Kill Shore flies (Scatella stagnalis) with Entomopathogenic Nematodes

• The shore fly, Scatella stagnalis (Fallén) (Diptera: Ephydridae) is an important insect pest of greenhouse plants.

• Larvae of these flies mainly feed on blue-green algae grown on the surface of plant growing media, walls, floors, benches, and pots.

• But larvae can also cause a serious damage to tender plant tissues thus reducing quality and productivity of plants.

• The adults are not considered as plant feeders but they are nuisance to people and disseminate pathogens such as Fusarium and Pythium from plant to plant as they disperse through the greenhouse.

• Currently, most growers rely on chemicals that kill host plants such as blue-green algae to reduce the incidence of shore flies. However, this method has not been proved effective in reducing shore fly incidence.

• Biological control agents including Bacillus thuringiensis var. thuringiensis (Bt) and entomopathogenic nematodes have been considered as alternatives to chemical pesticides.

• For successful control of shore flies, entomopathogenic nematodes can be easily applied in water suspension as spray application to the surface of plant growing medium.

• Entomopathogenice nematodes including Heterorhabditis megidis, Steinernema arenarium and Steinernema feltiae when applied at the rate of 50 nematodes/cm2 can cause 94- 100% mortality of shore flies.

How Entomopathogenic Nematodes kill Shore flies

• When the infective juveniles are applied to the surface of plant growing substrate, they start searching for their hosts, in this case shore fly larvae.

• Once a larva has been located, the nematode infective juveniles penetrate into the larval body cavity via natural openings such as mouth, anus and spiracles.

• Infective juveniles of Heterorhabditis spp also enter through the intersegmental members of the larval cuticle.

• Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the larval blood.

• In the blood, multiplying nematode-bacterium complex causes septicemia and kills shore fly larvae usually within 48 h after infection.

• Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new larvae in the potting medium/soil.

For more information on the interaction between entomopathogenic nematodes and leafminers, please read following research and extension publications.

• Foote, B.A. 1977. Utilization of blue-breen algae by larvae of shore flies. Environmental Entomology 6, 812-814.

• Goldberg, N.P. and Stanghellini, M.E. 1990. Ingestion-egestion and aerial transmission of Pythium aphanidermatum by shore flies (Ephydrinae: Scatella stagnalis). Phytopathology 80, 1244-1246.

• Lindquist, R., Buxton, J. and Piatkowski, J. 1994. Biological control of sciarid flies and shore flies in glasshouses. Brighton Crop Protection Conference, Pests and Diseases, BCPC Publications 3, 1067-1072.

• Morton, A., Garcia del Pino, F., 2007. Susceptibility of shore fly Scatella stagnalis to five entomopathogenic nematode strains in bioassays. Biocontrol 52: 533-545.

• Morton, A. and Garcia del Pino, F. 2003. Potential of entomopathogenic nematodes for the control of shore flies (Scatella stagnalis). Growing Biocontrol Markets Challenge Research and Development. 9th European Meeting IOBC/WPRS Working Group "Insect Pathogens and Entomopathogenic Nematodes", Abstracts, 67.

• Vanninen, I., Koskula, H. 2000. Biological control of the shore fly (Scatella tenuicosta) with steinernematid nematodes and Bacillus thuringiensis var. thuringiensis in peat and rockwool. Biocontrol Sci. Technol.. 13: 47-63.

• Zack, R.S. and Foote, B.A. 1978. Utilization of algal monoculture by larvae of Scatella stagnalis. Environmental Entomology 7, 509-511.

Kill Western Flower Thrips with Entomopathogenic Nematodes

• The Western flower thrips, Frankliniella occidentalis is a most economically important pest of many field- and glasshouse-grown vegetables and ornamentals.

• Adults lay eggs in the parenchyma tissue and there are two larval stages (first and second instars), prepupal and pupal stages are present in the life cycle of thrips.

• Adult thrips generally feed by piercing and scraping of the stem, leaf, flower and fruit tissues.

• Both instars also feed on all the aerial plant parts including leaves, flowers and fruits.

• Piercing and scraping of the plant tissues leads to discoloration and drying of the damaged area, in some cases, abortion of flower/leaf buds or distortion of emerging leaves, thus reducing field crop yield and aesthetic value of ornamental plants.

• Thrips are also capable of transmitting tospoviruses such as tomato spotted wilt virus (TSWV) and impatiens necrotic spot virus (INSV) during feeding, thus causing a tremendous loss to agricultural and horticultural greenhouse industries.

• Controlling western flower thrips is difficult because of their small size and cryptic behavior.

• Western flower thrips are commonly eradicated using endosulfan, chlorpyrifos, bendiocarb, and synthetic pyrethrinoids but use of these insecticides is restricted due to their environmental pollution and human health concerns, development of resistance to pesticides and removal of some of the most effective products from the market.

• Biological control agents including predacious mites (Neoseilus cucumeris and Neoseilus degenerans), predacious bugs (Orius insidiosus), entomopathogenic fungi (Beauveria bassiana, Metarhizium anisopliae) and entomopathogenic nematodes (see below) have been used as alternatives to chemical pesticides.

• The entomopathogenic nematodes species including Heterorhabditis bacteriophora, H. indica, H. marelata and Steinernema abassi, S. carpocapase, and S. feltiae have been found to be effective alternatives to chemical insecticides in controlling western flower thrips.

• The entomopathogenic nematodes specifically attack soil-dwelling second instar larval, prepupal and pupal stages.

• Generally, Heterorhabditis species are more effective than Steinernema species nematodes in controlling western flower thrips.

