Control oriental beetles, Anomala orientalis with an entomopathogenic nematode Steinernema scarabaei / January 18, 2010 by Ganpati Jagdale

The oriental beetle, Anomala orientalis is one of most damaging white grub species of turfgrass. An entomopathogenic nematode, Steinernema scarabaei has been used as effective biological control agent against these beetles. When infective juveniles of this nematode applied at the rate of 2.5 billion per hectare of turfgrass they can suppress over 77% population of oriental beetles (Koppenhofer and Fuzy, 2009). For more information on the effects of entomopathogenic nematodes on different species of white grubs.

Alm, S.R., Yeh, T., Hanula, J.L. and Georgis, R. 1992. Biological control of japanese, oriental and black turfgrass ataenius beetel (Coleoptera, Scarabidae) larvae with entomopathogenic nematodes (Nematoda, Steinernematidae, Heterorhabditidae). Journal of Economic Entomology. 85: 1660-1665.

Choo, H.Y., Kaya, H.K., Huh, J., Lee, D.W., Kim, H.H., Lee, S.M. and Choo, Y.M. 2002. Entomopathogenic nematodes (Steinernema spp. and Heterorhabditis bacteriophora) and a fungus Beauveria brongniartii for biological control of the white grubs, Ectinohoplia rufipes and Exomala orientalis, in Korean golf courses. Biocontrol. 47: 177-192.

Koppenhofer, A.M., Brown, I.M., Gaugler, R., Grewal, P.S., Kaya, H.K. and Klein MG. 2000. Synergism of entomopathogenic nematodes and imidacloprid against white grubs: Greenhouse and field evaluation. Biological Control. 19: 245-251.

Koppenhofer, A.M. and Fuzy, E.M. 2009. Long-term effects and persistence of Steinernema scarabaei applied for suppression of Anomala orientalis (Coleoptera: Scarabaeidae). Biological Control. 48: 63-72.

Koppenhofer, A.M. and Fuzy E.M. 2004. Effect of white grub developmental stage on susceptibility to entomopathogenic nematodes. Journal of Economic Entomology. 97: 1842-1849.

Koppenhofer, A.M. and Fuzy, E.M. 2003. Steinernema scarabaei for the control of white grubs. Biological Control. 28: 47-59.

Koppenhofer, A.M. and Fuzy, E.M. 2008. Effect of the anthranilic diamide insecticide, chlorantraniliprole, on Heterorhabditis bacteriophora (Rhabditida : Heterorhabditidae) efficacy against white grubs (Coleoptera : Scarabaeldae). Biological Control. 45: 93-102.

Koppenhofer, A.M., Fuzy, E.M., Crocker, R.L., Gelernter, W.D. and Polavarapu, S. 2004. Pathogenicity of Heterorhabditis bacteriophora, Steinernema glaseri, and S. scarabaei (Rhabditida : Heterorhabditidae, Steinernematidae) against 12 white grub species (Coleoptera : Scarabaeidae). Biocontrol Science and Technology. 14: 87-92.

Koppenhofer, A.M., Cowles, R.S., Cowles, E.A., Fuzy, E.M. and Baumgartner, L. 2002. Comparison of neonicotinoid insecticides as synergists for entomopathogenic nematodes. Biological Control 24: 90-97.

Koppenhofer, A.M., Grewal, P.S. and Fuzy, E.M. 2006. Virulence of the entomopathogenic nematodes Heterorhabditis bacteriophora, Heterorhabditis zealandica, and Steinernema scarabaei against five white grub species (Coleoptera : Scarabaeidae) of economic importance in turfgrass in North America. Biological Control 38: 397-404

Lee, D.W., Choo, H.Y., Kaya, H.K., Lee, S.M., Smitley, D.R., Shin, H.K. and Park, C.G. 2002. Laboratory and field evaluation of Korean entomopathogenic nematode isolates against the oriental beetle Exomala orientalis (Coleoptera : Scarabaeidae). Journal of Economic Entomology. 95: 918-926.

Li, X.Y., Cowles, R.S., Cowles, E.A., Gaugler, R. and Cox-Foster, D.L. 2007. Relationship between the successful infection by entomopathogenic nematodes and the host immune response. International Journal for Parasitology. 37: 365-374.

