Preferred name: Meloidogyne fallax
Authority: Karssen
Taxonomic position: Animalia: Nematoda: Chromadorea: Rhabditida: Meloidogynidae
Common names in English: false Columbia root-knot nematode
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Notes on taxonomy and nomenclature EPPO Categorization: A2 list
EU Categorization: A2 Quarantine pest (Annex II B)
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EPPO Code: MELGFA

M. fallax was detected on and described from potato (Solanum tuberosum). Host-range includes a wide range of dicotyledonous and monocotyledons plants, including weeds, ornamentals, and economically important crops such as carrot (Daucus carota), black salsify (Scorzonera hispanica) and tomato (Solanum lycopersicum). The experimental host range of M. fallax mostly overlaps that of M. chitwoodi, but differential hosts have been found. For example dwarf beans (Phaseolus vulgaris), valerian (Valeriana officinalis), maize (Zea mays), Erica cinerea and Potentilla fruticosa are good hosts for M. chitwoodi and not for M. fallax, while the reverse is the case for Oenothera glazioviana, Phacelia tanacetifolia, Hemerocallis cv. Rajah and Dicentra spectabilis (Brinkman et al., 1996). It is expected that many more plant species will be hosts of M. fallax than currently known, since this is the case also with other, closely related root knot nematodes.

Host list: Allium porrum, Asparagus officinalis, Avena strigosa, Beta vulgaris, Cichorium intybus, Cynara scolymus, Daucus carota subsp. sativus, Fragaria x ananassa, Hemerocallis sp., Hordeum vulgare, Lactuca sativa, Lamprocapnos spectabilis, Leptinella sp., Lolium multiflorum, Medicago sativa, Oenothera glazioviana, Phacelia tanacetifolia, Scorzonera hispanica, Solanum lycopersicum, Solanum nigrum, Solanum physalifolium, Solanum tuberosum, Trifolium repens, Triticum aestivum

After the first record near Baexem (NL) in 1992, M. fallax was recorded on potato at several locations in the southern and south-eastern part of the Netherlands (Karssen, 1996), close to the German and Belgium borders. Within the EPPO region it was detected locally in Belgium, France, Germany, Sweden, Switzerland and the United Kingdom (England). In addition, Topalović et al. (2017) revealed that a Meloidogyne species detected in Ireland in 1965 belongs to M. fallax. M. fallax has never been reported from the natural environment in Europe. Outside Europe it has been reported from Australia, Chile, New Zealand and South Africa. New Zealand is the only known country where M. fallax is widely distributed (North and South Island) and detected in cropping and pasture fields (Rohan et al., 2016), strongly suggesting that it could be the place of origin of this pest. 

EPPO Region: Belgium, France (mainland), Germany, Netherlands, Sweden, Switzerland, United Kingdom (England)
Africa: South Africa
Asia: Indonesia (Java)
South America: Chile
Oceania: Australia (Queensland, South Australia, Tasmania, Victoria, Western Australia), New Zealand

The life cycles of M. fallax and M. chitwoodi are, in general, the same with respect to root penetration, gall induction, symptomatology, number of moults, parthenogenetic reproduction and chromosome number: Both M. chitwoodi and M. fallax usually reproduce by parthenogenesis. They can have one to three generations per year in the Netherlands and produce several hundreds of eggs per female, deposited in an egg sac. These egg sacs allow the eggs to survive under unfavorable conditions (EFSA, 2019). Initial results by van der Beek (1997) indicated that M. fallax had a shorter life cycle than M. chitwoodi in a virulence study on potato.

Host races, as described for M. chitwoodi, have not been detected for M. fallax so far. Successful hybridization was not obtained when M. fallax and M. chitwoodi were crossed in two different experiments; the F1 was viable, but the F2 second-stage juveniles were not viable and showed morphological distortions (van der Beek & Karssen, 1997).

In addition, differences in terms of hatching responses to root diffusate and host age between M. chitwoodi and M. fallax suggests different survival strategies between these two species (Wesemael et al., 2006).

M. fallax, M. chitwoodi and M. minor Karssen are closely related, morphologically and at DNA level. Phylogenetically they appear to be in the same distinct clade within the genus Meloidogyne (Holterman et al., 2009; Elling, 2013).