• The insect- parasitic nematodes such as Thripinema nicklewoodii also have a potential to use as a biological control agent against western flower thrips.

• Application of entomopathgenic nematodes at the rate of 400 infective juveniles/ cm2 of soil surface can cause over 50% mortality of thrip population.

• Nematodes can be easily applied in water suspension as spray applications to the surface of plant growing medium or on the plant foliage infested with western flower thrips.

• Although larval stages, prepupae and pupae are susceptible to entomopathogenic nematodes, H. bacteriophora HK3 strain can cause higher mortality of larval and prepupal stages than pupal stages

How Entomopathogenic Nematodes kill Western Flower Thrips

• When the infective juveniles are applied to the surface of plant growing medium or injected in the potting medium, they start searching for their hosts, in this case Western Flower Thrip larvae, prepupae and pupae.

• Once a larvae, prepupae and pupae has been located, the nematode infective juveniles penetrate into the larvae, prepupae and pupae body cavity via natural openings (mouth, anus and spiracles).

• Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub/pupa cuticle.

• Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the larvae, prepupal and pupal blood.

• Multiplying nematode-bacterium complex in the blood causes septicemia and kills the grub usually within 48 h after infection.

• Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new larvae, prepupae and pupae in the potting medium/soil.

Biological Control of Black Vine Weevil using Insect Parasitic Nematodes

• Black vine weevil, Otiorhynchus sulcatus is a common insect pest of over 150 plant species that grown in the greenhouses and nurseries.

• Some of the plant species damaged by black vine weevils include Azalea, Cyclamen, Euonymus, Fuxia, Rosa, Rhododendron and Taxus.

• Grubs (Larvae) of these weevils generally girdle the main stem, and feed and damage roots leading to nutrient deficiencies.

• Adults feed on leaves and flowers by notching their edges thus reducing aesthetic value of plants.

• The entomopathogenic nematodes species including Heterorhabditis bacteriophora, H. megidis and Steinernema carpocapase, S. feltiae and S. glaseri have been found to be effective alternatives to chemical insecticides such as chlorpyrifos (Dursban) in controlling black vine weevils.

• Susceptibility of black vine weevil to nematodes is species and strain specific.

• The rate of application of the nematode species/strains that tested against black vine weevil varies (5,000- 60,000 infective juveniles/pot) among different studies but nematodes applied at the rate of 5000- 20,000 infective juveniles/pot can cause up to 100% grub mortality.

• Nematodes can be easily applied in water suspension as spray applications to the surface of plant growing medium but if nematodes are injected at depths deeper than 5 cm i.e. near to grubs they can cause highest mortality of grubs (70-93%) than those nematodes applied to the surface.

• All the four larval stages (instars) and pupae of black vine weevil are susceptible to all entomopathogenic nematode species.

• However, Heterorhabdtis bacteriophora can cause higher mortality of first and second instars than S. carpocapase and S. glaseri.

• Also, all the three nematodes species are equally effective against third and fourth instars of black vine weevil.

How Entomopathogenic Nematodes Kill Black Vine Weevil

• When the infective juveniles are applied to the surface of plant growing medium or injected in the potting medium, they start searching for their hosts, in this case black vine weevil grubs and pupae.

• Once a grub/pupa has been located, the nematode infective juveniles penetrate into the grub or pupa body cavity via natural openings (mouth, anus and spiracles).

• Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub/pupa cuticle.

• Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the grub blood.

• Multiplying nematode-bacterium complex in the blood causes septicemia and kills the grub usually within 48 h after infection.

• Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new grubs or pupae in the potting medium/soil.

Kill fungus gnats using biological control agents: Insect-parasitic nematodes

• Several fungus gnat species including Bradysia coprophila, B. impatiens and B. difformis are considered economically important indoor and greenhouse pests in Europe and the US.
• Fungus gnat flies are black or gray in color with clear wings, relatively small (3-4 mm) in size and commonly associated with compost and natural soils with high organic contents.
• You can see these hopping flies when you water your plants.
• Fungus gnat maggots (larvae) are white-bodied with black heads and can be found just under the surface of the potting medium/soil.
• These maggots primarily feed on fungi and organic matter but they can also cause a serious damage to many ornamental plants.
• Maggots often chew or strip plant roots and tunnel stems affecting water and nutrient absorption in severely injured plants resulting in lost vigor, turn off-color and eventually death.
• Maggots are also capable of transmitting fungal pathogens (Fusarium, Phoma, Pythium and Verticillium) during feeding.
• Adult flies are nuisance to people and disseminate fungal spores from plant to plant as they disperse through the greenhouse.
• Females often laying over 1000 eggs in a lifetime on the media surface and completing egg-to-egg life cycle within 20-25 days at 20-25oC.
• Continuous and overlapping generations of fungus gnats in the greenhouse have made most control strategies difficult.
• Currently, most growers rely on insecticides to manage fungus gnats in floriculture.
• However, use of these insecticides is restricted due to their environmental pollution and human health concerns, development of resistance to pesticides and removal of some of the most effective products from the market.
• Biological control agents including Bacillus thuringiensis (Bt), the predatory mite, Hypoaspis miles and entomopathogenic nematodes have been used as alternatives to chemical pesticides.
• The entomopathogenic nematodes species including Heterorhabditis bacteriophora GPS11 strain, H. indica LN2 strain and Steinernema feltiae UK strain have a potential to use as biocontrol agents against fungus gnats.
• These nematodes kill both maggots (larvae) and pupae, but the second and fourth stages are most susceptible than pupae.
• Nematodes are generally applied in water suspension as spray applications to the surface of plant growing medium to target larval and pupal stages.
• The potting medium (Ball-mix, Nursery-mix or Pro-mix) can influence the survival, persistence and efficacy of entomopathogenic nematodes in greenhouse production.
• In the Nursery-mix, H. bacteriophora can survive longer and perform better than H. indica, H. marelatus Oregon, H. zealandica X1 and Steinernema feltiae against fungus gnats.
• In the Pro-mix, only H. indica have performed better than all other nematode species that tested against fungus gnats.
• Application of S. feltiae can cause 40% reduction in fungus gnat population in Ball-mix, 50% in Metro-mix and 56% in Pro-mix, but only 27% in the Nursery-mix.
• In the greenhouse, temperature can influence efficacy of nematodes. For example, H. bacteriophora and H. indica can survive and cause very high mortality of fungus gnats at warmer (above 25oC) temperatures whereas S. feltiae is generally effective against fungus gnats at cooler (below 25oC) temperatures.
• Application of an appropriate concentration of nematodes is a crucial step in the cost effective control of fungus gnats in greenhouse production.
• Generally, application of one billion infective juveniles of H. bacteriophora, H. indica or S. feltiae per acre can kill over 50% fungus gnats in greenhouse productions.

