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    Bugs for Growers — Beneficial nematodes

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    Twelve Important Facts about Beneficial Entomopathogenic Nematodes

    1. What are insect-parasitic/entomopathogenic nematodes?

    By definition nematodes are thread-like microscopic, colorless and unsegmented round worms found in almost all habitats especially soil and water (Fig. 1).   [caption id="attachment_338" align="aligncenter" width="176" caption="Fig. 1. Nematodes are microscopic, non-segmented, thread-like round worms. Click on image for enlargement"]"Nematode"[/caption]

    Insect-parasitic nematodes:

    Nematodes that infect and complete their development, and reproduction at their insect host's expense are called as insect-parasitic nematodes.  In the phylum Nematoda, some members of a family Mermithidae (Order: Mermithida) including mosquito-parasitic nematode, Romanomermis culicivorax and grasshopper nematode Mermis nigrescens are considered as insect-parasitic nematodes but not as entomomopathogenic nematodes whereas the members of the two families Steinernematidae and Heterorhabditidae (Order: Rhabditida) including Steinernema spp. and Heterorhabditis spp., respectively are considered as both insect-parasitic and entomomopathogenic nematodes.

    Entomopathogenic nematodes:

    Members of both Steinernematidae and Heterorhabditidae families are also called as entomopathogenic nematodes because their infective juveniles are mutualistically associated with a specific kind of symbiotic bacteria, which are pathogenic to a variety of their insect hosts (Table 2). Although entomopathogenic nematodes are naturally present in the soil and responsible for suppressing the natural populations of insect pests, currently the main interest in them is to apply them inundatively as beneficial biological control agents to manage various economically important insect pests of different agricultural and horticultural crops, and ornamental plants (Grewal et al., 2005). Within last 30-40 years, 26 and 75 different species of Heterorhabditid (Table 3) and Steinernematid (Table 4) nematodes, respectively have been isolated and described from various parts of the world. A few of these described nematode species have been commercially produced and used as effective biological control agents against many insect pests of several economically important crops. These nematodes can infect and kill larvae/ caterpillars, pupae and adults of a variety of insect pests (Table 2; Fig. 2).   [caption id="attachment_704" align="aligncenter" width="300" caption="Fig. 2. Diagram showing that the entomopathogenic nematodes can infect and kill various stages (larvae, pupae and adults) of their host insects."]"Entomopathogenic nematodes can infect larval, pupal and adult stages of their insect hosts"[/caption] Therefore, these nematodes are also recognized and sold as beneficial nematodes. Unlike toxic chemical nematicides/pesticides, these beneficial nematodes are safe to the environment, human health, both pet and wild animals, and plants.  Also, they are not harmful to beneficial insects such as honeybees. Therefore, in this blog, we are providing some basic information on the mutualistic association between nematodes and their symbiotic bacteria, life cycle, host finding ability, production and application of entomopathogenic nematodes. Also, in our routine blog articles, we would like to provide a description of different insect and mollusk pests and their susceptibility to different species of entomopathogenic nematodes.

    2. What kinds of symbiotic bacteria are associated with entomopathogenic nematodes?

    • Two different kinds of symbiotic bacteria in the genus, Photorhabdus (Table 3) and Xenorhabdus (Table 4) are symbiotically associated with the species specific infective juveniles of Heterorhabditis spp. (Family: Heterorhabditidae) and Steinernema spp. (Family: Steinernematidae), respectively.
    • Species of both Xenorhabdus and Photorhabdus are motile gram-negative bacteria belong to the family Enterobacteriaceae and also exist in two main phenotypic forms (phase I and II), a phenomenon known as phase variation (Han and Ehlers, 2001).
    • The phase I form (also termed as primary form) varies physiologically and morphologically from phase II form (also called as secondary form).
    • Also, a main property distinguishing Xenorhabdus spp. from Photorhabdus spp. is that the only Photorhabdus bacteria have an ability to emit the light under stationary-phase culture conditions and in the infected host insect cadavers.