Symptoms

Above-ground symptoms of heavily infested plants include stunting and yellowing, while below ground galling is typical. 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 (these secondary roots are seen 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 infested, may be free from visible external symptoms, while the internal potato tissue is necrotic and brownish, just below the peel (EPPO, 2016a).

Morphology

Sedentary females are annulated, pearly white and globular to pear-shaped, 400-720 µm long and 250-460 µm wide. The stylet is dorsally curved, 13.9-15.2 µm long, with rounded to ovoid stylet knobs, slightly sloping posteriorly. The non-sedentary males are vermiform, annulated, slightly tapering anteriorly, bluntly rounded posteriorly, 735-1520 µm long and 27-44 µm wide. The stylet is 18.9-20.9 µm long, with large rounded knobs, set off from the shaft. The non-sedentary second-stage juveniles are vermiform, annulated, tapering at both ends, 380-435 µm long, 13.3-16.4 µm wide, with a tail length of 46-56 µm and a hyaline tail part 12.2-15.8 µm in length (Karssen, 1996).

M. fallax is closely related morphologically to M. chitwoodi, and this misleading resemblance was the reason for giving the species its name. The most striking differences for males and females are stylet length (longer for M. fallax) and stylet knob shape (M. fallax: prominent and rounded; M. chitwoodi: small and irregular). The second-stage juveniles differ in mean body length, tail length and hyaline tail length (all longer for M. fallax). With the scanning electron microscope, it can be observed that the male head of M. fallax has an elevated labial disk. Differences exist in the female perineal pattern (M. fallax has a relatively higher dorsal arch and thicker striae) (EPPO, 2016a; see also Karssen, 2002). 

The species can be reliably distinguished by morphological observation of females, males, and second-stage juveniles in combination with biochemical (isozyme electrophoresis) or molecular (PCR) methods; see EPPO, 2016. To predict the amount of potato tuber damage caused by M. fallax, a quantitative DNA-test was developed for soil (Hay et al., 2016).

Detection and inspection methods

Specific guidance on the sampling of soil and potato tubers is given in the EPPO Standards PM 9/17 (EPPO, 2013b), PM 3/71 (EPPO, 2007) and PM 3/69 (EPPO, 2019a). Populations in the soil rapidly decline in the absence of a host and nematodes reproduce better on a good host. Therefore, detection of the nematodes through field inspection and soil sampling is more sensitive if done as close as possible to the time of harvest of a host crop, targeting particularly susceptible plants (EPPO, 2013b). In each lot of potato (typically 25 tonnes), 200 tubers are randomly sampled and processed (EPPO, 2019a). 

Nematode extraction and identification should be carried out according to EPPO Standards PM 7/119 (EPPO, 2013a) and PM 7/41 Diagnostic protocol for Meloidogyne chitwoodi and M. fallax (EPPO, 2016a).

Historically, in order to distinguish M. fallax from other Meloidogyne spp., molecular methods have been used. Karssen et al. (1995) discriminated M. fallax and M. chitwoodi females by their esterase (EC 3.1.1.1) and malate dehydrogenase (EC 1.1.1.37) isozyme patterns, using the general method of Esbenshade & Triantaphyllou (1985) for identification of female Meloidogyne species by isozyme electrophoresis. Additionally, the isozyme glucose 6-phosphate dehydrogenase (EC 1.1.1.49) was used to differentiate the two species (van der Beek & Karssen, 1997). van der Beek et al. (1997) and Tastet et al. (1999) used mini two-dimensional gel electrophoresis to study the total soluble protein patterns of M. hapla, M. chitwoodi and M. fallax, and confirmed these species to be distinct biological groups.  An overwhelming number of molecular tests has been developed to identify M. fallax from soil, root or tubers and to separate it from related species such as M. chitwoodi, including PCR (Peterson & Vrain, 1996, Peterson et al., 1997; Wishart et al., 2002), PCR RFLP (Zijlstra et al. 1995, Zijlstra, 1997) AFLP (v.d. Beek et al., 1998; Fargette et al., 2005), SCAR (Zijlstra, 2000; Fourie et al., 2001), real time TaqMan PCR (Zijlstra & v. Hoof, 2006; de Haan et al., 2014), RAPD (Adam et al., 2007), satDNA (Castagnone-Sereno et al., 1999; Castagnone-Sereno, 2000; Mestrovich et al., 2009 & 2013), LAMP (Zhang & Gleason, 2019), HRMC (Holterman et al., 2012), barcoding (Hodgetts et al., 2016, EPPO, 2016b). A serological test was also developed to separate M. fallax from M. chitwoodi and other root-knot nematodes (Tastet et al., 2001). Only a selection of the above-mentioned tests are recommended for identification (see EPPO, 2016a).