How entomopathogenic nematodes kill fungus gnats

• When the infective juveniles are applied to the surface of plant growing medium, they start searching for hosts, in this case fungus gnat maggots (larvae) and pupae.
• Once a maggot/pupa has been located, the nematode infective juveniles penetrate into the maggot body cavity via natural openings such as mouth, anus and breathing pores called spiracles.
• Infective juveniles of Heterorhabditis spp also enter through the intersegmental members of the maggot/pupal cuticle.
• Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the fungus gnat blood.
• Multiplying nematode-bacterium complex causes septicemia and kills the host usually within 48 h after infection.
• Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new maggots in the potting medium/soil.

Nematodes are now commercially available from many suppliers distributed throughout in the USA.

For more information on biological control of fungus gnats, please read following research papers/book chapters:

• Binns, E.S., 1973.  Fungus gnats (Diptera: Mycetophilidae, Sciaridae) and the role of mycophagy in soil: a review. Rev. Ecol. Biol. Sol. 18, 77-90.
• Chambers, R.J., Wright, E.M., Lind, R.J., 1993.  Biological control of glasshouse sciarid larvae (Bradysia spp.) with the predatory mite, Hypoaspis miles on Cyclamen and Poinsettia. Biocontrol Sci. Technol. 3, 285-293.
• Ecke, P.Jr., Faust, J.E., Williams, J., Higgins, A., 2004.  The Poinsettia Manual. Ball Publishing, The Paul Ecke Ranch, Encinitas, California, USA.
• Freeman, P., 1983.  Sciarid flies, Diptera; Sciaridae. Handbooks for the identification of British insects 9, Part 6. London, Royal Entomol. Soc. pp 68.
• Gillespie, D.R., Menzies, J.G., 1993.  Fungus gnat vector Fusarium oxysporum f. sp. radicislycopersici.  Ann. Appl. Biol. 123, 539-544.
• Gouge, D.H., Hague, N.G.M., 1994.  Control of sciarids in glass and propagation houses with Steinernema feltiae. Brighton Crop Protection Conference: Pest Dis. 3, 1073-1078.
• Gouge, D.H., Hague, N.G.M., 1995.  Glasshouse control of fungus gnats, Bradysia paupera, on fuchsias by Steinernema feltiae. Fundam. Appl. Nematol. 18, 77-80.
• Grewal, P.S., Richardson, P.N., 1993.  Effects of application rates of Steinernema feltiae (Nematoda: Steinernematidae) on control of the mushroom sciarid fly, Lycoriella auripila (Diptera: Sciaridae).  Biocontrol Sci. Technol. 3, 29-40.
• Grewal, P.S., Tomalak, M., Keil, C.B.O., Gaugler, R., 1993. Evaluation of a genetically selected strain of Steinernema feltiae against the mushroom sciarid fly, Lycoriella mali. Ann. Appl. Biol. 123, 695-702.
• Harris, M.A., Oetting, R.D., Gardner, W.A., 1995.  Use of entomopathogenic nematodes and new monitoring technique for control of fungus gnats, Bradysia coprophila (Diptera: Sciaridae), in floriculture. Biol. Control 5, 412-418.
• 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. Biol. Contrl. 29: 296-305.
• 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. Biol. Control. 43: 23-30.
• Kim, H.H., Choo, H.Y., Kaya, H.K., Lee, D.W., Lee, S.M., Jeon, H.Y., 2004.  Steinernema carpocapsae (Rhabditida: Steinernematidae) as a biological control agent against the fungus gnat Bradysia agrestis (Diptera: Sciaridae) in propogation houses. Biocontrol Sci. Technol. 14, 171-183.
• Lindquist R., Piatkowski J. 1993. Evaluation of entomopathogenic nematodes for control of fungus gnat larvae. Bull. Int. Organiz. Biol. Integr. Control Noxious Animals and Plants. 16, 97-100.
• Lindquist, R.K., Faber, W.R., Casey, M.L., 1985.  Effect of various soilless root media and insecticides on fungus gnats.  HortScience. 20, 358-360.
• Menzel, F., Smith, J.E., Colauto, N.B., 2003.  Bradysia difformis Frey and Bradysia ocellaris (Comstock): two additional neotropical species of black fungus gnats (Diptera : Sciaridae) of economic importance: a redescription and review. Ann. Entomol. Soc. Am. 96, 448-457.
• Nielsen, G. R., 2003. Fungus gnats. http://www.uvm.edu/extension/publications/el/el50.htm
• Oetting, R.D., Latimer, J.G., 1991.  An entomogenous nematode Steinernema carpocapsae is compatible with potting media environments created by horticultural practices. J. Entomol. Sci. 26, 390-394.
• Olson, D.L., Oetting, R.D., van Iersel, M.W., 2002.  Effect of soilless media and water management on development of fungus gnats (Diptera: Sciaridae) and plant growth. HortScience. 37: 919-923.
• Richardson, P.N., Grewal, P.S., 1991.  Comparative assessment of biological (Nematoda: Steinernema feltiae) and chemical methods of control of mushroom fly, Lycoriella auripila (Diptera: Sciaridae).  Biocontrol Sci. Technol. 1, 217-228.
• 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.
• Wilkinson, J.D., Daugherty, D.M., 1970.  Comparative development of Bradysia impatiens (Diptera: Sciaridae) under constant and variable temperatures. Ann. Entomol. Soc. Am. 63, 1079-1083.