    3. What is an infective juvenile?

    A third-stage juvenile of an entomopathogenic nematode is called as an infective juvenile because it initiates the infection in its host. Infective juvenile is the only non-feeding and free-living stage found in the soil but all other stages including fourth and fifth (adult) and egg stages are completed inside the host.

    4. What is a dauer juvenile?

    The infective juveniles are actually third-stage juvenile that also called as dauer juveniles because they are enclosed in a second-stage cuticle, which arrests their further development (Fig.3; adopted from http://www.nematodeinformation.com) and helps to survive outside the host i.e. in the soil environment. Furthermore, these developmentally arrested dauer juveniles are physiologically adapted to remain in the environment (i.e. soil) without feeding until a perspective host is located. These dauer juveniles recover and resume their development only when they enter the perspective insect host’s body cavity via natural openings and shed their second stage cuticle. The dauer juveniles are also well known to tolerate harsh environmental conditions including extreme hot and cold temperatures, and desiccation (Jagdale and Gordon, 1997; Jagdale and Grewal, 2003; 2007; Jagdale et a., 2005). [caption id="attachment_470" align="aligncenter" width="300" caption="Fig. 3. A dauer juvenile of an entomopathogenic Steinernema carpocapsae nematode. adapted from www.nematodeinformation.com. Click the image for its enlargement"]"The dauer juvenile of entomopathogenic nematodes"[/caption]

    5. Life cycle of entomopathogenic nematodes

    As stated above, entomopathogenic nematodes complete most of their life cycle inside insect cadavers with an exception of infective/dauer juvenile, the only free-living stage found in the environment i.e. in the soil. Both Steinernema and Heterorhabditis infective juveniles locate an insect host and enter its body through natural body openings such as mouth, anus or spiracles. In addition, infective juveniles of Heterorhabditis species can also enter through the inter-segmental members of the host cuticle. Infective juveniles then actively penetrate through the mid-gut wall or tracheae into the insect body cavity also called hemocoel, which is filled with the insect blood also termed as haemolymph. Once in the hemocoel, infective juveniles release symbiotic bacteria from their intestine through anus in the insect haemolymph. Bacteria start multiplying in the nutrient-rich haemolymph and infective juveniles recover from their arrested state (dauer stage) and start feeding on multiplying bacteria and disintegrated host tissues. Toxins produced by the developing nematodes and multiplying bacteria in the body cavity kill the insect host usually within 48 hours.These bacteria also produce a plethora of metabolites, toxins and antibiotics with bactericidal, fungicidal and nematicidal properties, which ensures monoxenic conditions for nematode development and reproduction in the insect cadaver. Generally, if insect hosts such as wax worm larvae are infected with Steinernematid nematodes, they will turn creamy/beige/dark brown in color due to the metabolites produced by their symbiotic Xenorhabdus bacteria (Figs. 4 & 10) and if they are infected with Heterorhabditid nematodes, they will turn reddish/purplish in color to the metabolites produced by their symbiotic Photorhabdus bacteria (Figs. 5 & 11). [caption id="attachment_690" align="aligncenter" width="300" caption="Fig. 4. Beig colored Steinernematid nematode infected wax worm cadavers"]"Steinernematid nematodes infected wax worm cadavers"[/caption] [caption id="attachment_691" align="aligncenter" width="300" caption="Fig. 5. Red colored Heterorhabditis nematode infected wax worm cadavers"]"Heterorhabditis nematode infected wax worm cadavers"[/caption] Both heterorhabditid and steinernematid nematodes follow two slightly different reproduction pathways. For example, the first generation individuals of heterorhabditid nematodes are produced by self-fertile hermaphrodites (hermaphroditic) and their succeeding generations are produced by cross fertilization between males and females called as amphimictic type of reproduction.  In case of Steinernematid nematodes, with an exception of Steinernema hermaphroditum (Griffin et al., 2001; Stock et al., 2004), all generations are produced by cross fertilization between males and females. At the beginning eggs laid by females or hermaphrodites hatch and juveniles start feeding on the cadaver body tissue and bacterial soup. However, old females or hermaphrodites later do not lay eggs, which generally hatch only in the uterus of females. The hatched juveniles then start feeding on the mother’s tissues, the process is termed as “endotokia matricida” (Fig. 6; Johnigk and Ehlers, 1999). [caption id="attachment_447" align="aligncenter" width="300" caption="Fig. 6. After hatching from the eggs in the uterus, juveniles start feeding on mother’s tissues and this process is termed as Endotokia matricida"]“Endotokia matricida”[/caption] Depending on availability of food resources, both the heterorhabditid and steinernematid nematodes generally complete 2-3 generations within insect cadaver and emerge as infective juveniles to seek new hosts. Generally, life cycle of entomopathogenic nematodes starting from the penetration of infective juvenile into their hosts to the emergence of the infective juvenile from host cadavers is completed within 12- 15 days at room temperature (Fig. 7; adopted from http://www.nematodeinformation.com). The optimum temperature for growth and reproduction of most of the entomopathogenic nematode species is between 25 and 30oC (Grewal et al., 1994). [caption id="attachment_674" align="aligncenter" width="300" caption="Fig. 7. Life cycle of entomopathogenic nematodes. Adopted from www.nematodeinformation.com Click on a image for its enlargement."]"Life cycle of entomopathogenic nematodes"[/caption]

