Prevent corn damage from The western corn rootworms (Diabrotica virgifera virgifera) using beneficial entomopathogenic nematodes.Read More
Stored grain/ product pests: Nematode Information Several stored grain/product insect pests like Indian meal moth (Plodia interpunctella), Mediterranean flour moth (Ephestia kuehniella), Sawtoothed grain beetle (Oryzaephilus surinamensis), Mealworms (Tenebrio molitor), Red flour beetle (Tribolium castaneum) and Warehouse beetle (Trogoderma variabile) attack and destroy large quantities of stored grains and products during long-term storage in farm bins, grain processing facilities, warehouses, retail stores, and eventually also on the consumer shelves.Read More
- A presence of entomopathogenic nematode species including Steinernema khoisanae, Steinernema yirgalemense, Steinernema citrae, Heterorhabditis bacteriophora and Heterorhabditis zealandica have been reported in citrus orchards in the Western Cape, Eastern Cape and Mpumalanga provinces of South Africa (Malan et al., 2011).
- All the above nematode species have showed a very high virulence against false codling moth, Thaumatotibia leucotreta an economically important pest of citrus in South Africa. For example, S. yirgalemense can cause over 74% mortality of both larval and pupal mortality of false codling moth when applied at the rate of 50-200 infective juveniles/ larval or pupal stages of false codling moth.
- Two entomopathogenic nematode species including S. yirgalemense and S. citrae were reported for the first time from South Africa (Malan et al., 2011).
Read following papers on entomopathogenic nematodes from South Africa
de Waal, J.Y., Malan, A.P. and Addison, M.F. 2011. Evaluating mulches together with Heterorhabditis zealandica (Rhabditida: Heterorhabditidae) for the control of diapausing codling moth larvae, Cydia pomonella (L.) (Lepidoptera: Tortricidae). Biocontrol Science and Technology 21: 255-270.
de Waal, J.Y., Malan, A.P., Levings, J. and Addison, M.F. 2010. Key elements in the successful control of diapausing codling moth, Cydia pomonella (Lepidoptera: Tortricidae) in wooden fruit bins with a South African isolate of Heterorhabditis zealandica (Rhabditida: Heterorhabditidae). Biocontrol Science and Technology. 20: 489-502.
Hatting, J., Stock, S.P. and Hazir, S. 2009. Diversity and distribution of entomopathogenic nematodes (Steinernematidae, Heterorhabditidae) in South Africa. Journal of Invertebrate Pathology 102: 120-128.
Malan, A.P., Knoetze, R. and Moore, S.D. 2011. Isolation and identification of entomopathogenic nematodes from citrus orchards in South Africa and their biocontrol potential against false codling moth. Journal of Invertebrate Pathology 108: 115-125.
Malan, A.P., Nguyen, K. B. and Addison, M. F. 2006. Entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) from the southwestern parts of South Africa. African Plant Protection 12: 65-69.
Malan, A.P., Nguyen, K.B., de Waal, J.Y. and Tiedt, L. 2008. Heterorhabditis safricana n. sp (Rhabditida : Heterorhabditidae), a new entomopathogenic nematode from South Africa. Nematology 10: 381-396.
Entomopathogenic nematode Steinernema carpocapsae for the control of red palm weevil, Rhynchophorus ferrugineus- Nematode Information /
It has been demonstrated that the curative applications of the entomopathogenic nematode Steinernema carpocapsae in a chitosan formulation can reduce the population of red palm weevil, Rhynchophorus ferrugineus infesting Cretan Date Palm, Phoenix theophrasti (Dembilio et al., 2011). Read following papers for more information.
Dembilio, O., Karamaouna, F., Kontodimas, D. C., Nomikou, M. and Jacas, J. A. 2011. Short communication. Susceptibility of Phoenix theophrasti (Palmae: Coryphoideae) to Rhynchophorus ferrugineus (Coleoptera: Curculionidae) and its control using Steinernema carpocapsae in a chitosan formulation. Spanish Journal of Agricultural Research 9: 623-626.