Published Books and Related Literature on Beneficial Nematodes: Insect- and Slug- parasitc nematodes

• Acarology IX. Edited by Roger, M., Horn, D.J., Needham, G.R., Welbourn, W.C., Ohio Biological Survey, Columbus, Ohio (1996).
• A Handbook of Biology and Techniques. By Woodring, J.L., Kaya, H.K., Sothern Cooperative Bulletin 331, Arkansas Agricultural Experiment Station, Fayettville, Arkansas (1988).
• Annual Report, CSIRO Division of Entomology (1981).
• A Taxonomic Review of the Suborder Rhabditina (Nematoda: Secernentia) by Andrassy, I. ORSTROM, Paris (1983).
• A worldwide guide to beneficial animals (insects, mites, nematodes) used for pest control purposes. By Thomson, W.T, Thomson Publications; Fresno, CA (1992).
• Bioassays of Entomopathogenic Microbes and Nematodes. Edited by Navon, A., Ascher, K.R.S., CAB International, Wallingford, UK (2000).
• Biological Control: Benefits and Risks. Edited by Hokkanen, H.M.T., Lynch, J.M., Cambridge University Press, UK (1995).
• Biological Control of Mosquitoes. Edited by Chapman, H.C., American Mosquito Control Association Bulletin 6 (1985).
• Biorational Pest Control Agents: Formulation and Delivery. Edited by Hall, F.R., Barry, J.W., American Chemical Society, Maryland (1995).
• Cranberry Research Compilation. Progress and Final Reports on Cranberry Research Conducted in 1998. Edited by Deziel,G., Hogan, M., Cranberry Institute, Warenham, Massachsetts (1999).
• Conservation Biological Control. Edited by Barbosa, P., Academic Press, San Diego, California (1998).
• Control of Insect Pests with Entomopathogenic Nematodes. edited by Smith, K.A., Hatsukade, M., Food and Fertilizer Technology Center, Republic of China in Taiwan (1994).
• Cost Action 819 Developments in Entomopathogenic Nematode/Bacterial Research. European Commission, Luxembourg (2001).
• Cost 819 Entomopathogenic Nematode: Application and Pesistence of Entomopathogenic nematodes. European Communities, Luxembourg (1999).
• Cost 819 Biotechnology-Ecology and Transmission Strategies of Entomopathogenic Nematodes. EOfficial Publication of EC, Luxembourg, EN 16269 EN (1995).
• Cost 819. Entomopathogenic Nematodes- Pathogenicity of Entomopathogenic Nematodes Versus Insect Defence Mechanisms: Impact on Selection of Virulent Strains. Edited by Simoes, N. Boemare, N & Ehlers, R., Office for Official Publications of the European Communities (1988).
• Ecology and Biology of Soil Organisms. Edited by Bhandari, S.C., Somani, L.L., Agrotech Publishing Academy, Udaipur (1994).
• Ecology and Classification of North American Freshwater Invertebrates, 2nd edn., Edited by Thorp, J.H., Covich, A.P., Academic Press, San Diego, California (2001).
• Ecology and Transmission Strategies of Etomopathogenic Nematodes. Edited by Griffin, C.T., Gwynn, R.L., Masson, J.P., European Commission, Luxembourg (1995).
• Entomogenous Nematodes, A Manual and Host List Insect- Nematode Associations. By Poinar, G.O.Jr., E.J. Brill, Leiden, The Netherlands (1975).
• Entomopathogenic Nematodes in Biological Control. Edited by Gaugler, R., Kaya, H.K., CRC Press, Boca Raton, Florida (1990).
• Entomopathogenic Nematodes: Systematics, Phylogeny and Bacterial Symbionts. Edited by Nguyen, K.B. & Hunt, D. Brill, The Neitherland, (2007).
• Entopathogenic Nematology. Edited by Gaugler, R., CAB International, Wallingford, UK (2002).
• Environmental Persistance of Entomopathogens and Nematodes. Edited by Baur, M.E., Kaya, H.K., Fuxa, J., Southern Cooperative Series Bulletin 398, Oklahoma Agricultural Experiment Station, Stillwater, Oklahoma (2001).
• Genetics of Entomopathogenic Nematode Bacterium-Complexes. European Commission, Luxembourg City, Luxembourg (1994).
• Field Manual of Techniques in Invertebrate Pathology. Edited by Lacy, L.A., Kaya, H.K., Kluwer Academic Publishers, The Netherlands (2000).
• Formulation of Microbial Biopesticides - Beneficial Microorganisms, Nematodes, Seed Treatments. Edited by Burges, H.D. 1998.Kluwer Academic Pub. Boston and Dordrecht 496p (1998).