Mannion, C.M., McLane, W., Klein, M.G., Moyseenko, J., Oliver, J.B. and Cowan D. 2001. Management of early-instar Japanese beetle (Coleoptera : Scarabaeidae) in field-grown nursery crops. Journal of Economic Entomology. 94: 1151-1161.

Polavarapu, S., Koppenhoefer, A.M., Barry, J.D., Holdcraft, R.J. and Fuzy, E.M. 2007. Entomopathogenic nematodes and neonicotinoids for remedial control of oriental beetle, Anomala orientalis (Coleoptera : Scarabaeidae), in highbush blueberry. Crop Protection. 26: 1266-1271.

Yeh, T. and Alm, S.R. 1995. Evaluation of Steinernema glaseri (Nematoda: Steinernematidae) for biological control of japanese and apanese and oriental beetles (Coleoptera, Searabaeidae). Journal of Economic Entomology. 88: 1251-1255.

Yi, Y.K., Park, H.W., Shrestha, S., Seo, J., Kim, Y.O., Shin, C.S. and Kim, Y. 2007. Identification of two entomopathogenic bacteria from a nematode pathogenic to the oriental beetle, Blitopertha orientalis. Journal of Microbiology and Biotechnology. 17: 968-978.

Parasitization of subterranean termite Heterotermes aureus by beneficial nematodes / December 28, 2009 by Ganpati Jagdale

It has been reported that three entomopathogenic nematode species including Steinernema carpocapsae Mexican 33 strain, S. feltiae UK76 strain and Heterorhabditis bacteriophora HP88 strain can infect and kill desert subterranean termite s Heterotermes aureus under laboratory conditions (Yu et al., 2008). These nematodes can also develop and reproduce in termite cadavers and emerge as infective juveniles.

Please read following literature for more information on interaction between insect-parasitic nematodes and termites.

Yu, H., Gouge, D.H., Stock, S.P. and Baker, P.B. 2008. Development of entomopathogenic nematodes (Rhabditida: Steinernematidae; Heterorhabditidae) in desert subterranean termite Heterotermes aureus (Isoptera: Rhinotermitidae). Journal of Nematology. 40: 311-317.

Susceptibility of longicorn beetle (Dorcadion pseudopreissi) to entomopathogenic nematodes / December 27, 2009 by Ganpati Jagdale

Recently, it has been reported that a new insect pest of turf called longicorn beetle (Dorcadion pseudopreissi) was susceptible to three species entomopathogenic nematodes including Steinernema carpocapsae, S. feltiae and Heterorhabditis bacteriophora under laboratory condition. The results of this study suggests that the entomopathogenic nematodes have a potential to use as biological control agents against longicorn beetles (Susurluk et al., 2009). Susurluk, I.A., Kumral, N.A., Peters, A., Bilgili, U. and Acikgoz, E. 2009. Pathogenicity, reproduction, and foraging behaviours of some entomopathogenic nematodes on a new turf pest,

Plants can call for help for their protection against insect pests / December 27, 2009 by Ganpati Jagdale

It has been demonstrated that the plants when attacked by herbivorous insects can emit volatile compounds that can attract natural enemies of the insects. For example, the roots of maize plants when attacked by western corn root-worms (a noxiuos insect pest of corn) can synthesize and emit a volatile compound called (E)-beta-caryophyllene that attracts insect-parasitic nematodes that infect and kill many soil dwelling insect pests (Rasmann et al., 2005; Degenhardt et al., 2009). Read following scientific papers for more information on insect induced plant volatiles that attract natural enemies of insect pests.

Degenhardt, J., Hiltpold, I., Kollner, T.G., Frey, M., Gierl, A., Gershenzon, J., Hibbard, B.E., Ellersieck, M.R. and Turlings, T.C.J. 2009. Restoring a maize root signal that attracts insect-killing nematodes to control a major pest. Proceedings of the National Academy of Sciences of the United States of America. 106: 13213-13218.

Rasmann, S., Kollner, T.G., Degenhardt, J., Hiltpold, I., Toepfer, S., Kuhlmann, U., Gershenzon, J., Turlings T.C.J. 2005. Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature 434: 732–737.