Dembilio, O., Llacer, E., de Altube, M.D.M. and Jacas, J.A. 2010. Field efficacy of imidacloprid and Steinernema carpocapsae in a chitosan formulation against the red palm weevil Rhynchophorus ferrugineus (Coleoptera: Curculionidae) in Phoenix. Pest Management Science 66: 365-370.
Heterorhabditis indica and Steinernema carpocapsae for controlling alfalfa weevil Application of Heterorhabditis indica and S. carpocapase at the rate 1 billion nematodes per hectare can reduce 72 and 50% population of alfalfa weevil, Hypera postica grubs, respectively. Another entomopathogenic nematode, Steinemema thermophillum was also effective in killing H. postica grubs (Shah et al., 2011).
Read following paper for information on the effect of entomopathogenic nematodes on alfalfa weevil
Shah, N.K., Azmi, M.I. and Tyagi, P.K. 2011. Pathogenicity of Rhabditid nematodes (Nematoda: Heterorhabditidae and Steinernematidae) to the grubs of alfalfa weevil, Hypera postica (Coleoptera: Curculionidae). Range Management and Agroforestry 32: 64-67.
Use an entomopathogenic nematode, Heterorhabditis bacteriophora to control long-horned beetle, Dorcadion pseudopreissi infesting turf. /
The application of an entomopathogenic nematode Heterorhabditis bacteriophora at the rate of 0.5 million infective juveniles per square meter can significantly reduce the population of Dorcadion pseudopreissi infesting turf grass (Lolium perenne) in the field (Susurluk et al. (2011). Read following papers for more information.
Susurluk, I.A., Kumral, N.A., Bilgili, U. and Acikgoz, E. 2011. Control of a new turf pest, Dorcadion pseudopreissi (Coleoptera: Cerambycidae), with the entomopathogenic nematode Heterorhabditis bacteriophora. Journal of Pest Science 84: 321-326.
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, Dorcadion pseudopreissi (Coleoptera: Cerambycidae). Biocontrol Science and Technology 19: 585-594.
The lesser peachtree borer, Synanthedon pictipes is a serious pest of commercially grown peach (Prunus spp.), orchards. It has been demonstrated that this insect pest can be controlled using entomopathogenic nematodes including Steinernema carpocapsae, S. riobrave and Heterorhabditis spp. Please read following article for interaction between the lesser peachtree borer and entomopathogenic nematodes.
Cottrell, T. E., Shapiro-Ilan, D. I., Horton, D. L., and Mizell, R. F., III. 2011. Laboratory virulence and orchard efficacy of entomopathogenic nematodes against the lesser peach tree borer (Lepidoptera: Sesiidae). Journal of Economic entomology 104: 47-53.
It has been reported that entomopathogenic nematodes can be used as biological control agent to manage species of the American (Periplaneta americana) and the German (Blattella germanica) cockroaches. Read following paper for more information
Maketon, M., Hominchan, A. and Hotaka, D. 2010. Control of American cockroach (Periplaneta americana) and German cockroach (Blattella germanica) by entomopathogenic nematodes. Revista Colombiana de Entomologia 36: 249-253.
Filbertworm, Cydia latiferreana is considered as an economically important insect pest of hazelnuts, Corylus avellana in North America. Three entomopathogenic nematode species including Heterorhabditis marelatus Pt. Reyes strain, Steinernema carpocapsae All strain and Steinernema kraussei L137 strain have been tested as biological control agents against filbertworm under both laboratory and field condition (Chambers et al., 2010; Bruck and Walton, 2007). These studies showed that these nematodes can cause about 73–100% mortality of filbertworms (Bruck and Walton, 2007) and can be used to manage overwintering worms on the hazelnut orchard floor (Chambers et al., 2010). Read following literature for information on the interaction between entomopathogenic nematodes and filbertworm.
Bruck, D.J. and Walton, V.M. 2007. Susceptibility of the filbertworm (Cydia latiferreana, Lepidoptera:Tortricidae) and filbert weevil (Curculio occidentalis, Coleoptera: Curculionidae) to entomopathogenic nematodes. Journal of Invertebrate Pathology. 96: 93–96.