• Helminths of Insects. Edited by Sonin, M.D., Amerind Publishing, New Delhi (1987).
• Integrated Pest Management for Turfgrass and Ornamentals. Edited by Leslie, A.R., CRC Press, Boca Raton, Florida (1994).
• Manual of Agricultural Nematology. Edited by Nickle, W.R., Marcel Dekker, New York (1991).
• Manual of Techniques in Insect Pathology. Biological Techniques Series. Edited by Lacey, L.A., Academic Press, San Diego, California (1997).
• Manual of Techniques in Invertebrate Pathology. Edited by Lacey, L.A., Kaya, H.K., Kluwer Academic Publishers, Dordrecht, The Netherlands (2000).
• Methods in Biotechnology: Biopesticides: Use and Delivery. Edited by Hall, F.R., Menon, J. Humanan Press Inc, Totowa, New Jersey (1998).
• Microorganismos patógenos empleados en el control microbiano de insectos plaga. Edited by Lecuona, R., Talleres Gráficos Mariano Mas, Buenos Aires (1996).
• Natural Enemies of Terrestrial Molluscs. Edited by Barker, G.M., CAB International, Wallingford, UK (2003).
• Nematodes and the Biological Control of Insect Pests. Edited by Bedding, R.A., Akhurst, R., Kaya, H.K., CSIRO Publications, East Melbourne, Australia (1993).
• Nematodes as Biocontrol Agents. Edited by Grewal P. S., Ehlers, R.-U., Shapiro-Ilan, D.I., CABI Publishing, CAB International, Wallingford, UK (2005).
• Nematodes for Biological Control of Insects. Edited by Poinar, G.O.Jr., CRC Press, Boca Raton, Florida (1979).
• Nematology, Advances and Perspective. Edited by Chen, Z.X., Chen, S.Y. and Dickson, D.W.,Tsinghua University Press, TUP, China (2001).
• New Directions in Biological Control. Edited by Baker, R.P., Dunn, P.E., Liss, New York (1990).
• Plant and Insect Nematodes. Edited by Nickle, W. R., Marcel Dekker, New York (1984).
• Parasites and Pathogens of Insects. Edited by Beckage, N.E., Thompson, S.N., Federici, B. Academic Press, New York (1993).
• Pest management in the subtropics. Biological control - A Florida perspective. Edited by Rosen, D., Bennett, F.D. and Capinera, J.L., Intercept, Andover, UK (1994).
• Proceedings of Workshop on Optimal Use of Insecticidal Nematodes in Pest Management. Edited by Polavarapu, S., Rutgers University, New Brunswick, New Jersey (1999).
• Recent Advances in Biological Control of Insect Pests by Entomogenous Nematodes in Japan. Edited by Ishibashi, N., Saga University, Japan (1987).
• Recent Advances in Nematology. Edited by Dwivedi, B.K., Bioved Research Society, Allahabad, India (1992).
• Slug and Snail Pests in Agriculture. Edited by Henderson, I., Symposium Proceedings No. 66, British Crop Protection Council Farnham, UK (1996).
• Slugs & Snails: Agricultural, Veterinary & Environmental Perspectives. Monograph No. 80, British Crop Protection Council, Thornton Health, UK (2003).
• Survival Strategies of Entomopathogenic Nematodes. Edited by Glazer, I., Richardson, P., Boemare, N., Coudert, F., EUR 18855 EN Report (1999).
• The Biology of Nematodes. Edited by Lee, D.L., Taylor & Francis, London (2002).
• The Natural History of Nematodes by Pionar, G.O.Jr., Prentice-Hall, Englewood Cliffs, New Jersey (1983).
• Tick-Born Diseases and their Vectors. Edited by Dusbabek, R., Bukva, V., SPB Academic Publishers, The Hague, The Netherlands (1991).
• Tick Born Pathogens at the Host Vector Interface: A Global Perspective. Edited by Coons, L., Rothschild, M., Krugel, National Park, South Africa (1995).
• Tortricid Pests, Their Biology, Natural Enemies and Control. Edited by Van der Geest,L.P.S., Evenhius, H.H., elsevier Science, Amsterdam, The Netherlands (1991).
• Tylenchida Parasites of Plants and Insects. By Siddiqi, M.R., CAB Farnham Royal, Slough, UK (1986).
• Tylenchida Parasites of Plants and Insects. By Siddiqi, M.R., CAB International, Wallingford, UK (2000).
• Use of Pathogens in Scarab Pest Management. Edited by Jackson, T.A., Glare, T.R., Ag Research, Lincoln, New Zealand (1992).

Visit this blog again for new updates (if any) on books & related literature on insect and slug parasitic nematodes

List of Research and Extension Journals Accepting Papers on Beneficial Nematodes

• Acta Entomologia Bohemoslovaca
• Acta Horticulturae
• Acta Parasitologica
• Acta Phytopathologica et Entomologica Hungarica
• Acta Phytophylacica Sinica
• Actes des Colloques Insectes Sociaux
• Advances in Parasitology
• Agricultural Ecosystems and Environment
• Agricultural Systems
• Agro Food Industry Hi-Technology
• American Bee Journal
• American Naturalist
• Annals of Applied Biology
• Annals of the Entomological Society of America
• Annals of the New York Academy of Science
• Annual Review of Entomology
• Annual Review of Microbiology
• Annual Review of Phytopathology
• Applied Entomology and Zoology
• Applied and Environmental Microbiology
• Applied Microbiology and Biotechnology
• Applied Soil Ecology
• Arthropod Management Tests
• Australian Journal of Experimental Agriculture
• Behaviour
• Biocontrol Science and Technology
• Biocontrol
• Biodiversity and Conservation
• Biological Control
• Biology and Fertility of Soils
• Biotechnology and Bioengineering
• Biotechnology Progress
• Brighton Crop Protection Conference- Pest and Disease
• Bulletin of Entomological Research
• Bulletin of Entomological Society of America
• Bulletin of the Faculty of Agriculture, Saga University
• Bulletin of the Georgian Academy of Sciences
• Bulletin of the Institute of Maritime Tropical Medicine, Gdynia
• Bulletin of the Institute of Zoology, Academia Sinica
• Bulletin of the International Organization for Biological and Integrated Control of Noxious Animals and Plants
• Bulletin OILB/SROP
• California Agriculture
• Canadian Entomologist
• Canadian Journal of Zoology
• Cellular and Molecular Life Sciences
• Chinese Journal of Tropical Crops
• Comparative Biochemistry and Physiology B
• Crop Protection
• Current Genetics
• Current Opinion in Microbiology
• Ecology
• Egyptian Journal of Biological Pest Control
• Egyptian Journal of Agronematology
• Entomologia Expermentalis et Applicata
• Entomophaga
• Environmental Entomology
• Experientia
• Experimental and Applied Acarology
• Experimental Parasitology
• FEMS Microbiology Letters
• Florida Entomologist
• Folia Parasitologica
• Forest Ecology and Management
• Forest Research
• Gene
• Indian Journal of Agricultural Sciences
• Indian Journal of Entomology
• Indian Journal of Nematology
• Insect Science and Its Application
• Israel Journal of Medical Science
• Integrated Pest Management Reviews
• International Journal for Parasitology
• International Journal of Nematology
• International Journal of Parasitology
• International Journal of Systematic Bacteriology
• International Journal of Systematic and evolutionary Microbiology
• International Organization for Biological and Integrated Control Bulletin
• International Research Communications System Medical Science: Microbiology, Parasitology and Infectious Diseases
• IOBC/WPRS Bulletin
• Irish Journal of Agricultural and Food Research
• Japanese Journal of Applied Entomology and Zoology
• Japanese Journal of Nematology
• Journal for Hawaiian and Pacific Agriculture
• Journal of Agricultural Research
• Journal of American Mosquito Control Association
• Journal of Animal Ecology
• Journal of Applied Ecology
• Journal of Applied Entomology- Zeitschrift fur Angewandte Entomologie
• Journal of Arboriculture
• Journal of Australian Entomological Society
• Journal of Bacteriology
• Journal of Economic Entomology
• Journal of Egyptian Society of Parasitology
• Journal of Entomological Science
• Journal of Environmental Horticulture
• Journal of chemical Ecology
• Journal of Clinical Microbiology
• Journal of General Microbiolgy
• Journal of Helminthology
• Journal of Industrial Microbiology and Biotechnology
• Journal of Insect Pathology
• Journal of Invertebrate Pathology
• Journal of Kansas Entomological Society
• Journal of Medical Entomology
• Journal of Molluscan Studies
• Journal of Natural Products
• Journal of Nematology
• Journal of Parasitology
• Journal of Genetic Microbiology
• Journal of the Australian Entomological Society
• Journal of the Entomological Society of British Columbia
• Journal of the Georgia Entomological Society
• Journal of Thermal Biology
• Journal of West China University of Medical Sciences
• Korean Journal of Applied Entomology
• Korean Journal of Turfgrass Science
• Medical and Veterinary Entomology
• Memoirs of the Entomological Society of Canada
• Microbial Ecology
• Molecular Phylogenetics and Evolution
• Mosquito News
• Mushroom News
• Natural Enemies of Insects
• Nature
• Nature Biotechnology
• Nematology
• Nematropica
• Netherlands Journal of Plant Pathology
• New Zealand Entomologist
• New Zealand Journal of Experimental Agriculture
• New Zealand Journal of Zoology
• Oecologia
• Pakistan Journal of Nematology
• Parasitology
• Parasitology Research
• Pedobiologica
• Pest Management Science
• Phytoparasitica
• Phytoprotection
• Plant Protection Quarterly
• Polskie Pismo Entomologiczne
• Proceedings of the American Chemical Society, Division of Environmental Chemistry
• Proceedings of the Florida State Horticultural Society
• Proceedings of the Entomological Society of Washington
• Proceedings of the Helminthological Society of Washington
• Proceedings of the North Central Branch of Entomological Society of America
• Research and Reviews in Parasitology
• Revista de la Sociedad Entomologica Argentina
• Revista de Proteccion Vegetal
• Revue de Nematologie
• Russian Journal of Nematology
• Science
• Sri Lanka Journal of Tea Science
• SPOR/WPRS Bulletin
• Systematic and Applied Microbiolgy
• Systematic Parasitology
• The Canadian Entomologist
• Trends in Parasitology
• Veterinary Dermatology
• Zhonghua Kunchong

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Symbiotic bacteria of Heterorhabdits nematodes- Photorhabdus species

1. Heterorhabditis amazonensis- undescribed
2. H. argentinensis- P. temperata
3. H. bacteriophora- Photorhabdus luminescens subsp. laumondii TT01, P. luminescens kayaii subsp. nov., P. luminescens thracensis subsp. nov., P. temperate
4. H. baujardi- undescribed
5. H. brevicaudis- P. luminescens
6. H. downesi- Photorhabdus sp
7. H. floridensis- undescribed
8. H. georgiana- undescribed
9. H. hambletoni- undescribed
10. H. hawaiiensis- P. luminescens
11. H. heliothidis- undescribed
12. H. hepialius- P. luminescens
13. H. hoptha- undescribed
14. H. indica- P. luminescens
15. H. marelata- P. luminescens
16. H. megidis- P. temperata subsp. temperata XlNach
17. H. mexicana- undescribed
18. H. poinari- Photorhabdus sp
19. H. safricana- undescribed
20. H. taysearae- undescribed
21. H. zealandica- P. temperata

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Symbiotic bacteria of Steinernematid nematodes- Xenorhabdus species

1. Steinernema abbasi- undescribed
2. S. aciari- undescribed
3. S. affine-Xenorhabdus bovienii
4. S. akhursti- undescribed
5. S. anatoliense- undescribed
6. S. apuliae- undescribed
7. S. arenarium- X. kozodoii
8. S. ashiuense- undescribed
9. S. asiaticum- undescribed
10. S. australe- X. magdalenensis
11. S. backanense- undescribed
12. S. beddingi- undescribed
13. S. bicornutum- X. budapestensis
14. S. carpocapsae- X. nematophila
15. S. caudatum- undescribed
16. S. ceratophorum- undescribed
17. S. cholashanense- undescribed
18. S. cubanum- X. poinarii
19. S. cumgarense- undescribed
20. S. diaprepesi- undescribed
21. S. eapokense- undescribed
22. S. feltiae- X. bovienii
23. S. glaseri- X. poinarii
24. S. guangdongense- undescribed
25. S. hebeiense- undescribed
26. S. hermaphroditum- undescribed
27. S. intermedium - X. bovienii
28. S. jollieti-undescribed
29. S. karii- undescribed
30. S. khoisanae- undescribed
31. S. kraussei- X. bovienii
32. S. kushidai- X. japonica
33. S. leizhouense- undescribed
34. S. litorale- undescribed
35. S. loci- undescribed
36. S. longicaudum- undescribed
37. S. monticolum- undescribed
38. S. neocurtillae- undescribed
39. S. oregonense- undescribed
40. S. pakistanense- undescribed
41. S. puertoricense- X. romanii
42. S. rarum- X. szentirmaii
43. S. riobrave- Xenorhabdus sp
44. S. ritteri- Xenorhabdus sp
45. S. robustispiculum- undescribed
46. S. sangi- undescribed
47. S. sasonense- undescribed
48. S. scapterisci- X. innexi
49. S. scarabaei- X. koppenhoeferi
50. S. serratum- X. ehlersii
51. S. siamkayai- X. stockiae
52. S. sichuanense- X. bovienii
53. S. silvaticum- undescribed
54. S. tami- Xenorhabdus sp
55. S. texanum- undescribed
56. S. thanhi- undescribed
57. S. thermophilum- X. indica
58. S. websteri- undescribed
59. S. weiseri- undescribed
60. S. yirgalemense- undescribed

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Beneficial Nematodes: Steinernema and Heterorhabditis species

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 on Entomopathogenic 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.

Symbiotic bacterial genus, Photorhabdus

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.luminescens subsp. sonorensis, Orozco, Hill & Stock, 2013
8. P. temperata sp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
9. P. temperata subsp. temperata subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
10. P. luminescens subsp. thracensis subsp. nov., Hazir, Stackebrandt, Lang, Schumann, Ehlers & Keskin, 2004

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Symbiotic bacterial genus, Xenorhabdus Thomas and Poinar 1979

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. magdalenensis, Tailliez, Pages, Edgington, Tymo, & Buddie, 2012
15. X. mauleonii Tailliez, Pagès, Ginibre & Boemare, 2006
16. X. miraniensis Tailliez, Pagès, Ginibre & Boemare, 2006
17. X. nematophila (Poinar and Thomas 1965) Thomas and Poinar 1979
18. X. poinarii (Akhurst 1983) Akhurst and Boemare 1993
19. X. romanii Tailliez, Pagès, Ginibre & Boemare, 2006
20. X. stockiae Tailliez, Pagès, Ginibre & Boemare, 2006
21. X. szentirmaii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005

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Life cycle of entomopathogenic nematodes (EPNs)

 

Entomopathogenic nematode life cycle

• 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.

  

Species of the genus Heterorhabditis Poinar, 1976

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. atacamensis, Edgington, Buddie, Moore, France, Merino, & Hunt, 2011
4. H. bacteriophora Poinar, 1976
5. H. baujardi Phan, Subbotin, Nguyen & Moens, 2003
6. H. brevicaudis Liu, 1994
7. H. downesi Stock, Griffin & Burnell, 2002
8. H. floridensis Nguyen, Gozel, Koppenhofer, & Adams, 2006
9. H. georgiana Nguyen, Shapiro-Ilan, & Mbata, 2008
10. Heterorhabditis gerrardi, Plichta, Joyce, Clarke, Waterfield, & Stock, 2009
11. H. hambletoni (Pereira, 1937) Poinar, 1976
12. H. hawaiiensis Gardner, Stock & Kaya, 1994
13. H. heliothidis (Khan, Brooks & Hirschman, 1976) Poinar, Thomas & Hess, 1977
14. H. hepialius Stock, Strong & Gardner, 1996
15. H. hoptha (Turco, 1970), Poinar, 1979
16. H. indica Poinar, Karunakar & David, 1992
17. H. marelata Liu & Berry, 1996
18. H. megidis Poinar, Jackson & Klein, 1988
19. H. mexicana Nguyen, Shapiro-Ilan, Stuart, MCCoy, James & Adams, 2004
20. H. poinari Kakulia & Mikaia, 1997
21. H. safricana Malan, Nguyen, deWaal, & Tiedt, 2008
22. Heterorhabditis sonorensis, Stock, Rivera-Orduno, & Flores-Lara, 2009
23. H. taysearae Shamseldean, El-Sooud, Abd-Elgawad & Saleh, 1996
24. H. zealandica Poinar, 1990