M. fallax has very limited potential for natural movement; only second-stage juveniles can move in the soil and, at most, only a few tens of centimetres. The most likely pathway for introducing M. fallax into a new area is through the movement of infested or contaminated planting material. Infested host plants or host products such as bulbs or tubers can easily transport the nematode. The movement of non-host plants for planting (e.g. seedling transplants, nursery stock), non-host plant products (e.g. bulbs, tubers, corms and rhizomes), equipment and machinery which are contaminated with soil infested with M. fallax could also result in spread (EPPO, 2013b). Soil as such is also a possible pathway. Infective juveniles of this genus have been known to persist for more than one year in the absence of host plants. Nematode movement can also be facilitated by contaminated irrigation water.

Economic impact

In trials, M. fallax caused the same symptoms on potato tubers, black salsify and carrots as M. chitwoodi, i.e. external galling and internal necrosis just below the skin (Brinkman et al., 1996; van Riel & Goossens, 1996). The reported natural outbreaks of M. fallax on potato showed these external symptoms (Karssen, 1996). Goosens (1995) reported infected Asparagus officinalis and several ornamentals with root-knots in an experimental field with an infestation of M. fallax. M. fallax sometimes occurs in mixed infestations with M. chitwoodi (Wesemael et al., 2006). 

Meloidogyne fallax mainly induces quality losses caused by cosmetic damage on black salsify, leek, carrot and potato. If the level of infection is high a complete rejection of these crops is possible. So far, this nematode has a limited damaging effect on other known host crops (EFSA, 2012).

In countries where M. fallax is regulated, traded plants and plant products infested with M. fallax may need to be destroyed.

Control

There is no direct practical experience of the control of M. fallax. Research on M. fallax in the Netherlands has focused on host suitability, damage thresholds, effect of fallow, the use of green manure crops and time of sowing. The first results indicate that fallow for one year reduced the population by more than 95%, but this reduction was not sufficient to ensure that subsequent crops met quality standards. There was less damage in sugar beet and carrot when these crops were sown later in spring. Farmers are advised to be careful when growing green-manure crops on infested fields, because some species are suitable hosts for M. fallax. Phaseolus vulgaris was the only tested crop with no reproduction of M. fallax, while maize and cereals were poor hosts (Brommer, 1996). Several green manures were tested, and no reproduction of M. fallax was found on Eruca Sativa cv Trio, Tagetus patula and Borago spp., while Raphanus sativus was a poor host. Avena strigosa turned out to be a very good host (Visser & Molendijk, 2015).

Janssen et al. (1996) tested several wild tuber-bearing Solanum species, to determine the level of resistance to M. hapla, M. chitwoodi and M. fallax. High resistance to M. chitwoodi and M. fallax was observed in genotypes of S. bulbocastanum, S. hougasii, S. cardiophyllum, S. fendleri and S. brachistotrichum. Differential resistance between M. chitwoodi and M. fallax was observed in S. chacoense, S. stoloniferum and S. gourlayi. Resistance to M. fallax was also found in Solanum sparsipilum (Kouassi et al., 2004) and the resistance gene was studied (Kouassi et al., 2006). By crossing wild tuber bearing Solanum species with Solanum tuberosum, successful introgression of resistance to M. fallax and M. chitwoodi into potato was obtained (Janssen et al., 1997).

In addition, useful resistance against M. fallax was found in sea beet (Beta maritima) (Yu et al., 1999) and used to develop a resistant sugar beet line (Beta vulgaris) (Yu, 2001).

Wesemael & Moens (2012) tested ten common bean (Phaseolus vulgaris) cultivars and noted that all these cultivars were non-hosts or poor hosts for M. fallax.

An interesting potential control method consists of using the hyper-parasitic bacterium Pasteuria spp. So far three different species of Pasteuria are reported to be able to parasitize M. fallax: P. penetrans, P. nishizawae and P. hartismeri (Wishart et al., 2004; Bishop et al., 2007).