Chambers, U. Bruck, D.J., Olsen, J. and Walton, V.M. 2010. Control of overwintering filbertworm (Lepidoptera: Tortricidae) larvae with Steinernema carpocapsae. Journal of Economic Entomology. 103: 416-422.
Entomopathogenic nematodes including Steinernema riobrave and Heterorhabditis indica were evalusted against a small hive beetle Aethina tumida Murray (Coleoptera: Nitidulidae) in the field. According to Ellis et al. (2010) both nematode species caused over 76% mortality of hive beetles. Shapiro-Ilan et al. (2010) tested efficacy of H. indica and Steinernema carpocapsae against hive beetles and demonstrated that both nematode species when applied through infected host cadavers can cause up to 78% control in hive beetles. This suggests that entomopathogenic nematodes have a potential to use as biological control agents against hive beetles. Read following papers for detail information on effect of entomopathogenic nematodes on the small hive beetles.
Ellis, J.D., Spiewok, S., Delaplane, K.S., Buchholz, S., Neumann, P. and Tedders, W.L. 2010. Susceptibility of Aethina tumida (Coleoptera: Nitidulidae) larvae and pupae to entomopathogenic nematodes. Journal of Economic Entomology. 103: 1-9.
Shapiro-Ilan, D.I., Morales-Ramos, J.A., Rojas, M.G. and Tedders, W.L. 2010. Effects of a novel entomopathogenic nematode-infected host formulation on cadaver integrity, nematode yield, and suppression of Diaprepes abbreviatus and Aethina tumida. Journal of Invertebrate Pathology. 103: 103-108.
How to apply nematodes Insect-parasitic nematodes can be easily applied using conventional pesticide and fertilizer sprayers that have up to 300 PSI pressures. However, nematodes will be easily damaged, if they are agitated through excessive recirculation of spray mix or if the temperature in the tank increases beyond 86 degrees F. Nematodes can also be applied through different types of irrigation systems but pumps should have proper pressure to avoid damage to nematodes and screen sizes should be larger than 50 mesh so that nematodes will pass through them live. Watering cans are used to apply nematodes in small areas including vegetable and ornamental gardens.
How many nematodes should be applied
For the suscessful control most of the soil dweling insect pests, the optimal rate of 1 billion infective juvenile nematodes in 100 to 260 gallons of water per acre is generally recommended.
Optimal soil and environmental condtions to apply nematodes
All nematodes require proper soil moisture for their optimal movement and infectivity. The activity and infectivity of nematodes can be enhanced by maintaining optimum moisture levels in the soil before and after their application. In case of nematode application in turf, turf should be irrigated immediately after applicationwith at least 1/2 inch of water to rinse off nematodes from the folliage and move them into the soil and thatch. As nematodes are very sensitiv to heat and cold, their infectivity will be reduced if soil temperature is below 4 degrees C and above 35 degrees C. Soil temperatures between 20 to 30 degrees C are considered favourable for application of majority of nematode species and their virulence. Nematode survival and activity also influenced by soil type. Both survival and activity of nematodes is higher in sandy-loam soils than in heavy clay soils.
When to apply nematodes
Since nematodes are very sensitive to UV light, they will die within a minute or two when exposed to full sun. Therefore, nematodes should be applied early in the morning or late in the evening to avoid exposure to UV light.
