Nematode Information

• 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 spry applications to the surface of plant growing medium or on the plant foliage infecsted 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.

• 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.

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 spry 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 (mouth, anus and breathing pores called spiracles).
• Infective juveniles of Heterorhabditis 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.
  

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

• Acta Entomologia Bohemoslovaca
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• Acta Parasitologica
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• Acta Phytophylacica Sinica
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• Advances in Parasitology
• Agricultural Ecosystems and Environment
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• American Bee Journal
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• Australian Journal of Experimental Agriculture
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• 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
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• Comparative Biochemistry and Physiology B
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• Egyptian Journal of Biological Pest Control
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• Entomologia Expermentalis et Applicata
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• FEMS Microbiology Letters
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• Research and Reviews in Parasitology
• Revista de la Sociedad Entomologica Argentina
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• 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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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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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. backanense- undescribed

11. S. beddingi- undescribed

12. S. bicornutum- X. budapestensis

13. S. carpocapsae- X. nematophila

14. S. caudatum- undescribed

15. S. ceratophorum- undescribed

16. S. cholashanense- undescribed

17. S. cubanum- X. poinarii

18. S. cumgarense- undescribed

19. S. diaprepesi- undescribed

20. S. eapokense- undescribed

21. S. feltiae- X. bovienii

22. S. glaseri- X. poinarii

23. S. guangdongense- undescribed

24. S. hebeiense- undescribed

25. S. hermaphroditum- undescribed

26. S. intermedium - X. bovienii

27. S. jollieti-undescribed

28. S. karii- undescribed

29. S. khoisanae- undescribed

30. S. kraussei- X. bovienii

31. S. kushidai- X. japonica

32. S. leizhouense- undescribed

33. S. litorale- undescribed

34. S. loci- undescribed

35. S. longicaudum- undescribed

36. S. monticolum- undescribed

37. S. neocurtillae- undescribed

38. S. oregonense- undescribed

39. S. pakistanense- undescribed

40. S. puertoricense- X. romanii

41. S. rarum- X. szentirmaii

42. S. riobrave- Xenorhabdus sp

43. S. ritteri- Xenorhabdus sp

44. S. robustispiculum- undescribed

45. S. sangi- undescribed

46. S. sasonense- undescribed

47. S. scapterisci- X. innexi

48. S. scarabaei- X. koppenhoeferi

49. S. serratum- X. ehlersii

50. S. siamkayai- X. stockiae

51. S. sichuanense- X. bovienii

52. S. silvaticum- undescribed

53. S. tami- Xenorhabdus sp

54. S. texanum- undescribed

55. S. thanhi- undescribed

56. S. thermophilum- X. indica

57. S. websteri- undescribed

58. S. weiseri- undescribed

59. S. yirgalemense- undescribed

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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. 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.

2. 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.

3. 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.

4. 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.

5. 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.

6. 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.

7. 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.

8. 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.

9. 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.

10. 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.

11. 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.

12. Jagdale, G. B. and Gordon, R. 1998. Effect of propagation temperatures on temperature tolerances of entomopathogenic nematodes. Fundamental and Applied Nematology 21: 177-183.

13. Jagdale, G. B. and Gordon, R. 1998. Variable expression of isozymes in entomopathogenic nematodes follows laboratory recycling. Fundamental and Applied Nematology 21: 147-155.

14. 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.

15. 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.

16. Jagdale, G. B. and Gordon, R. 1997. Effect of recycling temperature on the infectivity of entomopathogenic nematodes. Canadian Journal of Zoology 75: 2137-2141.

17. 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.

18. 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.

19. 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.

20. Jagdale, G.B. and Gordon, R. 1994. Role of catecholamines in the reproduction of Romanomermis culicivorax. Journal of Nematology 26: 40-45.

21. Jagdale, G.B. and Gordon, R. 1994. Caudal papillae in Romanomermis culicivorax. Journal of Nematology 26: 235-237.

22. Saeb, A. T. M., Jagdale, G. B. and Grewal, P. S. 2008. Genetic variation and sub-structuring in the entomopathogenic nematode genus Heterorhabditis using random amplified polymorphic DNA. Biological Control. (in preparation).

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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