Use insect-parasitic nematodes to control citrus root weevils / December 26, 2009 by Ganpati Jagdale

The citrus root weevil also called as Diaprepes root weevil (Diaprepes abbreviatus) is one of the major insect pests of citrus and many ornamental plants in Florida and California. Several researchers have demonstrated that the application of an insect-parasitic nematode can supress the populations of root weevils in citrus orchards. For example, Steinernema riobrave infective juveniles when applied in citrus orchards or greenhouses can provide 50 to 90% reduction in populations of D. abbreviatus (Bullock et al., 1999; Duncan and McCoy, 1996; Duncan et al., 1996; Shapiro and McCoy, 2000ab). Applications of S. carpocapsae (All strain), Heterorhabditis bacteriophora (HP-88 strain) or H. bacteriophora (Florida strain) in the citrus grove can also reduce 50-70% adult emergence of D. abbreviatus (Duncan et al., 1996; Schroeder, 1992). According to Shapiro et al. (1999), S. riobrave, H. bacteriophora and H. indica were highly virulent against younger (50-day-old) than older (100-day-old) D. abbreviatus larvae at 24 or 27 degrees C temperature. Heterorhabditis indica was more virulent than H. bacteriophora in 50-day-old D. abbreviatus larvae at all temperatures. However, H. bacteriophora was more virulent than S. riobrave in 20-day-old larvae at 24 degrees C but it was less virulent than S. riobrave in 50-day-old larvae at 21 degrees C.

Please Read following literature for detailed information on interaction between insect-parasitic nematodes and citrus root weevil.

Bullock, R.C., Pelosi, R.R. and Killer, E.E. 1999. Management of citrus root weevils (Coleoptera : Curculionidae) on Florida citrus with soil-applied entomopathogenic nematodes (Nematoda : Rhabditida). Florida Entomologist. 82: 1-7.

Duncan, L.W and McCoy, C.W. 1996 Vertical distribution in soil, persistence, and efficacy against citrus root weevil (Coleoptera: Curculionidae) of two species of entomogenous nematodes (Rhabditida: Steinernematidae; Heterorhabditidae). Environmental Entomology. 25: 174-178.

Duncan, L.W. McCoy, C.W. and Terranova, A.C. 1996. Estimating sample size and persistence of entomogenous nematodes in sandy soils and their efficacy against the larvae of Diaprepes abbreviatus in Florida. Journal of Nematology. 28: 56-67.

Schroeder, W.J. 1992. Entomopathogenic nematodes for control of root weevils of citrus. Florida Entomologist 75: 563-567.

Shapiro, D.I. and McCoy, C.W. 2000a. Susceptibility of Diaprepes abbreviatus (Coleoptera : Curculionidae) larvae to different rates of entomopathogenic nematodes in the greenhouse. Florida Entomologist. 83: 1-9.

Shapiro, D.I. and McCoy, C.W. 2000b. Effects of culture method and formulation on the virulence of Steinernema riobrave (Rhabditida: Steinernematidae) to Diaprepes abbreviatus (Coleoptera: Curculionidae). Journal of Nematology 32: 281-288.

Shapiro, D.I., Cate, J. R., Pena, J., Hunsberger, A. and McCoy, C.W. 1999. Effects of temperature and host age on suppression of Diaprepes abbreviatus (Coleoptera : Curculionidae) by entomopathogenic nematodes. Journal of Economic Entomology. 92: 1086-1092.

Biological control of Colorado potato beetle, Leptinotarsa decemlineata with entomopathogenic nematodes / December 16, 2009 by Ganpati Jagdale

Colorado potato beetle, Leptinotarsa decemlineata: This is an economically important pest of potatoes with more than 40 species have been reported from North America. The larvae of this beetle are voracious feeder of potato leaves costing hundreds of millions of dollars for pesticide control and yield loss each year in the United States.

Use entomopathogenic nematodes to control insect pests of peaches (Prunus persicae, Miller) / December 14, 2009 by Ganpati Jagdale

South American fruit fly, Anastrepha fraterculus: It has been demonstrated that an entomopathogenic nematode Heterorhabditis bacteriophora when applied at the concentration of 250 infective juveniles per square cm in the field can cause 28 to 51% mortality of South American fruit fly larvae.