Phytosanitary risk

Because M. fallax occurs in crops and situations which are similar to those for M. chitwoodi, and the two species are closely related and difficult to distinguish, M. fallax presents a phytosanitary risk similar to that of M. chitwoodi (EFSA, 2012). M. fallax may be able to establish in a large proportion of its host area but may only cause significant damage in certain areas and under certain conditions, causing complete crop rejection (e.g. on potato, carrot and/or black salsify). Soils with a coarse texture such as sandy and sandy-loam soils have a higher probability of being contaminated. Narrow rotation or rotation with alternative hosts facilitates a rapid build-up of population levels and therefore increases the risk for establishment of M. chitwoodi and M. fallax in a particular field. As observed since 1992, M. fallax is expected to spread slowly (EFSA, 2012).

Measures similar to those for other root-knot nematodes would appear relevant, i.e. that rooted host plants for planting (with or without soil), non-host plants for planting with soil attached and plant products with soil attached come from a pest free area, a pest free place of production or are produced under protected cultivation. Alternatively, soil from non-host plants for planting or plant products can be removed. Soil as such can originate from a pest free area or a pest free place of production. Used machineries, equipment, vehicles, and passengers’ shoes can be cleaned. Publicity would allow to enhance public awareness on M. fallax when travelling. 

Specific requirements are recommended in EPPO Standard PM 8/1 Commodity-specific phytosanitary measures for Potato (EPPO, 2017) for seed and ware potatoes to be imported from third countries. In this Standard, seed potato freedom from M. fallax can also be guaranteed by testing the seed potatoes after harvest following EPPO Standard PM 3/69 Meloidogyne chitwoodi and M. fallax: sampling potato tubers for detection (EPPO, 2019a). Ware potatoes freedom can also be guaranteed by implementing EPPO Standard PM 9/17 National regulatory control system for Meloidogyne chitwoodi and M. fallax (EPPO, 2013b). EPPO Standard PM 3/61 details conditions for establishing pest-free areas and pest-free production and distribution systems for quarantine pests of potato (EPPO, 2019b).

Measures to contain or eradicate M. chitwoodi and M. fallax are described in the national regulatory control system (EPPO, 2013b).

Adam MAM, Phillips MS & Blok VC (2007) Molecular diagnostic key for identification of single juveniles of seven common and economically important species of root-knot nematode (Meloidogyne spp.). Plant Pathology 56, 190-197.

Bishop AH, Gowen SR, Pembroke B & Trotter JR (2007) Morphological and molecular characteristics of a new species of Pasteuria parasitic on Meloidogyne ardenensis. Journal of Invertebrate Pathology 96, 28-33.

Brinkman H, Goossens JJM & van Riel HR (1996) Comparative host suitability of selected crop plants to Meloidogyne chitwoodi and M. fallax. Anzeiger für Schädlingskunde, Planzenschutz, Umweltschutz69, 127-129.

Brommer E (1996) [Control of the root-knot nematode Meloidogyne fallax]. Publicatie Proefstation voor de Akkerbouw en de Groenteteelt in de Vollegrond no. 81B, pp. 159-163 (in Dutch). 

Castagnone-Sereno P, Leroy F, Bongiovanni M, Zijlstra C & Abad P (1999) Specific diagnosis of the two root-knot nematodes, Meloidogyne chitwoodi and M. fallax. Phytopathology 89, 380-384.

Castagnone-Sereno P (2000) Use of satellite DNA for specific diagnosis of the quarantine root-knot nematodes Meloidogyne chitwoodi and M. fallax. EPPO Bulletin 30, 581-584.

de Haan EG, Dekker CCEM, Tameling WIL, den Nijs LJMF, Bovenkamp GW & van den Kooman-Gersmann M (2014) The MeloTuber Test: a real-time TaqManReq. PCR-based assay to detect the rootknot nematodes Meloidogyne chitwoodi and M. fallax. Bulletin OEPP/EPPO Bulletin 44, 166-175.

EFSA (2012) Macleod A, Anderson H, Follak S, van der Gaag DJ, Potting R, Pruvost O, Smith J, Steffek R, Vloutoglou I, Holt J, Karadjova O, Kehlenback H, Labonne G, Reynaud P, Viaene N, Anthoine G, Holeva M, Hostachy B, Ilieva Z, Karssen G, Krumov V, Limon P, Meffert J, Niere B, Petrova E, Peyre J, Pfeilstetter E, Roelofs W, Rothlisberger F, Sauvion N, Schenck N, Schrader G, Schroeder T, Steinmöller S, Tjou-Tam-Sin L, Ventsislavov V, Verhoeven K & Wesemael W. Pest risk assessment for the European Community plant helath: a comparative approach with case studies. Cases: Meloidogyne chitwoodi and M. fallax. EFSA Supporting publications 2012 EN-319. 1053 pp.