Pulse (legume) grains are considered as the important sources of protein, fats, carbohydrates, sugar and vitamin. B. In developing countries pulses are a cheaper protein source than meat. Many insect pests including red flour beetle Tribolium castaneum (Herbst), India meal moth Plodia interpunctella, Mediterranean flour moth Ephestia kuehniella (Zeller), saw thoothed grain beetle Oryzaephilus surinomensis (L.), yellow mealworm Tenebrio molitor (L.) and the ware house beetle Trogoderma variable (Ballion) cause a serious damage to these crops in the field and grains in the storage. The efficacy of entomopathogenic nematodes against many stored grain/product pests have been studied by many researchers (Athanassiou et al., 2008; Moris, 1985; Romos-Rodriguez et al., 2006). In the laboratory, an entomopathogenic nematode, Steinernema feltiae when applied at the rate 900 infective juveniles per insect caused over 66% mortality of both adults and larvae of T. confusum. This nematode when applied at the same rate also caused over 52% mortality of E. kuehniella. (Athanassiou et al., 2008) Under laboratory conditions, another species of nematode, S. riobrave can cause about 70% mortality of T. castaneum (Ramos-Rodríguez et al., 2007). It has also been demonstrated that nematodes including S. carpocapsae, Heterorhabditis bacteriophora and H. megidis have a potential to control the adults of two stored grain pests including, Sitophilus granarius and O. surinamensis (Tradan, 2006). Mbata and Shapiro-IIan (2005) also showed that various heterorhabditis nematodes including H. bacteriophora (HP88, Lewiston, and Oswego strains); H. indica (Homl strain); H. marelatus (Point Reyes strain); H. megidis (UK211 strain); and H. zealandica (NZH3 strain) have potential to kill larvae and adults of P. interpunctella. For more information on biological control of stored grain pets with entomopathogenice nematodes; please read following research papers:
Athanassiou CG, Palyvos NE, Kakoull-Duarte T. 2008. Insecticidal effect of Steinernema feltiae (Filipjev) (Nematoda : Steinernematidae) against Tribolium confusum du Val (Coleoptera : Tenebrionidae) and Ephestia kuehniella (Zeller) (Lepidoptera: Pyralidae) in stored wheat Journal of Stored Products Research. 44: 52-57.
Mbata, G.N., and Shapiro-Ilan, D.I. 2005. Laboratory evaluation of virulence of heterorhabditid nematodes to Plodia interpunctella Hübner (Lepidoptera: Pyralidae). Environmental Entomology 34: 676 - 682.
Ramos-Rodríguez, O., Campbell, J. F., and Ramaswamy, S. 2006. Pathogenicity of three species of entomopathogenic nematodes to some major stored- product insect pest. Journal of Stored Product Research 42: 241 - 252.
Ramos-Rodríguez,O.,Campbell, J. F.,and Ramaswamy, S. 2007. Efficacy of the entomopathogenic nematodes Steinernema riborave against the stored-product pests Tribolium castaneum and Plodia interpunctella. Biological Control 40:15 -21.
Tradan, S., Vidric, M., and Valic, N. 2006. Activity of four entomopathogenic nematodes against young adult of Sitophilus granarious (Coleptera: Curculionidae ) and Oryzophilus surinamensis ( Coleoptera: Silvanidae ) under laboratory condition. Plant Disease and Protection. 113: 168 - 173.
- 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.
Buy Steinernema feltiae
Control oriental beetles, Anomala orientalis with an entomopathogenic nematode Steinernema scarabaei /
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.
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.
Before starting to write about this topic, I would like to make it clear that taxonomically all bugs are insects but all the insects are not bugs. As far as I know, both in the USA and Canada, almost all people except entomologists call each and every insect as a bug. Even extension entomologists when they are giving extension seminars to farmers/growers about insect pests of different crops, they often refer them as bad bugs for the understanding of growers. "True" bugs are mainly belong to two insect orders including Hemiptera and Homoptera. All natural enemies of insect pests are considered as good bugs because they can kill and feed on insect pests that cause tremendous yield losses to many economically important crops. Since many of these natural enemies are commercially produced and used in the integrated pest management program (IPM), they are called as biological control agents. These biological control agents can be parasitic or predatory insects. In addition to these predators and parasites (good bugs), there are some microorganisms such as bacteria, fungi, protozoa and viruses that can cause diseases and kill insect pests. These microorganisms are termed as insect pathogens and also considered as biological control agents. Nematodes belonging to two families, Steinernematidae and Heterorhabditidae are also considered as insect parasites or pathogens and used as biological agents in controlling many soil dwelling insect pests of many economically important crops (in this blog, please read several posts that are devoted to insect- parasitic nematodes). Furthermore, mites are closely related to spiders but not considered as insects. Some species of mites are predatory in nature but others are serious pests of many plant species.