Use insect parasitic nematodes to manage western corn rootworms (Diabrotica virgifera virgifera) / December 8, 2009 by Ganpati Jagdale

The western corn rootworm (Diabrotica virgifera virgifera) is a very serious pest of corn in the North America and Europe. Larvae of this insect exclusively feed on maize roots, often causing plant lodging whereas adults may reduce yields through silk feeding and interfering maize pollination.

Biological control of various insect pests with entomopathogenic nematode S. carpocapsae / November 23, 2009 by Ganpati Jagdale

Apopka weevil (Diaprepes abbreviatus): This insect was named as Apopka weevil (Snout beetles) because it was first reported from Apopka, Florida. This is also recognized as a Diaprepes root weevil and considered as a very damaging pests of Citrus, many agricultural crops and ornamental plants throughout the United States.

Control of Black Vine Weevils with Insect Parasitic Nematodes / September 18, 2009 by Ganpati Jagdale

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 leaf beetles, Altica quercetorum and Agelastica alni are serious pests of urban trees including Quercus sp and Alnus sp, respectively. The elm leaf beetle Xanthogaleruka luteola is a serious pest that causes defoliation of eml trees (Ulmus spp.) in North America. Adults of these beetles generally feed on leaves by chewing holes through the leaf tissue. Larvae skelotonize leaves by feeding on leaf tissues leaving veins and upper epidermis intact.

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.

List of insects susceptible to various species of entomopathogenic nematodes / April 10, 2009 by Ganpati Jagdale

Insect Species: Entomopathogenic nematode species

Ø Apopka weevil (Diaprepes abbreviatus): S. carpocapsae All strain

Ø Armyworm (Heliothis armigera): S. carpocapsae All strain

Ø Billbugs (Sphenophorus purvulus): H. bacteriophora & S. carpocapsae All strain

Ø Black vine weevil (Otiorhynchus salcatus): S. carpocapsae All & UK strains, S. feltiae, S. glaseri & H. megidis UK 211 strain

Ø Blue grass weevil (Listronotus maculicollis): H. bacteriophora & S. carpocapsae

Ø Carpenter worms (Cossus cossus): S. carpocapsae

Ø Carrot weevil (Listronotus oregonensis): S. feltiae

Ø Cat fleas (Ctenocephalides felis): S. carpocapsae

Ø Citrus root weevil (Pachnaeus litus): S. carpocapsae All strain

Ø Clover root weevil (Sitona hispidulus): S. feltiae & H. bacteriophora

Ø Codling moth (Cydia pomonella): S. carpocapsae

Ø Crane flies (Tipula spp.): S. carpocapsae & H. megidis

Ø Cutworms (Agrotis ipsilon, A. segetum): S. carpocapsae All strain

Ø Dog fleas (Ctenocephalides cannis): S. carpocapsae

Ø Face fly (Musca autumnalis): S. carpocapsae, H. bacteriophora & S. feltiae

Ø Fall web worms (Hyphantria cunea): S. carpocapsae

Ø Flea beetles (Phyllotreta spp.): S. carpocapsae

Ø Fungus gnats (Bradysis spp.): H. bacteriophora, H. indica, H. zealandica, S. anomali, S. carpocapsae, S. feltiae SN strain & S. riobrave

Ø House flies (Musca domestica): S. carpocapsae, H. bacteriophora & S. feltiae

Ø Hunting billbug (Sphenophorus venatus venatus): S. carpocapsae All strain

Ø Japanese beetle (Popillia japonica): H. bacteriophora, H. indica, H. marelata, H. megidis, H. zealandica, S. anomali, S. carpocapsae, S. feltiae, S. glaseri, S. kushidai, S. riobrave, S. scapterisci & S. scarabae

Ø Leaf minors (Liriomyza trifolii): S. carpocapsae & S. feltiae

Ø Leopard moth (Zeuzera pyrina): S. carpocapsae

Ø Mole crickets (Gryllotapla gryllotapla): S. riobravis & S. scapterisci

Ø Peach borer moth (Synanthedon exitiosa): S. carpocapsae

Ø Pecan weevil (Curculio caryae): H. bacteriophora

Ø Pine weevil (Hylobius abietis): S. carpocapsae, S. feltiae & H. downesi

Ø Plum weevil (Conotrachelus nenuphar): S. riobrave 355 strain

Ø Shore flies (Scatella stagnalis): H. megidis, S. carpocapsae, S. feltiae & S. anomaly

