a well-known root-knot nematode species like M. hapla or M javanica as well as the M. chitwoodi and M. fallax reference controls. Relative movements (RM%) of the individual isozyme bands are compared with those of van Meggelen et al. (1994), Karssen (1996a) or Karssen (2002). For M. chitwoodi, an N1a Mdh type and S1 Est type should be obtained; for M. fallax an N1b Mdh type and a weak F3 Est type. Additional information on isozyme diversity and coding can be found in Esbenshade & Triantaphyllou (1985). If a method other than Karssen et al. (1995) has been used, it should be described in detail (sample & gel preparation, staining procedure and running conditions). Prepare original photographs of gel(s). PCR tests Several PCR methods are recommended for the identification of M. chitwoodi and M. fallax as species-specific primers are applied: • Zijlstra (2000), a highly sensitive PCR method based on species-specific primers or sequence-characterized amplified regions (SCARs) designed from RAPD fragments • Wishart et al. (2002), a sensitive PCR method based on species-specific primers designed from ribosomal IGS regions • Zijlstra et al. (1997), a relatively simple ITS-RFLP PCR method based on the well-known Vrain primers. The first two methods are useful for the identification of (single or more) second-stage juveniles, while females, eggs, males and even galls of infested roots can also be used. DNA isolation: see the cited references. Alternatively, the High Pure PCR Template Preparation Kit (Roche, Almere, NL) can be used for convenience. Primers: Zijlstra (2000) describes different specific SCAR primer sets for normal PCR, multiplex PCR (used to detect species mixtures) and nested PCR, respectively. For the normal PCR, the M. fallax primer set: Ff: 5′-CCAAACTATCGTAATGCATTATT-3′ Rf: 5′-GGACACAGTAATTCATGAGCTAG-3′ results in a 530 bp amplicon. The M. chitwoodi primer set: Fc: 5′-TGGAGAGCAGCAGGAGAAAGA-3′ Rc: 5′-GGTCTGAGTGAGGACAAGAGTA-3′ results in a 800 bp amplicon. The multiplex and nested primer sets results in different sized amplicons. Wishart et al. (2002) designed the specific primers: JMV1 5′-GGATGGCGTGCTTTCAAC-3′ JMV2 5′-TTTCCCCTTATGATGTTTACCC-3′ resulting in a 540 bp amplicon for M. chitwoodi and a 670 bp for M. fallax. Zijlstra et al. (1997) use the ITS-specific Vrain primers: 5′-TTGATTACGTCCCTGCCCTTT-3′ 5′-TTTCACTCGCCGTTACTAAGG-3′ resulting for both species in a 760 bp amplicon. 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