Predators: Although, there are many kinds of vertebrate predators including birds, amphibians, reptiles, fish and mammals that feed on insects, in this blog I am going to focus on the predatory insects that are generally used in biological control programs. These insects are called predators because they feed and complete their entire life cycle by remaining outside of their prey host as opposed to parasites that complete at least part of their life cycle inside their hosts. Predators are generally larger than their prey, they kill and feed on both immature and adult stages of many different kinds of hosts.
Following are the examples of insect predators that can be used as biological control agents against many kinds of insect pests.
Aphid midge (Aphidoletes aphidimyza): This predatory midge fly often found in many vegetable crops (potatoes, cabbage and cauliflower), fruit orchards (apple, blueberries and peaches) and many ornamental plants throughout North America. The larval stages of this midge fly are mainly predators of aphids. This midge fly is commercially available and widely used as biocontrol agents in the greenhouses against over 60 species of aphids infesting both vegetable and ornamental plants.
Bigeyed bug (Geocoris spp.): There are four most common species of bigeyed bug (G. punctipes, G. pallens, G. bullatus and G. uliginosus) found in almost all cropping systems in North America. Bigeyed bugs generally feed on many small insects including aphids, mites and whiteflies, eggs and nymphs of many plant bugs. They can also feed on eggs and small larval stages of cotton ballworms, pink ballworms and tobacco budworms. Since this bug is very susceptible to broad spectrum pesticides, care should be taken to avoid killing of this important biocontrol agent. This predator is commercially available from insectories in the USA.
Brown lacewings (Hemerobius stigma): These lacewings found throughout North American forests and are mainly predators of aphids and many other soft-bodied small insects including balsam woolly adelgis (Adelges piceae), pine bark adelgid (Pineus strobi) and Cooley's spruce gall adelgid (Adelges cooleyi). These lacewings are not commercially available.
Deraeocoris bug (Deraeocoris nebulosus): This is a very important predator of many insect and mite pests different agricultural, horticultural and landscape plants in the Canada and USA. This is a true predatory bug, which is generally found in many fruit orchards including apple, peach and pecan. They also found in cotton fields and many landscape settings. These bugs are natural enemies of many small insects including aphids, lace bugs, psyllids, scales and whiteflies. They also feed on mites. These bugs are not commercially available.
Dragon and damselflies: Adult dragon and damsel flies generally feed on small flying small adult insects including midge flies, mayflies, mosquitoes, ants and termites in the air where as dragon/damsel fly nymphs feed on mosquito larvae in the water.
Green lacewing (Chrysoperla carnea, C. rufilabris): Lacewings adults are not predatory in nature but mainly feed on nectar, honeydew and pollens. However, larvae of lacewings are predatory in nature and feed on insect pests of many crops including apples, asparagus, cotton, corn, cole crops, eggplants, leafy vegetables, potatoes, tomatoes, peppers and strawberries. Lacewing larvae generally prey on aphids, leafhopper eggs, eggs of butterflies and moths, mealybugs, mites, thrips, small larvae of beetles and moths. Both species of lacewings are commercially available and sold in all stages (eggs, larvae and adults).
Ladybird beetles (Hippodamia parenthesis and Harmonia axyridis): These beetles are also recognized as lady beetles or ladybugs and more than 450 of this beetles have been reported from North America. Both larval and adult stages of this predator found on many agricultural and ornamental plants and they primarily feed on aphids. In addition, they can feed on small insects, mites, scales, thrips and eggs of many moths and beetles. they can eat nectar or pollen if insect hosts are not around. These predators are now commercially available to use against many crop pests, especially aphids.