Ø Sod webworm (Herpetogramma phaeopteralis): S. carpocapsae All strain

Ø Stable fly (Stomoxys calcitrans): S. carpocapsae, H. bacteriophora & S. feltiae

Ø Strawberry root borer (Nemocestes incomptus): S. carpocapsae

Ø Sugarcane borer (Diaprepes abbreviatus): S. carpocapsae All strain

Ø Sweet potato weevil (Cylasformicarius elegantulus): S. carpocapsae All strain & H. bacteriophora HP88 strain

Ø Western flower thrips (Frankliniella occidentalis): H. bacteriophora, H. indica, H. marelata, S. abassi, S. arenarium, S. bicornutum, S. carpocapsae, S. feltiae

Ø White grubs (Amphimallon solstitiale): S. glaseri

Ø White grubs (Anomala orientalis): H. bacteriophora, H. megidis, H. zealandica, S. carpocapsae, S. glaseri, S. longicaudum, S. scarabae

Ø White grubs (Ataenius spretulus): H. bacteriophora, S. glaseri & S. scarabae

Ø White grubs (Costelytra zealandica): H. bacteriophora & S. glaseri

Ø White grubs (Cotinus nitida): H. bacteriophora, S. carpocapsae, S. feltiae, S. glaseri & S. scarabae

Ø White grubs (Cyclocephala borealis): H. bacteriophora, H. indica, H. marelata, H. megidis, H. zealandica, S. glaseri & S. scarabae

Ø White grubs (Cyclocephala hirta): H. bacteriophora, H. megidis, S. carpocapsae, S. feltiae, S. glaseri, S. kushidai, S. riobrave & S. scarabae

Ø White grubs (Cyclocephala lurida): H. bacteriophora, S. glaseri & S. scarabae

Ø White grubs (Cyclocephala pasadenae): H. bacteriophora, S. glaseri, S. kushidai & S. scarabae

Ø White grubs (Hoplia philanthus): H. megidis, S. feltiae & S. glaseri

Ø White grubs (Maladera castanea): H. bacteriophora, S. glaseri & S. scarabae

Ø White grubs (Melolontha melolontha): H. bacteriophora, H. marelata, H. megidis, S. arenaria, S. feltiae, S. glaseri & S. riobrave

Ø White grubs (Phyllophaga congrua): H. bacteriophora, S. glaseri & S. scarabae

Ø White grubs (Phyllophaga crinita): H. bacteriophora, S. glaseri & S. scarabae

Ø White grubs (Phyllophaga georgiana): H. bacteriophora, S. glaseri & S. scarabae

Ø White grubs (Rhizotrogus majalis): H. bacteriophora, H. megidis, H. zealandica, S. carpocapsae, S. feltiae, S. glaseri & S. scarabae

For more information on insect pathogenic nematodes read following books:

Ø Nematodes As Biocontrol Agents by Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon.

Ø Entomopathogenic Nematodes in Biological Control by Gaugler, R. and Kaya, H. K. (eds.), CRC Press, Boca Raton

Ø Entomopathogenic Nematology by Gaugler, R. (Ed.), CABI

White grub species susceptible to entomopathogenic nematodes / January 15, 2009 by Ganpati Jagdale