EFSA (2019) den Nijs L, Camilleri M, Diakaki M, Schenk M & Vos S. Pest survey card on Meloidogyne chitwoodi and Meloidogyne fallax. EFSA supporting publication 2019 EN-1572. 20 pp.

Elling AA (2013) Major emerging problems with minor Meloidogyne species. Phytopathology 103, 1092-1102.

EPPO (2007) EPPO Standard PM 3/71(1) General crop inspection procedure for potatoes + corrigendum. EPPO Bulletin 37, 592-597. Available at: https://gd.eppo.int/standards/PM3/ 

EPPO (2013a) EPPO Standard PM 7/119 (1) Nematode extraction. EPPO Bulletin 43, 471-495. Available at: https://gd.eppo.int/standards/PM7/

EPPO (2013b). EPPO Standard PM 9/17 (1) Meloidogyne chitwoodi and M. fallax. EPPO Bulletin 43, 527-533. Available at: https://gd.eppo.int/standards/PM9/ 

EPPO (2016a) EPPO Standard PM 7/41 (3). Meloidogyne chitwoodi and M. fallax. EPPO Bulletin 46, 171-189. Available at: https://gd.eppo.int/standards/PM7/

EPPO (2016b) EPPO Standard PM 7/129(1) DNA barcoding as an identification tool for a number of regulated pests. EPPO Bulletin 46, 501-537. Available at: https://gd.eppo.int/standards/PM7/

EPPO (2019a) EPPO Standard PM 3/69(2) Meloidogyne chitwoodi and M. fallax: sampling potato tubers for detection. EPPO Bulletin 49, 486-487. Available at: https://gd.eppo.int/standards/PM3/ 

EPPO (2019b) EPPO Standard PM 3/61(2) Pest-free areas and pest-free production and distribution systems for quarantine pests of potato. EPPO Bulletin 49, 480-481. Available at: https://gd.eppo.int/standards/PM3/ 

Esbenshade PR & Triantaphyllou AC (1985) Use of enzyme phenotypes for identification of Meloidogyne species. Journal of Nematology 17, 6-20.

Fargette M, Lollier V, Phillips M, Blok V & Erutos R (2005) AFLP analysis of the genetic diversity of Meloidogyne chitwoodi and M. fallax, major agricultural pests. Comtes Rendus Biologies 328, 455-462.

Fourie H, Zijlstra C & McDonald AH (2001) Identification of root-knot nematode species occurring in South Africa using SCAR-PCR technique. Nematology 3, 675-680.

Goossens JJM (1995) Host range test of Meloidogyne n.sp. In: Annual Report 1994 Diagnostic Centre, pp. 95-97. Plant Protection Service, Wageningen (NL).

Hay FS, Ophel-Keller K, Hartley DM & Pethybridge SJ (2016) Prediction of potato tuber damage by root-knot nematodes using quantitative DNA assay of soil. Plant Disease 100, 592-600.

Hodgetts J, Ostoja-Starzewski JC, Prior T, Lawson R, Hall J & Boonham N (2016) DNA barcoding for biosecurity: case studies from the UK plant protection program. Genome 59, 1033-1048. 

Holterman MHM, Karssen G, van den Elsen SJJ, van Megen HHB, Bakker J & Helder H (2009) SSU rDNA-based phylogeny of the Tylenchida sheds light on the evolution of plant feeding & establishes relationships among high impact plant parasitic nematodes. Phytopathology 99, 227-235.

Holterman MHN, Oggenfuss M, Frey JE & Kiewnick S (2012) Evaluation of high-resolution melting curve analysis as a new tool for root-knot nematode diagnostics. Journal of Phytophatology 160, 59-66.Janssen GJW, van Norel A, Verkerk-Bakker B & Janssen R (1996) Resistance to Meloidogyne chitwoodi, M. fallax and M. hapla in wild tuber-bearing Solanum spp. Euphytica 92, 287-294.

Janssen GJW, Janssen R, Van Norel A, Verkerk-Bakker B & Hoogendoorn J (1996) Expression of resistance to the root-knot nematodes, Meloidogyne hapla and M. fallax, in wildSolanum spp. under field conditions. European Journal of Plant Pathology 102, 859-865.

Janssen GJW, van Norel A, Verkerk-Bakker B & Janssen R (1997) Intra- and interspecific variation of root-knot nematodes, Meloidogyne spp., with regard to resistance in wild tuber bearing Solanum species. Fundamental and Applied Nematology 20, 449-457.

Karssen G (1994) The use of isozyme phenotypes for the identification of root-knot nematodes (Meloidogyne species). In: Annual Report 1992 Diagnostic Centre, pp. 85-88. Plant Protection Service, Wageningen (NL).

Karssen G (1995) Morphological and biochemical differentiation in Meloidogyne chitwoodi populations in the Netherlands. Nematologica 41, 314-315.

Karssen G (1996) Description of Meloidogyne fallax n.sp. (Nematoda: Heteroderidae), a root-knot nematode fromthe Netherlands. Fundamental and applied Nematology 19, 593-599.

Karssen G, van Hoenselaar T, Verkerk-Bakker B & Janssen R (1995) Species identification of cyst and root-knot nematodes frompotato by electrophoresis of individual females. Electrophoresis 16, 105-109.

Karssen G (2002) The plant-parasitic nematode genus Meloidogyne Göldi, 1892 in Europe. Brill Academic Publishers, the Netherlands. 157pp.

Kouassi AB, Kerlan MC, Sobczak M, Bantec JP, Rouaux C, Ellisseche D & Mugniery D (2004) Resistance to the root-knot nematode Meloidogyne fallax in Solanum sparsipilum. Nematology 6, 389-400.

Kouassi AB, Kerlan MC, Caromel B, Dantec JP, Fouville D, Manzanares-Dauleux M, Ellisseche D & Mugniery D (2006) A major gene mapped on chromosome XII is the main factor of a quantitatively inherited resistance to Meloidogyne fallax in Solanum sparsipilum. Theoritical and Applied Genetics 112, 699-707. 

Mestrovic N, Phlol M & Castagnone-Serone P (2009) Relevance of satellite DNA genomic distribution in phylogenetic analysis: a case study with root-knot nematodes of the genus Meloidogyne. Molecular Phylogenetics and Evolution 50, 204-208.

Mestrovic N, Pavlek M, Car A, Castagnone-Serone P, Abad, P & Plohl M (2013) Conserved DNA motifs, including CENP-B box-like, are possible promotors of satellite DNA array rearrangesments in nematodes. PloS ONE 8, e67328. 

Peterson DJ & Vrain TC (1996) Rapid identification of Meloidogyne chitwoodi, M. hapla and M. fallax using PCR primers to amplify their ribosomal intergenic spacer. Fundamental and applied Nematology 19, 601-605.

Peterson DJ, Zijlstra C, Wishart J, Blok V & Vrain T (1997) Specific probes efficiently distinguish root-knot nematode species using signature in the ribosomal intergenic spacer. Fundamental and applied Nematology 20, 619-626.

Rohan TC, Aalders LT, Bell NL & Shah FA (2016) First report of Meloidogyne fallax hosted by Trifolium repens (white clover): implications for pasture and crop rotations in New Zealand. Australasian Plant Disease Notes. 11, 14.

Tastet, C, Val F, Lesage M, Renault L, Marche L, Bossis, M & Mugniery D (2001) Application of a putative fatty-acid binding protein to discriminate serologically the two European quarantine root-knot nematodes, Meloidogyne chitwoodi and M. fallax from other Meloidogyne species. European Journal of Plant Pathology 107, 821-832.

Topalović O, Moore JF, Janssen T, Bert W & Karssen G (2017) An early record of Meloidogyne fallax from Ireland. ZooKeys 643, 33–52. https://doi.org/10.3897/zookeys.643.11266

van der Beek JG (1997) Isolate-by-cultivar interaction in M. hapla, M. chitwoodi and M. fallax on potato. In: Interaction between root-knot nematodes and Solanum spp. variation in pathogenicity, cytology, proteins and DNA. pp. 41-53. Thesis, Wageningen Agricultural University, Wageningen (NL).

van der Beek JG & Karssen G (1997) Interspecific hybridization of meiotic parthenogenetic Meloidogyne chitwoodiandM. fallax. Phytopathology 87,1061-1066.

van der Beek JG, Folkertsma R,  Poleij LM, van Koert PHG & Bakker J (1997) Molecular evidence that Meloidogyne hapla, M. chitwoodi and M. fallax are distinct biological entities. Fundamental and applied Nematology 20, 513-520.

van der Beek JG, Folkertsma R, Zijlstra C, van Koert PHG, Poleij LM & Bakker J (1998) Genetic variation among parthenogenetic Meloidogyne species revealed by AFLPs and 2D-protein electrophoresis contrasted to morphology. Fundamental and Applied Nematology 21, 401-411.

van Meggelen JC, Karssen G, Janssen GJW, Verkerk B & Janssen R (1994) A new race of Meloidogyne chitwoodi?. Fundamental and appliedNematology 17,93.

van Riel HR & Goossens JJM (1996) Response of Meloidogyne fallax to differential hosts of M. chitwoodi. In: Annual Report 1995 Diagnostic Centre, pp. 100-101. Plant Protection Service, Wageningen (NL).

Visser JHM & Molendijk LPG (2015) [Host suitability new green manures for plant-parasitic nematodes]. Publicatie WUR, Praktijkonderzoek Plant & Omgeving no. 633, 29pp. (in Dutch).

Wesemael WML, Perry RN & Moens M (2006) The influence of the root diffusate and host age on hatching of the root-knot nematodes. Nematology 8, 895-902.

Wesemael WML & Moens M (2012) Screening of common bean (Phaseolus vulgaris) for resistance against temperate root-knot nematodes (Meloidogyne spp.). Pest Management Science 68, 702-708. 

Wishart J, Phillips MS & Blok VC (2002) Ribosomal intergenic spacer: a polymerase chain reaction diagnostic for Meloidogyne chitwoodi, M. fallax and M. hapla. Phytophatology 92, 884-892.

Wishart J, Blok VC, Phillips MS & Davies KG (2004) Pasteuria penetrans and P. nishizawae attachment to Meloidogyne chitwoodi, M. fallax and M. hapla. Nematology 6, 507-510.

Yu M, Heijbroek W & Pakish LM (1999) The sea beet source of resistance to multiple species of root-knot nematode. Euphytica 108, 151-155.

Yu MH (2001) Registration of M6-1 root-knot nematode resistant sugarbeet germplasm. Crop Science 41, 278-279.

Zhang L & Gleason C (2019) Loop-mediated isothermal amplification for the diagnostic detection of Meloidogyne chitwoodi and M. fallax. Plant Disease 103, 12-18.

Zijlstra C (1997) A fast PCR assay to identify Meloidogyne hapla, M. chitwoodi and M. fallax, and to sensitively differentiate them from each other and from M. incognita in mixtures. Fundamental and applied Nematology 20, 505-511.

Zijlstra C, Lever AEM, Uenk BJ & van Silfhout CH (1995) Differences between ITS regions of isolates of root-knot nematodes Meloidogyne hapla and M. chitwoodi. Phytopathology 85, 1231-1237.

Zijlstra C (2000) Reliable identification of quarantine root-knot nematodes Meloidogyne chitwoodi and M. fallax by PCR-based techniques. EPPO Bulletin 30, 575-579.

Zijlstra C & van Hoof RA (2006) A multiplex real-time polymerase chain reaction (TaqMan) assay for the simultaneous detection of Meloidogyne chitwoodi and M. fallax. Phytopathology 96, 1255-1262. 

This datasheet was extensively revised in 2020 by Karssen G. His valuable contribution is gratefully acknowledged.

EPPO (2024) Meloidogyne fallax. EPPO datasheets on pests recommended for regulation. https://gd.eppo.int (accessed 2024-05-09)

This datasheet was first published in the EPPO Bulletin in 1999 and is now maintained in an electronic format in the EPPO Global Database. The sections on 'Identity', ‘Hosts’, and 'Geographical distribution' are automatically updated from the database. For other sections, the date of last revision is indicated on the right.

EPPO (1999) EPPO Data sheets on quarantine pests - Meloidogyne fallax.  EPPO Bulletin 29(4), 493-496. https://doi.org/10.1111/j.1365-2338.1999.tb01425.x