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Entomopathogenic nematodes in the genera Steinernema and Heterorhabditis are recognized as insect-parasitic nematodes, beneficial nematodes, biocontrol agents, biological control agents, biological insecticides or biopesticides.

These nematodes are also recognized as pathogens or microbial control agents because of their symbiotic association with bacteria (Xenorhabdus and Photorhabdus spp.) that are mainly pathogenic to insects. Because of mutualistic relationship with pathogenic bacteria these nematodes are named as entomopathogenic nematodes.

These beneficial nematodes contribute to the regulation of natural populations of insects.  However, the population of naturally occurring entomopathogenic nematodes is normally not high enough to manages soil dwelling plant pests. Therefore, during last 3-4 decades, these live nematodes have been commercially mass produced and inundatively applied to control many garden insects, turfgrass insects, nursery insects, greenhouse insects and insects that feed on different field crops.

Use of this natural control of insects is beneficial for both the environment and humans because it reduces use of chemical insecticides/pesticides.

These biopesticides (entomopathogenic nematodes and their symbiotic bacteria) are safe to produce and not harmful to users, application personnel, mammals, most beneficial insects or plants.

Since entomopathogenic nematodes do not cause any health risk to the consumers of nematode treated agricultural produce and damage to the environment, they are exempted from registration requirements in most countries.

These biological control agents have also no detrimental effect on other benefical nematodes including bacterial feeders, some fungal feeders (Aphelenchus sp.), predatory nematodes and other soil microbial communities.

But entomopathogenic nematodes can be detrimental to plant-parasitic nematodes that are responsible for causing a tremendous economic loss to our agriculture industry throughout world. It has been demonstrated that entomopathogenic nematodes can suppress the populations of many economically important plant-parasitic nematodes including foliar nematodes, potato cyst nematodes, ring nematodes, root-knot nematodes,  root lesion nematodes, sting nematodes, stubby root nematodes and stunt nematodes.

Scientific Publications by Dr. Ganpati B. Jagdale on insect-parasitic nematodes (EPNs)

I. Book Chapters

Tomalak, M., Piggott, S. and Jagdale, G. B. 2005. Glasshouse applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166.

II. Research Publications

1. Jagdale, G.B., Kamoun, S., Grewal, P.S. 2009. Entomopathogenic nematodes induce components of systemic resistance in plants: Biochemical and molecular evidence. Biol. Control.51: 102-109
2. Hoy, C. W., Grewal, P. S., Lawrence, J. L., Jagdale, G., Acosta, N. 2008. Canonical correspondence analysis demonstrates unique soil conditions for entomopathogenic nematode species compared with other free-living nematode species. Biol. Control. 46: 371-379.
3. Jagdale, G. B. and Grewal, P. S. 2008. Influence of the entomopathogenic nematode Steinernema carpocapsae infected host cadavers or their extracts on the foliar nematode Aphelenchoides fragariae on Hosta in the greenhouse and laboratory. Biological Control 44: 13-23.
4. Shabeg, S .B., Jagdale, G. B., Cheng, Z, Hoy, C. W., Miller, S. A. and. Grewal, P. S. 2007. Indicative value of soil nematode food web indices and trophic group abundance in differentiating habitats with a gradient of anthropogenic impact. Environmental Bioindicators 2: 146-160.
   Jagdale, G. B., Casey, M. L., Grewal, P. S. and Luis Cañas. 2007. Effect of entomopathogenic nematode species, split application and potting medium on the control of the fungus gnat, Bradysia difformis (Diptera : Sciaridae), in the greenhouse at alternating cold and warm temperatures. Biological Control 43: 23-30.
   Jagdale, G. B. and Grewal, P. S. 2007. Storage temperature influences desiccation and ultra violet radiation tolerance of entomopathogenic nematodes. Journal of Thermal Biology 32: 20-27.
   Jagdale, G. B., Saeb, A. T., Nethi Somasekhar and Grewal, P. S. 2006. Genetic variation and relationships between isolates and species of the entomopathogenic nematode genus Heterorhabditis deciphered through isozyme profiles. Journal of Parasitology 92: 509- 516.
   Sandhu, S. K., Jagdale, G. B., Hogenhout, S. A. and Grewal, P. S. 2006. Comparative analysis of the expressed genome of the entomopathogenic nematode, Heterorhabditis bacteriophora. Molecular and Biochemical Parasitology 145: 239-244.
   Grewal, P. S., Susan Bornstein-Forst, S., Burnell, A. M., Glazer, I. and Jagdale, G. B. 2006. Physiological, genetic, and molecular mechanisms of chemoreception, thermobiosis and anhydrobiosis in entomopathogenic nematodes. Biological Control 38: 54- 65.
   Jagdale, G. B., Grewal, P. S. and Salminen, S. O. 2005. Both heat-shock and cold-shock influence trehalose metabolism in entomopathogenic nematodes. Journal of Parasitol 91: 988-994.
   Jagdale, G. B., Casey, M. L., Grewal, P. S. and Lindquist, R. K. 2004. Application rate and timing, potting medium and host plant on the efficacy of Steinernema feltiae against the fungus gnat, Bradysia coprophila, in floriculture. Biological Control 29: 296-305.
   Jagdale, G. B., and Grewal, P. S. 2003. Acclimation of entomopathogenic nematodes to novel temperatures: trehalose accumulation and the acquisition of thermotolerance. International Journal for Parasitology 33: 145-152.
   Grewal, P. S. and Jagdale, G. B. 2002. Enhanced trehalose accumulation and desiccation survival of entomopathogenic nematodes through cold preacclimation. Biocontrol Science and Technology 12: 533- 545.
   Jagdale, G. B. and Gordon, R. 1998. Effect of propagation temperatures on temperature tolerances of entomopathogenic nematodes. Fundamental and Applied Nematology 21: 177-183.
   Jagdale, G. B. and Gordon, R. 1998. Variable expression of isozymes in entomopathogenic nematodes follows laboratory recycling. Fundamental and Applied Nematology 21: 147-155.
   Jagdale, G. B. and Gordon, R.1997. Effect of temperature on the activities of glucose-6-phosphate dehydrogenase and hexokinase in entomopathogenic nematodes (Nematoda: Steinernematidae). Comparative Biochemistry and Physiology 118A: 1151-1156.
   Jagdale, G. B. Gordon, R. 1997. Effect of temperature on the composition of fatty acids in total lipids and phospholipids of entomopathogenic nematodes. Journal of Thermal Biology 22: 245-251.
   Jagdale, G. B. and Gordon, R. 1997. Effect of recycling temperature on the infectivity of entomopathogenic nematodes. Canadian Journal of Zoology 75: 2137-2141.
   Jagdale, G. B., Gordon, R. and Vrain, T. C. 1996. Use of cellulose acetate electrophoresis in the taxonomy of steinernematids (Rhabditida, Nematoda). Journal of Nematology 28: 301-309.
   Jagdale, G. B. and Gordon, R. 1994. Distribution of catecholamines in the nervous system of a mermithid nematode, Romanomermis culicivorax. Parasitology Research 80: 459-466.
   Jagdale, G. B. and Gordon, R. 1994. Distribution of FMRF-amide-like peptide in the nervous system of a mermithid nematode, Romanomermis culicivorax. Parasitology Research 80: 467-473.
   Jagdale, G.B. and Gordon, R. 1994. Role of catecholamines in the reproduction of Romanomermis culicivorax. Journal of Nematology 26: 40-45.
   Jagdale, G.B. and Gordon, R. 1994. Caudal papillae in Romanomermis culicivorax. Journal of Nematology 26: 235-237.

known species of symbiotic bacterial genus Photorhabdus associated with a nematode genus Heterorhabditis.

Identification based on colony morphology and molecular techniques

1. Photorhabdus luminescens (Thomas and Poinar 1979) Boemare et al. 1993
2. P. temperata
3. P. luminescens subsp. luminescens subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
4. P. luminescens subsp. akhurstii subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
5. P. luminescens subsp. kayaii subsp. nov., Hazir, Stackebrandt, Lang, Schumann, Ehlers & Keskin, 2004
6. P. luminescens subsp. laumondii subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
7. P. temperata sp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
8. P. temperata subsp. temperata subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
9. P. luminescens subsp. thracensis subsp. nov., Hazir, Stackebrandt, Lang, Schumann, Ehlers & Keskin, 2004

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known species of symbiotic bacterial genus Xenorhabdus Thomas and Poinar 1979 associated with a nematode genus Steinernema.

Identification based on colony morphology and molecular techniques

1. Xenorhabdus beddingii (Akhurst 1986) Akhurst and Boemare 1993
2. X. bovienii (Akhurst 1983) Akhurst and Boemare 1993
3. X. budapestensis Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
4. X. cabanillasii Tailliez, Pagès, Ginibre & Boemare, 2006
5. X. doucetiae Tailliez, Pagès, Ginibre & Boemare, 2006
6. X. ehlersii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
7. X. griffiniae Tailliez, Pagès, Ginibre & Boemare, 2006
8. X. hominickii Tailliez, Pagès, Ginibre & Boemare, 2006
9. X. indica Somvanshi, Lang, Ganguly, Swiderski, Saxena, & Stackebrandt 2006
10. X. innexi Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
11. X. japonica Nishimura et al. 1995
12. X. koppenhoeferi Tailliez, Pagès, Ginibre & Boemare, 2006
13. X. kozodoii Tailliez, Pagès, Ginibre & Boemare, 2006
14. X. mauleonii Tailliez, Pagès, Ginibre & Boemare, 2006
15. X. miraniensis Tailliez, Pagès, Ginibre & Boemare, 2006
16. X. nematophila (Poinar and Thomas 1965) Thomas and Poinar 1979
17. X. poinarii (Akhurst 1983) Akhurst and Boemare 1993
18. X. romanii Tailliez, Pagès, Ginibre & Boemare, 2006
19. X. stockiae Tailliez, Pagès, Ginibre & Boemare, 2006
20. X. szentirmaii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005

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• EPNs complete most of their life cycle in insects with an exception of infective juveniles, the only free-living stage found in soil.
• Infective juveniles of both Steinernema and Heterorhabditis locate a host and enter through its natural body openings such as mouth, anus or spiracles.
• Infective juveniles of Heterorhabditis also enter through the intersegmental members of the host cuticle.
• Infective juveniles then actively penetrate through the midgut wall or tracheae into the insect body cavity (hemocoel) containing insect blood (haemolymph).
• Once in the body cavity, infective juvenile releases symbiotic bacteria from its intestine in the insect haemolymph.
• Bacteria start multiplying in the nutrient-rich haemolymph and infective juveniles recover from their arrested state (dauer stage) and start feeding on multiplying bacteria and disintegrated host tissues.
• Toxins produced by the developing nematodes and multiplying bacteria in the body cavity kill the insect host usually within 48 hours.
• These bacteria also produce a plethora of metabolites, toxins and antibiotics with bactericidal, fungicidal and nematicidal properties, which ensures monoxenic conditions for nematode development and reproduction in insect cadaver.
• Heterorhabditid and Steinernematid nematodes differ in their mode of reproduction. For example, in heterorhabditid nematodes, the first generation individuals are produced by self-fertile hermaphrodites (hermaphroditic) but subsequent generation individuals are produced by cross fertilization involving males and females (amphimictic). In Steinernematid nematodes with an exception of one species, all generations are produced by cross fertilization involving males and females (amphimictic).
• Depending on availability of food resource, both heterorhabditid and steinernematid nematodes generally complete 2-3 generations within insect cadaver and emerge as infective juveniles to seek new hosts.
• Generally, life cycle of entomopathogenic nematodes (from infective juvenile penetration to infective juvenile emergence) is completed within 12- 15 days at room temperature. The optimum temperature for growth and reproduction of nematodes is between 25 and 30⁰C.

Known species of Heterorhabditis Poinar, 1976 with a biocontrol potential- Identification based on morphological and molecular techniques

1. Heterorhabditis amazonensis Andalo, Nguyen, & Moino, 2006
2. H. argentinensis Stock, 1993
3. H. bacteriophora Poinar, 1976
4. H. baujardi Phan, Subbotin, Nguyen & Moens, 2003
5. H. brevicaudis Liu, 1994
6. H. downesi Stock, Griffin & Burnell, 2002
7. H. floridensis Nguyen, Gozel, Koppenhofer, & Adams, 2006
8. H. georgiana Nguyen, Shapiro-Ilan, & Mbata, 2008
9. H. hambletoni (Pereira, 1937) Poinar, 1976
10. H. hawaiiensis Gardner, Stock & Kaya, 1994
11. H. heliothidis (Khan, Brooks & Hirschman, 1976) Poinar, Thomas & Hess, 1977
12. H. hepialius Stock, Strong & Gardner, 1996
13. H. hoptha (Turco, 1970), Poinar, 1979
14. H. indica Poinar, Karunakar & David, 1992
15. H. marelata Liu & Berry, 1996
16. H. megidis Poinar, Jackson & Klein, 1988
17. H. mexicana Nguyen, Shapiro-Ilan, Stuart, MCCoy, James & Adams, 2004
18. H. poinari Kakulia & Mikaia, 1997
19. H. safricana Malan, Nguyen, deWaal, & Tiedt, 2008
20. H. taysearae Shamseldean, El-Sooud, Abd-Elgawad & Saleh, 1996
21. H. zealandica Poinar, 1990

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Known species of Steinernema Travassos, 1927 with a biocontrol potential- Identification based on morphological and molecular techniques

1. Steinernema abbasi Elawad, Ahma & Reid, 1997
2. S. aciari Qiu, Yan, Zhou, Nguyen & Pang, 2004
3. S. affine (Bovien, 1937) Wouts, Mrácek, Gerdin & Bedding, 1982
4. S. akhursti Qiu, Hu, Zhou, Mei, Nguyen, & Pang, 2005
5. S. anatoliense Hazir, Stock & Keskin, 2003
6. S. apuliae Triggiani, Mracek & Reid, 2004
7. S. arenarium (Artyukhovsky, 1967) Wouts, Mrácek, Gerdin & Bedding, 1982
8. S. ashiuense Phan, Takemoto & Futai, 2006
9. S. asiaticum Shahina, Reid & Rowe, 2002
10. S. backanense Phan, Spiridonov, Subbotin & Moens, 2006
11. S. beddingi Qiu, Hu, Zhou, Pang & Nguyen, 2005
12. S. bicornutum Tallosi, Peters & Ehlers 1995
13. S. carpocapsae (Weiser, 1955) Wouts, Mrácek, Gerdin & Bedding, 1982
14. S. caudatum Xu, Wang & Li, 1991
15. S. ceratophorum Jian, Reid & Hunt 1997
16. S. cholashanense Nguyen, Puža & Mrácek, 2008
17. S. costaricense Uribe, Mora & Stock, 2007
18. S. cubanum Mrá¡cek, Hernandez & Boemare, 1994
19. S. cumgarense Phan, Spiridonov, Subbotin & Moens, 2006
20. S. diaprepesi Nguyen, & Duncan, 2002
21. S. eapokense Phan, Spiridonov, Subbotin & Moens, 2006
22. S. feltiae (Filipjev, 1934) Wouts, Mrácek, Gerdin & Bedding, 1982
23. S. glaseri (Steiner, 1929) Wouts, Mracek, Gerdin & Bedding, 1982
24. S. guangdongense Qiu, Fang, Zhou, Pang, & Nguyen, 2004
25. S. hebeiense Chen, Li, Yan, Spiridonov & Moens 2006
26. S. hermaphroditum Stock, Griffin, & Chaerani, 2004
27. S. intermedium (Poinar, 1985) Mamiya, 1988
28. S. jollieti Spiridonov, Krasomil-Osterfeld & Moens, 2004
29. S. karii Waturu, Hunt & Reid, 1997
30. S. khoisanae Nguyen, Malan, & Gozel, 2006
31. S. kraussei (Steiner, 1923) Travassos, 1927
32. S. kushidai Mamiya, 1988
33. S. leizhouense Nguyen, Qiu, Zhou, & Pang, 2006
34. S. litorale Yoshida, 2004
35. S. loci Phan, Nguyen & Moens, 2001
36. S. longicaudum Shen & Wang, 1992
37. S. monticolum Stock, Choo & Kaya, 1997
38. S. neocurtillae Nguyen & Smart, 1992
39. S. oregonense Liu & Berry, 1996
40. S. pakistanense Shahina, Anis, Reid, Rowe & Maqbool, 2001
41. S. puertoricense Roman & Figueroa, 1994
42. S. puntauvense Uribe, Mora & Stock, 2007
43. S. rarum (Doucet, 1986) Mamiya, 1988
44. S. riobrave Cabanillas, Poinar & Raulston, 1994
45. S. ritteri de Doucet & Doucet, 1992
46. S. robustispiculum Phan, Subbotin, Waeyenberge, & Moens, 2005
47. S. sangi Phan, Nguyen & Moens, 2001
48. S. sasonense Phan, Spiridonov, Subbotin & Moens, 2006
49. S. scapterisci Nguyen & Smart, 1992
50. S. scarabaei Stock & Koppenhöfer 2003
51. S. serratum Liu, 1992
52. S. siamkayai Stock, Somsook & Kaya, 1998
53. S. sichuanense Mrácek, Nguyen, Tailliez, Boemare & Chen, 2006
54. S. silvaticum Sturhan, Spiridonov & Mracek, 2005
55. S. tami Luc, Nguyen, Reid & Spiridonov, 2000
56. S. texanum Nguyen, Stuart, Andalo, Gozel, & Roger, 2007
57. S. thanhi Phan, Nguyen & Moens, 2001
58. S. thermophilum Ganguly & Singh, 2000
59. S. websteri Cutler & Stock, 2003
60. S. weiseri Mrácek, Sturhan & Reid, 2003
61. S. yirgalemense Nguyen, Tesfamariam, Gozel, Gaugler, & Adams, 2005

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Entomopathogenic nematodes (EPNs) of the two genera Steinernema Travassos, 1927 and Heterorhabditis Poinar, 1976 in the order Rhabdita kill most insects but they are harmless to some beneficial insects (e.g. honey bees), higher animals and environment.

Third-stage juvenile is the only free-living stage in the life cycle of the nematode known as the infective juvenile or dauer juvenile that found in soil and can seek, infect and kill their insect hosts.

These infective juveniles are mutualistically associated with symbiotic bacteria (Xenorhabdus spp. or Photorhabdus spp.) in the family Enterobacteriaceae, which are capable of causing disease in insect pests and killing them.

Species of genus, Xenorhabdus are specifically assocaited with the members of the nematode genusSteinernema and Photorhabdus species are associated with the members of nematode genusHeterorhabditis.

In this mutualistic relationship, the nematode infective juveniles provides protection for bacterium outside the insect host (as bacterium is unable to survive in the outside environment i.e. soil or water) and a means of transmission to new hosts and in return bacteria provides nutrients required for the nematode development and reproduction.

Infective juveniles are adapted to remain in the soil environment without feeding until they find a suitable host.

They are also resistant to unfavorable environmental conditions such as desiccation, heat and freezing.

EPNs can infect soil dwelling stages of butterflies, moths, beetles, flies, crickets and grasshoppers.

Infective juveniles of different nematode species employ different foraging strategy to find and infect their insect hosts. For example, Heterorhabditis bacteriophora is a cruiser forager meaning that it actively finds out or hunts its prey, Steinernema carpocapsae is an ambusher forager that sits and wait for a pray to pass by and S. feltiae and S. rivobrave are intermediate foragers.

EPNs are now commercially produced using both in vivo (in living host) and in vitro (in artificial medium) techniques.

Since EPNs have a wide host range, they are currently used as potential biological control agents to manage insect pests of many field crops, greenhouse and nursery plants, horticultural crops, turfgrass, and in some instances insect pests of animals and humans.

EPNs also have a potential to use as biocontrol agents against plant-parasitic nematodes.

Commercially produced nematode infective juveniles can be stored for extended periods and easily applied in aqueous suspensions in the field using traditional sprayers.

Also, EPNs are compatible with several chemical fungicides, insecticides, nematicides and herbicides, and therefore, they can be easily included in IPM programs.

Under current pesticide regulations, the U.S. Environmental Protection Agency has exempted these biological control agents from registration.