Lebia beetles (Lebia grandis): These beetles are natural enemies of Colorado potato beetle, Leptinotarsa decemlineata. Adults of the predatory insect can feed on all immature stages of colorado potato beetle. Larval stages of Labia beetles are generally parasitic in nature and therefore, they are considered as ectoparasites of larval and pupal stages of colorado potato beetles. These predators are not commercially produced.
Pirate bugs (Orius spp.): Both adults and nymphs of these predatory insects have a sharp, needle-like beak that they use to suck body content of their prey. These insects found in many crops including alfalfa, corn, cotton, pea, peanuts, and strawberries. These are predators of aphids, mites, thrips, small larval stages of many insects, eggs of many different kinds of insects. These insect predators are commercially available in the USA and most often suscessfully used as biocontrol agents in controlling greenhouse pests.
Rove beetles (Aleochara bilineata): These beetles naturally found in many vegetable crops including onions, different cole crops, turnip, radish and sweet corn. Rove beetle adults are predatory in nature but their larval stages are parasitic in nature. Rove beetles generally feed on egg, larval and pupal stages of onion and cabbage maggots. These insects are not commercially available.
Soldier beetles (Chauliognathus marginatus and C. pennsylvanicus): These beetles are also called leatherwing beetle because of texture of their wings. Larvae of this insect mainly feed on grasshopper eggs, both adult and nymphal stages of aphids, soft bodied larvae of many insects (cutworms, gypsy moths) whereas adults mainly feed on adult aphids and other soft bodied insects. These predators also feed on snails and slugs. These insects are not pest any plant species but they can eat nector or pollen if insect hosts are not around.
Spined soldier bug (Podisus maculiventris): This is a "true bug" that also named as a stink bug because it emits a strong stinky odour when disturbed. Like Pirate bugs, this bug also uses its sharp beak to suck the body content of its prey. This predator feeds on immature stages of many insect pests including beet armyworm, cabbage loopers, cabbageworm, colorado potato beetle, corn earworm, diamond backmoth, Eropean corn borer, fall armyworms, flea beetles, Mexican bean beetle and velvetbean caterpillars. These insect predators are commercially available.
What is biological control of insect pests? Biological control is a method in which natural enemies are introduced in the fields or greenhouses to suppress the populations of economically important insect pests of many plant species. Natural enemies may include predators, parasities and pathogens.Read More
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.Read More
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
Biological control of slugs and snails with parasitic nematode, Phasmarhabditis hermaprodita
Slugs (Mollusca: Gastropoda) are considered as important pests of many agricultural and horticultural crops throughout the world.
Recently, a slug parasitic nematode, P. hermaprodita has been commercialized as a biological molluscicide by MicroBio Ltd, UK and sold under the trade name "Nemaslug".
Phasmarhabditis hermaprodita as been found to be associated with several different bacteria rather than one particular species but the association with a bacterium, Moraxella oslensis proved to be highly pathogenic to gray garden slug (Deroceras reticulatum) and preferred bacterium for mass production of this nematode in monoxenic culture.
Like entomopathogenic nematodes, slug parasitic nematode infective juveniles or dauer juveniles move through soil, locate slugs and infect. They penetrate slugs through a natural opening at the backside of the mantle. Once inside, the dauer juveniles release bacterial cells, start feeding on multiplying bacteria and develop into self-fertilizing hermaphrodites. Nematode- bacteria complex can cause the death of the slug within 7-21 days after infection.
Phasmarhabditis hermaprodita can attack and kill several species of slugs including Arion ater, A. intermedius, A. distinctus, A. silvaticus, D. reticulatum, D. caruanae, Tandonia budapestensis and T. sowerbyi.
Phasmarhabditis hermaprodita can also parasitize several species of snails including Cernuella virgata, Cochlicella acuta, Helis aspersa, Monacha cantiana, Lymnaea stagnalis and Theba pisana.
It has been demonstrated that slug parasitic nematodes when applied at the rate of 3x 109 infective juveniles/hectare can give better control of slugs than standard chemical molluscicide, Methiocarb pellets.
For more information on insect and slug parasitic nematodes read a book "Nematodes As Biocontrol Agents" by Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon.