Biological control of grape root borer Vitacea polistiformis using entomopathogenic nematodes.

Efficacy of two entomopathogenic nematodes including Heterorhabditis zealandica strain X1 and H. bacteriophora Strain GPS11 was studied in the field against grape root borer Vitacea polistiformis (Williams et al., 2010).  This borer can damage roots of both wild and cultivated Vitis and Muscadinia grapes and is considered as a major pest of grapes grown in the eastern United States.  According to Williams et al. (2010), both H. zealandica and H. bacteriophora can cause up to 92% control of grape root borer and they can also persist in the soil for a extended period after their application.

Read following literature for more information on interaction between entomopathogenic nematodes and the grape root borers.

Williams, R.N., Fickle, D.S., Grewal, P.S. and Dutcher, J. 2010.  Field efficacy against the grape root borer, Vitacea polistiformis (Lepidoptera: Sesiidae) and persistence of Heterorhabditis zealandica and H. bacteriophora (Nematoda: Heterorhabditidae) in vineyards. Biological Control. 53: 86-91.

Williams, D.S. Fickle, P.S. Grewal and J.R. Meyer. 2002.  Assessing the potential of entomopathogenic nematodes to control the grape root borer, Vitacea polistiformis (Lepidopetera: Sesiidae) through laboratory and greenhouse bioassays. Biocontrol Science and Technology 12: 35-42.

Can you kill small hive beetles (Aethina tumida) with entomopathogenic nematodes?

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.

A record of new entomopathogenic nematode species from Brazil

An entomopathogenic nematode in a soil sample collected from a natural forest in Mato Grosso do Sul state, Brazil was described using both morphological and molecular characteristics as a new species "Steinernema brazilense (Rhabditida: Steinernematidae)" (Nguyen et al., 2010). Reference:

Nguyen, K.B., Ginarte, C.M.A., Leite, L.G., dos Santos, J.M. and Harakava, R. 2010. Steinernema brazilense n. sp (Rhabditida: Steinernematidae), a new entomopathogenic nematode from Mato Grosso, Brazil. Journal of Invertebrate Pathology. 103: 8-20.

How and when to apply insect-parasitic nematodes

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.

Can we control plant-parasitic nematodes with entomopathogenic nematodes?

For the last several decades, entomopathogenic nematodes have been successfully used for the management of insect pests of many economically important crops (Grewal et al., 2005).  As an additional benefit, several researchers including Fallon et al. (2002), Gouge et al. (1997), Grewal et al. (1997; 1999), Jagdale et al. (2002), Jagdale and Grewal (2008), LaMondia and Cowles (2002), Lewis et al. (2001), Lewis and Grewal (2005), Molina et al. (2007), Nyczepir et al. (2004), Perez and Lewis (2002), Perry et al. (1998) and Shapiro et al. (2006) have demonstrated that entomopathogenic nematodes can also be used as biological control agents to control plant-parasitic nematodes infesting different crops in the fields and greenhouses . To control plant- parasitic nematodes, entomopathogenic nematodes can be applied using standard spraying equipments used for application of chemical pesticides. Entomopathogenic nematodes are generally applied against plant-parasitic nematodes at the rate of 1 billion infective juveniles per acre but this rate can vary with both entomopathogenic nematode and plant- parasitic nematode species.  Following are the examples of different species of entomopathogenic nematode that found to be successful in suppressing the population of different species of plant- parasitic nematodes.  Steinernema carpocapsae can reduce the population of ring nematodes (Mesocriconema spp., Criconemoides spp.) by 65%.  S. carpocapsae can reduce the population of stubby root nematodes (Paratrichodorus spp.) by 60%.  S. carpocapsae can reduce the population of potato cyst nematodes (Globodera rostochiensis).  S. carpocapsae can reduce the populations of foliar nematode Aphelenchoides fragariae.  Steinernema riobrave can reduce the population of stunt nematodes (Tylenchorynchu spp.) by 85%.  S. riobrave can reduce the population of lance nematodes (Hoplolaimus spp.).  S. riobrave can reduce the population of root-knot nematodes (Meloidogyne spp.) by 83%.  S. riobrave reduced egg masses of root-knot nematodes (Meloidogyne spp.).  S. riobrave can reduce the population of sting nematodes (Belonolaimus longocaudatus).  Steinernema feltiae can inhibit hatching root-knot nematode eggs and infection by hatched infective juveniles of root-knot nematodes (Meloidogyne spp.).  S. feltiae reduced egg masses of root-knot nematodes (Meloidogyne spp.) .  S. feltiae can reduce the population of root-knot nematodes (Meloidogyne spp.).  Steinernema glaseri reduced egg masses of root-knot nematodes (Meloidogyne spp.).  Heterorhabditis bacteriophora can reduce the population of ring nematodes (Mesocriconema spp., Criconemoides spp.) by 80%.  H. bacteriophora can reduce the population of stunt nematodes (Tylenchorynchus spp.) by 60%.  H. bacteriophora can reduce the population of lesion nematodes (Pratylenchus pratensis).   H. baujardi can inhibit hatching root-knot nematode eggs and infection by hatched infective juveniles of root-knot nematodes (Meloidogyne mayaguensis). Read following literature for more information on interaction between entomopathogenic nematodes and plant- parasitic nematodes:

1. Fallon, D.J., Kaya, H.K., Gaugler, R., Sipes, B.S., 2002. Effects of entomopathogenic nematodes on Meloidogyne javanica on tomatoes and soybeans. Journal of Nematology 34, 239-245.

2. Fallon, D.J., Kaya, H.K., Sipes, B.S., 2006. Enhancing Steinernema spp. suppression of Meloidogyne javanica. Journal of Nematology 38, 270-271.

3. Grewal, P.S., Ehlers, R.-U., Shapiro-Ilan, D.I. (Eds.), 2005. Nematodes As Biocontrol Agents. CABI Publishing, CAB International, Oxon, U.K.,

4. Grewal, P.S., Lewis, E.E., Venkatachari, S., 1999. Allelopathy: a possible mechanism of suppression of plant-parasitic nematodes by entomopathogenic nematodes. Nematology. 1, 735-743.

5. Grewal, P.S., Martin, W.R., Miller, R.W., Lewis E.E., 1997. Suppression of plant-parasitic nematode populations in turfgrass by application of entomopathogenic nematodes. Biocontrol Science and Technology 7, 393-399.

6. Jagdale, G.B., Grewal, P.S., 2008. Influence of the entomopathogenic nematode Steinernema carpocapsae in host cadavers or extracts from cadavers on the foliar nematode Aphelenchoides fragariae on Hosta. Biological Control 44, 13-23.

7. Jagdale, G.B., Somasekhar, N., Grewal, P.S., Klein, M.G., 2002. Suppression of plant parasitic nematodes by application of live and dead entomopathogenic nematodes on Boxwood (Buxus spp). Biological Control. 24, 42-49.

8. Lewis, E.E., Grewal, P.S., 2005. Interactions with plant-parasitic nematodes. In: Grewal, P.S., Ehlers, R.-U., Shapiro-Ilan, D.I. (Eds.), Nematodes As Biocontrol Agents. CABI Publishing, CAB International, Oxon, U.K., pp. 349-362.

9. Perry, R.N., Homonick, W.M., Beane, J., Briscose, B., 1998. Effects of the entomopathogenic nematodes, Steinernema feltiae and S. carpocapsae, on the potato cyst nematode, Globodera rostochiensis, in pot trials. Biocontrol Science and Technology 8:175 – 180.

10. Shapiro, D.I., Nyczepir, A.P., Lewis, E.E., 2006. Entomopathogenic nematodes and bacteria applications for control of the pecan root-knot nematode, Meloidogyne partityla in the greenhouse. Journal of Nematology 38, 449-454.

Can you control stored grain insect pests with entomopathogenic nematodes?

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:

Hello, World!

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.

Entomopathogenic Nematodes and fungus gnats

• 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 suppliersdistributed 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.

Manage insect pests of Strawberries with entomopathogenic nematodes

Strawberries are one of the most economically grown crops throughout the world and in North America with annual yields ranging from 4-20 tons per acre and average monitory values between $2,800 to $14000 per acre.  There are several kinds of insect pests have been reported that attack and cause significant economic losses (over 60%) to this crop.   Different species of entomopathogenic have been used as biological control agents against different  insect pests of strawberries. It has been demonstrated that  the entomopathogenic nematode, Steinernema kraussei can reduce over 81%  population of black vine weevil (Ansari et al., 2010; Susurluk and Ehlers, 2008; Willmott et al., 2002). Entomopathogenic nematodes, Heterorhabditis megidis and H. downesi also can reduce 93 and 51% population of black vine weevil, respectively (Boff et al., 2001, 2002; Lola-Luz et al., 2005; Fitters et al., 2001). Populations of black vine weevils were also reduced by application of infective juveniles of Steinernema carpocapsae and S. glaseri (Booth et la., 2002). Steinernema carpocapsae can reduce 51% population of strawberry crown moth (Bruck et al., 2008).