Species of white grubs : Species of entompathogenic nematodes

Asiatic garden beetle (Maladera castanea): H. bacteriophora, S. glaseri, S. scarabae
Black turfgrass ataenius (Ataenius spretulus): H. bacteriophora, S. glaseri, S. scarabae
Cockchafer (Melolontha melolontha): H. bacteriophora, H. marelata, H. megidis, S arenaria, S. feltiae, S. glaseri, S. riobrave
Cranberry root grub (Phyllophaga Georgiana): H. bacteriophora, S. glaseri, S. scarabae
European chafer (Rhizotrogus majalis): H. bacteriophora, H. megidis, H. zealandica, S. carpocapsae, S. feltiae, S. glaseri, S. scarabae
Grass grub beetle (Costelytra zealandica): H. bacteriophora, S. glaseri
Green June beetle (Cotinus nitida): H. bacteriophora, S. carpocapsae, S. feltiae, S. glaseri, S. scarabae
Japanese beetle (Popillia japonica): H. bacteriophora, H. indica, H. marelata, H. megidis, H. zealandica, S. anomali, S. carpocapsae, S. feltiae, S. glaseri, S. kushidai, S. riobrave, S. scapterisci, S. scarabae
Masked Chafer (Cyclocephala pasadenae): H. bacteriophora, S. glaseri, S. kushidai, S. scarabae
Northern Masked Chafer (Cyclocephala borealis): H. bacteriophora, H. indica, H. marelata, H. megidis, H. zealandica, S. glaseri, S. scarabae
Oriental beetle (Anomala orientalis): H. bacteriophora, H. megidis, H. zealandica, S. carpocapsae, S. glaseri, S. longicaudum, S. scarabae
Southern Masked Chafer (Cyclocephala lurida): H. bacteriophora, S. glaseri, S. scarabae
Southwestern Masked Chafer (Cyclocephala hirta): H. bacteriophora, H. megidis, S. carpocapsae, S. feltiae, S. glaseri, S. kushidai, S. riobrave, S. scarabae
Summer chafer (Amphimallon solstitiale): S. glaseri
White grub (Hoplia philanthus): H. megidis, S. feltiae, S. glaseri
White grub (Phyllophaga crinita): H. bacteriophora, S. glaseri, S. scarabae
White grub (Phyllophaga congrua): H. bacteriophora, S. glaseri, S. scarabae

For more information on insect pathogenic nematodes read book "Nematodes As Biocontrol Agents" by Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon.

Kill leaf beetles (Altica quercetorum, Agelastica alni and Xanthogaleruka luteola) with Entomopathogenic Nematodes / December 21, 2008 by Ganpati Jagdale

The leaf beetles, Altica quercetorum and Agelastica alni are serious pests of urban trees including Quercus sp and Alnus sp, respectively.
The elm leaf beetle Xanthogaleruka luteola is a serious pest that causes defoliation of eml trees (Ulmus spp.) in North America.
Adults of these beetles generally feed on leaves by chewing holes through the leaf tissue.
Larvae skelotonize leaves by feeding on leaf tissues leaving veins and upper epidermis intact.
Entomopathogenice nematodes including Heterorhabditis megidis, Steinernema carpocapsae and S. feltiae can be used as potential biocontrol agents against different species leaf beetles (read Grewal et al., 2005 for more information).
It has been shown that both the pre-pupal and pupal stages of A. quercetorum and A. alni are very susceptible to H. megidis when applied in the soil.
The last instar larvae of X. luteola are highle susceptible to S. carpocapsae when applied to the mulch.

How Entomopathogenic Nematodes kill leaf beetles

When the infective juveniles are applied to the soil surface or mulch, they start searching for their hosts, in this case leaf beetles grubs.
Once a beetle grub has been located, the nematode infective juveniles penetrate into the grub body cavity via natural openings such as mouth, anus and spiracles.
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in grub blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills grubs 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 soil.

References: Refer following book to read more about efficacy of entomopathogenic nematodes against leaf beetles

1. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). Nematodes As Biocontrol Agents. CAB publishing, CAB International, Oxon

Kill Japanese beetles (Popillia japonica) with Entomopathogenic Nematodes / December 20, 2008 by Ganpati Jagdale

The Japanese beetle, Popillia japonica, is a most economically important pest of many ornamental plants and turf grasses.
Larvae of these beetles are called white grubs that generally feed on roots of over 300 plants but their primary food source is grass roots. Severe damage caused by these grubs can result in dead patches of turf that can be picked up like a loose carpet.
Adults mostly feed on leaves and flowers by chewing the tissue between the veins, a type of feeding called skeletonizing.
Chemical insecticides including Imidacloprid (Merit), Chlorpyrifos, Isofenphos, and Diazinon are generally used to manage white grubs but due to human health and environment pollution concerns their use is restricted.
Currently, environmentally safe biological control agents including a milky disease causing bacterium Bacillus popilliae (Milky spores) and entomopathogenic nematodes have been used to control this pest.
Three entomopathogenic nematodes including Heterorhabditis bacteriophora GPS11 and TF strains, H. zealandica X1 strain and Steinernema scarabaei have been considered to be the most effective species against Japanese beetle grubs.
It has been demonstrated that the application of H. bacteriophora GPS11 and TF strains, H. zealandica X1 strain and S. scarabaei at rate of 2.5 billion infective juveniles per hectare can cause about 96, 98 and 100%, respectively control of Japanese beetle grubs infesting turfgrass (for more information read Grewal et a., 2005).
Nematodes can be applied using traditional sprayers that are used for the application of insecticides.
Nematodes perform better when they are applied to target small stages of grubs.
Nematodes also survive better and remain efficacious when field/lawns are irrigated before and after nematode applications.

