Biological control of the red palm weevil, Rhynchophorus ferrugineus with entomopathogenic nematodes

The red palm weevil, Rhynchophorus ferrugineus is considered as a major pest of palms in the Mediterranean Basin. Because of cryptic habitats of these weevils, their management with chemical insecticides is difficult.  It has been demonstrated that the entomopathogenic nematodes have a potential to use as biological control agents against red palm weevils.  For example, Steinernema carpocapsae can cause over 80% mortality of weevils under field conditions when applied in a chitosan formulation (Dembilio et al., 2010, Llacer et al., 2009).

Read following literature for more information

Abbas, M.S.T., Saleh, M.M.E. and Akil, A.M. 2001.  Laboratory and field evaluation of the pathogenicity of entomopathogenic nematodes to the red palm weevil, Rhynchophorus ferrugineus (Oliv.) (Col.: Curculionidae). Anzeiger Fur Schadlingskunde-Journal of Pest Science. 74: 167-168.

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 canariensis. Pest Management Science. 66: 365-370.

Llacer, E., de Altube, M.M.M. and Jacas, J.A. 2009.  Evaluation of the efficacy of Steinernema carpocapsae in a chitosan formulation against the red palm weevil, Rhynchophorus ferrugineus, in Phoenix canariensis. Biocontrol. 54: 559-565.

Monzer, A.E, and El-Rahman, R.A. 2003.  Effect on Heterorhabditis indica of substances occurring in decomposing palm tissues infested by Rhynchophorus ferrugineus. Nematology. 5: 647-652.

Salama, H.S., Abd-Elgawad, M. 2001.  Isolation of heterorhabditid nematodes from palm tree planted areas and their implications in the red palm weevil control. Anzeiger Fur Schadlingskunde-Journal of Pest Science. 74: 43-45.

Salama, H.S. and Abd-Elgawad, M. 2002.  Activity of heterorhabditid nematodes at high temperature and in combination with cytoplasmic polyhedrosis virus. Anzeiger Fur Schadlingskunde-Journal of Pest Science. 75: 78-80.

A first report of occurrence of entomopathogenic nematodes in Nepal

Recently a survey was conducted to study the occurrence and distribution of entomopathogenic nematodes in Nepal.  Although a total of 276 soil samples were collected from various habitats, entomopathogenic nematode were found only in 29 samples.  Nematodes were isolates using the Galleria-baiting technique (Bedding and Akhurst,1975). Both heterorhabditid and steinernematid nematodes were identified at their species level using both molecular and morphological techniques.  In this survey, the occurrence of only one species of heterorhabditids including Heterorhabditis indica and four described species of steinernematids such as Steinernema abbasi, S. cholashanense, S. feltiae and S. siamkayai were reported for the first time in Nepal (Khatri-Chhetri et al., 2010). 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.

Khatri-Chhetri, H.B., Waeyenberge, L., Manandhar, H.K. and Moens, M. 2010.  Natural occurrence and distribution of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) in Nepal. Journal of Invertebrate Pathology. 103: 74-78.

Kill cereal leaf beetles, Oulema melanopus with entomopathogenic nematodes

Recently, it has been demonstrated that the entomopathogenic nematodes including Steinernema feltiae strain B30, S. carpocapsae strain C101, and Heterorhabditis bacteriophora strain D54 have a potential to use as biological control agents against cereal leaf beetles (Oulema melanopus), which is a most common pest of many cereal crops including barley, corn, oats, wheat, rye, millet and rice.

For more information on interaction between entomopathogenic nematodes and cereal leaf beetles read following research paper.

Laznik, Z., Toth, I., Lakatos, T., Vidrih, M. and Trdan, S. 2010.  Oulema melanopus (L.) (Coleoptera: Chrysomelidae) adults are susceptible to entomopathogenic nematodes (Rhabditida) attack: results from a laboratory study. Journal of Plant Diseases and Protection. 117: 30-32.

Entomopathogenic nematodes can be applied through infected insect host cadavers

Entomopathogenic nematodes are generally applied as infective juveniles in aqueous suspensions using various techniques including irrigation systems, sprayers and water cans. These nematodes can also be applied through infected host cadavers. It has been demonstrated that the application of nematode infected insect cadavers can provide superior nematode dispersal (Shapiro and Glazer, 1996), infectivity (Shapiro and Lewis, 1999) and survival (Perez et al., 2003) when compared with the nematodes that applied in aqueous suspensions. Please read following literature to learn more about the advantages and disadvantages of applying nematodes through infected insect cadavers.

Creighton, C.S. and Fassuliotis, G. 1985.  Heterorhabditis sp. (Nematoda: Heterorhabditidae): a nematode parasite isolated from the banded cucumber beetle Diabrotica balteata. Journal of Nematology. 17: 150–153.

Del Valle, E.E., Dolinksi, C., Barreto, E.L.S. and Souza, R.M. 2009.  Effect of cadaver coatings on emergence and infectivity of the entomopathogenic nematode Heterorhabditis baujardi LPP7 (Rhabditida: Heterorhabditidae) and the removal of cadavers by ants. Biological Control 50: 21–24.

Del Valle, E.E., Dolinksi, C., Barreto, E.L.S., Souza, R.M. and Samuels, R.I. 2008.  Efficacy of Heterorhabditis baujardi LP77 (Nematoda: Rhabditida) applied in Galleria mellonella (Lepidoptera: Pyralidae) insect cadavers to Conotrachelus psidii (Coleoptera: Curculionidae) larvae. Biocontrol Science and Technology. 18: 33–41.

Perez, E.E., Lewis, E.E and Shapiro-Ilan, D.I. 2003.  Impact of host cadaver on survival and infectivity of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) under desiccating conditions. Journal of Invertebrate Pathology. 82: 111–118.

Shapiro, D.I and Lewis, E.E. 1999.  Comparison of entomopathogenic nematode infectivity from infected hosts versus aqueous suspension. Environmental Entomology. 28: 907–911.

Shapiro, D.I. and Glazer, I. 1996.  Comparison of entomopathogenic nematode dispersal from infected hosts versus aqueous suspension. Environmental Entomology. 25: 1455–1461.

Shapiro-Ilan, D.I., Lewis, E.E., Behle, R.W and McGuire, M.R. 2001.  Formulation of entomopathogenic nematode-infected-cadavers. Journal of Invertebrate Pathology 78: 17–23.

Shapiro-Ilan, D.I., Lewis, E.E., Tedders, W.L. and Son, Y. 2003.  Superior efficacy observed in entomopathogenic nematodes applied in infected-host cadavers compared with application in aqueous suspension, Journal of Invertebrate Pathology 83: 270–272.

Shapiro-Ilan, D.I., Tedders, W.L. and Lewis, E.E., 2008. Application of entomopathogenic nematode-infected cadavers from hard-bodied arthropods for insect suppression. US Patent 7374,773.

Biological control of the cattle tick Rhipicephalus microplus with entomopathogenic nematodes

Recently, it has been demonstrated that the entomopathogenic nematode, Heterorhabditis amazonensis strain RSC-5 have a potential to use as a biological control agent against cattle tick,  Rhipicephalus (Boophilus) microplus (Monteiro et al., 2010), which is considered to be the most important tick parasite of livestock in the world.  This hardy tick can be found on many hosts including cattle, buffalo, horses, donkeys, goats, sheep, deer, pigs, dogs and some wild animals. This tick can also transmit babesiosis (cattle fever), which is caused by the protozoal parasites,  Babesia bigemina and Babesia bovis.  Also, transmit  anaplasmosis caused by Anaplasma marginale. Read following literature for more information on interaction between entomopathogenic nematodes and animal parasitic ticks

Freitas-Ribeiro G.M., Furlong, J., Vasconcelos, V.O., Dolinski, C. and Loures-Ribeiro, A. 2005.  Analysis of biological parameters of Boophilus microplus Canestrini, 1887 exposed to entomopathogenic nematodes Steinernema carpocapsae Santa Rosa and all strains (Steinernema : Rhabditida). Brazilian Archives of Biology and Technology. 48: 911-919.

Kocan, K.M., Pidherney, M.S., Blouin, E.F., Claypool, P.L., Samish, M. and Glazer, I. 1998.  Interaction of entomopathogenic nematodes (Steinernematidae) with selected species of ixodid ticks (Acari : Ixodidae). Journal of Medical Entomology. 35: 514-520.

Monteiro, C.M.D., Prata, M.C.D., Furlong, J., Faza, A.P., Mendes, A.S., Andalo, V. and Moino, A.2010.  Heterorhabditis amazonensis (Rhabditidae: Heterorhabditidae), strain RSC-5, for biological control of the cattle tick Rhipicephalus (Boophilus) microplus (Acari: Ixodidae). Parasitology Research. 106: 821-826.

Reis-Menini, C.M.R., Prata, M.C.A., Furlong, J. and Silva, E.R. 2008.  Compatibility between the entomopathogenic nematode Steinernema glaseri (Rhabditida : Steinernematidae) and an acaricide in the control of Rhipicephalus (Boophilus) microplus (Acari : Ixodidae). Parasitology Research. 103: 1391-1396.

How do entomopathogenic nematodes kill their insect hosts?

When the infective juveniles of entomopathogenic nematodes are applied to the soil surface in the fields or thatch layer on glf courses, they start searching for their insect hosts. Once insect larva has been located, the nematode infective juveniles penetrate into the larval body cavity via natural openings such as mouth, anus and spiracles. Infective juveniles of Heterorhabditis nematodes can also enter through the intersegmental membranes of the grub cuticle. Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in insect blood. In the blood, multiplying nematode-bacterium complex causes septicemia and kill their insect host usually within 48 h after infection. Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the host cadaver to seek new larvae in the soil.

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.