Please read following literature for more information on interaction between insect pests of strawberries and different species entomopathogenic nematodes.

Ansari, M.A., Shah, F.A. and Butt, T.M. 2010.  The entomopathogenic nematode Steinernema kraussei and Metarhizium anisopliae work synergistically in controlling overwintering larvae of the black vine weevil, Otiorhynchus sulcatus, in strawberry growbags. Biocontrol Science and Technology. 20: 99-105.

Berry, R.E., Liu, J. and Groth, E. 1997.  Efficacy and persistence of Heterorhabditis marelatus (Rhabditida: Heterorhabditidae) against root weevils (Coleoptera: Curculionidae) in strawberry. Environmental Entomology. 26: 465-470.

Boff, M.I.C., van Tol, R.H.W.M. and Smits, P.H. 2002.  Behavioural response of Heterorhabditis megidis towards plant roots and insect larvae. Biocontrol. 47: 67-83.

Boff, M.I.C., Wiegers, G.L. and Smits, P.H. 2001.  Influence of insect larvae and plant roots on the host-finding behaviour of Heterorhabditis megidis. Biocontrol Science and Technology. 11: 493-504.

Boff, M.I.C., Zoon, F.C. and Smits, P.H. 2001.  Orientation of Heterorhabditis megidis to insect hosts and plant roots in a Y-tube sand olfactometer. Entomologia Experimentalis et Applicata. 98: 329-337.

Booth, S.R., Tanigoshi, L.K., Shanks, C.H. 2002.  Evaluation of entomopathogenic nematodes to manage root weevil larvae in Washington state cranberry, strawberry, and red raspberry. Environmental Entomology. 31: 895-902.

Bruck, D.J., Edwards, D.L. and Donahue, K.M. 2008.  Susceptibility of the strawberry crown moth (Lepidoptera : Sesiidae) to entomopathogenic nematodes. Journal of Economic Entomology. 101: 251-255.

Curran, J. 1992. Influence of application method and pest population-size on the field efficacy of entomopathogenic nematodes. Journal of Nematology. 24: 631-636.

Fitters, P.F.L., Dunne, R. and Griffin, C.T. 2001.  Vine weevil control in Ireland with entomopathogenic nematodes: optimal time of application. Irish Journal of Agricultural and Food Research. 40: 199-213.

KakouliDuarte, T., Labuschagne, L. and Hague, N.G.M. 1997.  Biological control of the black vine weevil, Otiorhynchus sulcatus (Coleoptera: Curculionidae) with entomopathogenic nematodes (Nematoda: Rhabditida). Annals of Applied Biology. 131: 11-27.

Lola-Luz, T. and Downes, M. 2007.  Biological control of black vine weevil Otiorhynchus sulcatus in Ireland using Heterorhabditis megidis. Biological Control. 40: 314-319.

Lola-Luz, T., Downes, M. and Dunne, R. 2005.  Control of black vine weevil larvae Otiorhynchus sulcatus (Fabricius) (Coleoptera : Curculionidae) in grow bags outdoors with nematodes. Agricultural and Forest Entomology. 7: 121-126.

Simser, D. and Roberts, S. 1994.  Suppression of strawberry root weevil, Otiorhynchus-ovatus, in cranberries by entomopathogenic nematodes (Nematoda, Steinernematidae and Heterorhabditidae). Nematologica. 40: 456-462.

Susurluk, A. and Ehlers, R.U. 2008.  Sustainable control of black vine weevil larvae, Otiorhynchus sulcatus (Coleoptera: Curculionidae) with Heterorhabditis bacteriophora in strawberry. Biocontrol Science and Technology. 18: 635-640.

Vainio, A. and Hokkanen, H.M.T. 1993.  The potential of entomopathogenic fungi and nematodes against Otiorhynchus-ovatus L and O. dubius strom (Col, Curculionidae) in the field. Journal of Applied Entomology-Zeitschrift fur Angewandte Entomologie. 115: 379-387.

Willmott, D.M., Hart, A.J., Long, S.J., Edmondson, R.N. and Richardson, P.N. 2002.  Use of a cold-active entomopathogenic nematode Steinernema kraussei to control overwintering larvae of the black vine weevil Otiorhynchus sulcatus (Coleoptera: Curculionidae) in outdoor strawberry plants. Nematology. 4: 925-932.

Wilson, M., Nitzsche, P. and Shearer, P.W. 1999.  Entomopathogenic nematodes to control black vine weevil (Coleoptera : Curculionidae) on strawberry. Journal of Economic Entomology. 92: 651-657.

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.

Occurrence of entomopathogenic nematode Steinernema feltiae in Slovenia

Presence of an entomopathogenic nematode, Steinernema feltiae (Rhabditida: Steinernematidae) was recorded for first time in soil samples collected from grasslands and field crops in central part of Slovenia. Nematodes were isolated using Galleria-baiting technique (Bedding and Akhurst, 1975) and identified using molecular technique. Read following literature for more information

Bedding, R.A. and R.J. Akhurst. 1975. A simple technique for detection of insect parasitic rhabditid nematodes in soil. Nematologica. 21: 109-110.

Laznik, Z., Toth, T., Lakatos, T., Vidrih, M. and Trdan, S. 2009.  First record of Steinernema feltiae (Filipjev) (Rhabditida: Steinernematidae) in Slovenia. Helminthologia. 46: 135-138.

Parasitization of subterranean termite Heterotermes aureus by beneficial nematodes

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

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

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

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.

Bugs for growers: Predatory Mites

Bugs for growers Predatory mites: 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 also includes predatory mites, which are closely related to spiders but they are not considered as insects. Although some species of mites are predatory in nature, there are several species of mites considered as pests because they cause a serious damage to many plant species. In general, predatory mites feed on spider mites, which are serious pests of many vegetable crops and ornamental plants. Predator mites also prey on thrips, fungus gnat eggs and other small soft bodied insects. Predatory mites are commercially available and used as biological control agents against many species of spider mites and fungus gnats in the Canadian, European, USA greenhouses.

Following are the examples of predatory mites that can be used as biological control agents against many kinds of insect pests.

Euseius tularensis: This predatory mite often found in citrus orchards and generally feed on citrus thrips and red mites.  Color of E. tularensis depends on the type of insect host they feed on. They are red, when they are feeding on red spider mites, yellow when they are feeding on thrips. These predatory mites also feed on scale insects, nymphs of white flies and two-spotted mites. This predatory mites are not available commercially.

Western predatory mite, Galendromus occidentalis: This predatory mite generally found in apple, cherry, peach, pear, and plum orchards. These pear shaped predatory mites generally feed on apple rust mites, blister mites, European red mites, yellow spider mites, pear and prunus rust mites and two-spotted spider mites. This important biocontrol agent is available commercially.

Galendromus pyri: This predatory mite is also found in many fruit orchards. This predatory mite also feeds on Apple rust mite, European red mites and two-spotted spider mites. These predatory mites are available commercially.

Neoseiulus fallacis: This pear shaped predatory mite found in many fruit orchards in the USA and also feeds on European red mites and two-spotted spider mites. These predatory mites are available commercially.

Phytoseiulus persimilis: This predatory mite is commercially available and most often used as biological control agents against different species of spider mites infesting vegetable crops and ornamental plants grown in the Canadian, European, US greenhouses.

Zetzellia mali: This pear shaped predatory found in many fruit orchards in the USA and also feeds on European red mites and two-spotted spider mites. These predatory mites are not available commercially.

Use Good Bugs to Control Bad Bugs: Predatory insects

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.

VOTiVO: A new biological control Product from Bayer CropScience for plant-parasitic nematode control

Recently, Bayer CropScience, Research Triangle Park, North Carolina announced that the company will launch a new Plant Growth Promoting Rhizobacteria (PGPR) based biological seed treatment product called "VOTiVO" for the biological control of different species of plant-parasitic nematodes that are responsible for reducing yields of several economically important crops. For more information on this product please visit following links: www.plantmanagementnetwork.org/pub/php/news/2009/VOTiVO/ www.VOTiVO.us

A record of new entomopathogenic nematode species from China

An entomopathogenic nematode in the soil samples that collected from the northern part of the Yunnan province of China was described using both morphological and molecular characteristics as new species "Steinernema xueshannense"  (Mracek et al., 2009). Reference:

Mracek, Z., Liu, Q., Nguyen, K. B. 2009. Steinernema xueshanense n. sp (Rhabditida, Steinernematidae), a new species of entomopathogenic nematode from the province of Yunnan, southeast Tibetan Mts., China. Journal of Invertebrate Pathology. 102: 69-78.