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Species of the genus Steinernema Travassos, 1927

Known species of Steinernema Travassos, 1927 with a biocontrol potential- Identification was 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. australe, Edgington, Buddie, Tymo, Hunt, Nguyen, France, Merino, & Moore, 2009
11. S. backanense Phan, Spiridonov, Subbotin & Moens, 2006
12. S. balochiense Fayyaz, Khanum, Ali, Solangi, Gulsher & Javed, 2015
13. S. beddingi Qiu, Hu, Zhou, Pang & Nguyen, 2005
14. S. bicornutum Tallosi, Peters & Ehlers 1995
15. S. brazilense, Nguyen, Ginarte, Leite, dos Santos, & Harakava, 2010
16. S. carpocapsae (Weiser, 1955) Wouts, Mrácek, Gerdin & Bedding, 1982
17. S. caudatum Xu, Wang & Li, 1991
18. S. ceratophorum Jian, Reid & Hunt 1997
19. S. cholashanense Nguyen, Puža & Mrácek, 2008
20. S. citrae Stokwe, Malan, Nguyen, Knoetze, & Tiedt, 2011
21. S. costaricense Uribe, Mora & Stock, 2007
22. S. cubanum Mrá¡cek, Hernandez & Boemare, 1994
23. S. cumgarense Phan, Spiridonov, Subbotin & Moens, 2006
24. S. dharanaii , Kulkarni, Rizvi, Kumar, Paunikar& Mishra, 2012
25. S. diaprepesi Nguyen, & Duncan, 2002
26. S. eapokense Phan, Spiridonov, Subbotin & Moens, 2006
27.  S. fabii Abate, Malan, Tiedt, Wingfield, Slippers, Hurley, 2016. 
28. S. feltiae (Filipjev, 1934) Wouts, Mrácek, Gerdin & Bedding, 1982
29. S. glaseri (Steiner, 1929) Wouts, Mracek, Gerdin & Bedding, 1982
30. S. guangdongense Qiu, Fang, Zhou, Pang, & Nguyen, 2004
31. S. hebeiense Chen, Li, Yan, Spiridonov & Moens, 2006
32. S. hermaphroditum Stock, Griffin, & Chaerani, 2004
33. S.  innovation Cimen, Lee, Hatting, Hazir, Stock 2015
34. S. intermedium (Poinar, 1985) Mamiya, 1988
35. S. jeffreyense Malan, Knoetze & Tiedt, 2016
36. S. jollieti Spiridonov, Krasomil-Osterfeld & Moens, 2004
37. S. karii Waturu, Hunt & Reid, 1997
38. S. khoisanae Nguyen, Malan, & Gozel, 2006
39. S. kraussei (Steiner, 1923) Travassos, 1927
40. S. kushidai Mamiya, 1988
41. S. leizhouense Nguyen, Qiu, Zhou, & Pang, 2006
42. S. litorale Yoshida, 2004
43. S. loci Phan, Nguyen & Moens, 2001
44. S. longicaudum Shen & Wang, 1992
45. S. monticolum Stock, Choo & Kaya, 1997
46. S. neocurtillae Nguyen & Smart, 1992
47. S. oregonense Liu & Berry, 1996
48. S. pakistanense Shahina, Anis, Reid, Rowe & Maqbool, 2001
49. S. papillatum  San-Blas, Portillo, Nermut, Puza, & Morales-Montero 2015
50. S. phyllophagae Nguyen and Buss, 2011
51. S. puertoricense Roman & Figueroa, 1994
52. S. puntauvense Uribe, Mora & Stock, 2007
53. S. rarum (Doucet, 1986) Mamiya, 1988
54. S. riobrave Cabanillas, Poinar & Raulston, 1994
55. S. ritteri de Doucet & Doucet, 1992
56. S. robustispiculum Phan, Subbotin, Waeyenberge, & Moens, 2005
57. S. sangi Phan, Nguyen & Moens, 2001
58. S. sasonense Phan, Spiridonov, Subbotin & Moens, 2006
59. S. scapterisci Nguyen & Smart, 1992
60. S. scarabaei Stock & Koppenhöfer 2003
61. S. serratum Liu, 1992
62. S. siamkayai Stock, Somsook & Kaya, 1998
63. S. sichuanense Mrácek, Nguyen, Tailliez, Boemare & Chen, 2006
64. S. silvaticum Sturhan, Spiridonov & Mracek, 2005
65. S. tami Luc, Nguyen, Reid & Spiridonov, 2000
66. S. texanum Nguyen, Stuart, Andalo, Gozel, & Roger, 2007
67. S. thanhi Phan, Nguyen & Moens, 2001
68. S. thermophilum Ganguly & Singh, 2000
69. S. websteri Cutler & Stock, 2003
70. S. weiseri Mrácek, Sturhan & Reid, 2003
71. S. xinbinense Ma, Chen, De Clercq, Waeyenberge, Han & Moens, 2012
72. S. xueshanense, Mracek, Liu, & Nguyen, 2009
73. S. yirgalemense Nguyen, Tesfamariam, Gozel, Gaugler, & Adams, 2005

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Entomopathogenic Nematode Facts

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.