How Entomopathogenic Nematodes kill Japanese beetles

When the infective juveniles are applied to the soil surface or thatch layer, they start searching for their hosts, in this case Japanese beetle grubs.
Once a Japanese beetle grub has been located, the nematode infective juveniles penetrate into the Japanese beetle grub body cavity via natural openings such as mouth, anus and spiracles.
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in grub blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills Japanese beetle grubs 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 soil.

References

Grewal, P.S., Koppenhofer, A.M., and Choo, H.Y., 2005. Lawn, turfgrass and Pasture applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166.

Kill black cutworms (Agrotis ipsilon Hufnagel) with Entomopathogenic Nematodes / November 29, 2008 by Ganpati Jagdale

The black cutworm, Agrotis ipsilon (Hufnagel), is a polyphagous pest, feeding on almost all vegetables, many grain crops, ornamentals, turf grasses and weeds.
The plants damaged by black cutworms include beans, broccoli, cabbage, carrot, Chinese broccoli, Chinese cabbage, Chinese spinach, clover, corn, cotton, eggplant, flowering white cabbage, green beans, head cabbage, lettuce, mustard, potato, spinach, sugarcane, sweet potato, tomato, turnip, alfalfa, rice, sorghum, strawberry, sugarbeet, tobacco, bluegrass (Poa pratensis), curled dock (Rumex crispus); lambsquarters (Chenopodium album), yellow rocket (Barbarea vulgaris) and redroot pigweed (Amaranthus retroflexus).
There are five to nine larval instars that generally feed on seedlings at ground level by cutting off the stem causing a significant damage especially in newly planted fields. They also feed on roots and the below ground stem.
They can damage turfgrass by clipping off their blades and shoots.
The biological control agents including a bacterium Bacillus thuringiensis var. kurstaki, and entomopathogenic nematodes have a great potential against black cutworms.
Bacillus thuringiensis var. kurstaki produces a toxin that paralyzes the gut of the caterpillar. This toxin does not kill the caterpillars quickly, but it does cause the caterpillars to stop feeding, which in turn reducing the intensity of the damage.
Since caterpillars of cutworms are highly mobile insects, the entomopathogenic nematodes with ambush type of foraging strategy can be used very effectively for the management of cutworms.
For example, Steinernema carpocapsae, is an ambusher nematode species that can control black cutworms very effectively if applied at a rate of 1 billion nematodes/acre on golf course greens.

How Entomopathogenic Nematodes Kill Black Cutworms

When the infective juveniles are applied to the soil surface or thatch layer, they start searching for their hosts, in this case caterpillares.
Once a caterpillar has been located, the nematode infective juveniles penetrate into the caterpillar body cavity via natural openings such as mouth, anus and spiracles.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae) from their gut in the caterpillar 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.

Kill leafminers (Liriomyza spp.) with Entomopathogenic Nematodes / November 23, 2008 by Ganpati Jagdale

Leafminers (Liriomyza spp.) are considered as economically important polyphagous pests of many indoor vegetable crops and flowering plants.
Vegetable host crops included beans, beet, carrots, celery, cucumbers, eggplants, lettuce, melons, onions, peas, peppers, potatoes, squash and tomatoes.
Flowering host plants included ageratum, aster, calendula, chrysanthemum, dahlia, gerbera, gypsophila, marigold, petunia, snapdragon, and zinnia.
Leafminer maggots generally feed on leaf parenchyma tissues by tunneling/mining between the upper and lower epidermal leaf surfaces.
Adults generally feed on sap exuding from the punctures caused by maggots during mining.
Infested leaves appear stippled due to the punctures made by leafminers while feeding, mining and oviposition especially at the leaf tip and along the leaf margins.
Widespread mining and stippling on the leaves generally decreases the level of photosynthesis in the plant leading towards the premature leaf drop reducing the amount of shade, which in turn causes sun scalding of fruits.
Injuries caused by maggots on the foliage also allow entry of bacterial and fungal disease causing pathogens.
Life cycle of leafminers contains four stages including egg, maggot, pupa and adult.
Life cycle can be completed within 15-21 days depending upon the host and temperature.
Adult females lay eggs in leaf tissues, eggs hatch within 2-3 days into maggots, hatched maggots starts feeding immediately and become mature within 3-4 days. Mature larvae eventually cut through the leaf epidermis and move to the soil for pupation and adults emerge within 3 weeks of pupation in the summer.
Although, chemical insecticides are generally used to protect foliage from injury caused by leafminers, but development of insecticide resistance among leafminer populations is a major problem.
Insecticides also are highly disruptive to naturally occurring biological control agents, particularly parasitoids.
Therefore, biological control agents including Bacillus thuringiensis var. thuringiensis (Bt), parasitic wasps (Diglyphus begina, D. intermedius, D. pulchripes and Chrysocharis parksi) and entomopathogenic nematodes (Heterorhabditis spp, Steinernema carpocapase and S. feltiae) have been considered as alternatives to chemical pesticides.
For successful control of leafminers, entomopathogenic nematodes can be easily applied in water suspension as spray application on plant foliage.
Entomopathogenice nematodes including S. carpocapase and S. feltiae when applied at the rate of 5.3 X 108 nematodes/ha can cause over 64% mortality of leafminers but need at least 92% relative humidity.

How Entomopathogenic Nematodes kill leafminers

When the infective juveniles are applied as spray to plant foliage, they enter the leaf mines through the leaf miner feeding punctures or exit holes made by the adults.
Once inside the mine the nematodes swim to find a leafminer maggot, nematodes then penetrate into the maggot body cavity via natural openings such as mouth, anus and spiracles.
Infective juveniles of Heterorhabditis 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 maggot blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills maggots usually within 48 h after infection.

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

Hara, A.H., Kaya, H.K., Gaugler, R., Lebeck, L.M. and Mello, C.L. 1993. Entomopathogenic nematodes for biological control of the leafminer, Liriomyza trifolii (Dipt.: Agromyzidae). Entomophaga 38, 359-369.
Head, J. and Walters, K.F.A. 2003. Augmentation biological control utilising the entomopathogenic nematode, Steinernema feltiae, against the South American Leafminer, Liriomyza huidobrensis. Proceedings of the 1st International Symposium on Biological Control, (Hawaii, USA, 13-18 January 2002). USDA Forest Service, FHTET-03-05, 136-140.
Olthof, T.H.A. and Broadbent, A.B. 1992. Evaluation of steinernematid nematodes for control of a leafminer, Liriomyza trifolii, in greenhouse chrysanthemums. Journal of Nematology 24, 612.
Tong-Xian Liu, Le Kang, K.M.Heinz, J.Trumble. 2008. Biological control of Liriomyza leafminers: progress and perspective. CAB Reviews: Perspectives in Agriculture, Veterinary Science, Nutrition and Natural Resources, 2009, 4, No. 004, 16 pp.
Williams, E.C. and Walters, K.F.A. 1994. Nematode control of leafminers: Efficacy, temperature and timing. Brighton Crop Protection Conference - Pests and Disease. 1079-1084.
Williams, E.C. and MacDonald, O.C., 1995. Critical factors required by the nematode Steinernema feltiae for the control of the leafminers Liriomyza huidobrensis, Liriomyza bryoniae and Chromatomyia syngenesiae. Annals of Applied Biology. 127, 329-341.
Williams, E.C. and Walters, K.F.A. 2000. Foliar application of the entomopathogenic nematode Steinernema feltiae against leafminers on vegetables. Biocontrol Science and Technology 10, 61-70.

Kill Shore flies (Scatella stagnalis) with Entomopathogenic Nematodes / November 22, 2008 by Ganpati Jagdale

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.