    6. How do entomopathogenic nematodes locate their insect hosts?

    Entomopathogenic nematode infective juveniles use following three types of foraging strategies to locate their insect hosts.

    a. Ambush foraging:

    Some entomopathogenic nematodes like Steinernema carpocapsae and S. scapterisci have adapted ambush foraging behavior known as “sit and wait” strategy to attack highly mobile insects including billbugs, sod webworms, cutworms, mole-crickets and armyworms at the surface of the soil.  These nematodes do not respond to host released cues but infective juveniles of some Steinernema spp can stand on their tails (nictate) and easily infect passing insect hosts by jumping on them.  Since highly mobile insects live in the upper soil or thatch layer, ambushers are generally effective in infecting more insects on the surface than deep in the soil.

    b. Cruise foraging:

    Cruiser entomomatogenic nematodes such as Heterorhabditis bacteriophora, H. megidis, Steinernema glaseri and S. kraussei are generally move actively in search of hosts and therefore, they found throughout the soil profile and more effective against less mobile hosts such as white grubs and larvae of black vine weevils.  These cruisers never nictate but generally respond to carbon dioxide released by insect hosts as cues.

    c. Intermediate foraging:

    Some entomopathogenic nematode species such as Steinernema feltiae and S.riobrave have adapted a foraging behavior that lie in between ambush and cruise strategies called an intermediate strategy to attack both the mobile and sedentary/less mobile insects at the surface or immobile stages deep in the soil.  Steinernema feltiae is highly effective against fungus gnats and mushroom flies whereas S.riobrave is effective against corn earworms, citrus root weevils and mole crickets.

    7. How are entomopathogenic nematodes produced?

    Currently, two different techniques including in vivo and in vitro are used for the mass production of entomopathogenic nematodes (Ehlers and Shapiro-Ilan, 2005).  Generally for a small-scale nematode production, in vivo technique is used whereas for a large-scale nematode production in vitro technique is used. In in vivo production technique, the nematode production is carried out in insect hosts; most commonly in last instar larvae of wax worms, Galleria mellonella (Fig. 8 ) or mealworms, Tenebrio molitor whereas in vitro production is carried out in solid or liquid media. Since in vitro technique is costly, needs a large infrastructure and installation, a thorough knowledge of bioreactor technology and biology of both entomopathogenic nematodes and their symbiotic bacteria, this blog focuses only on in vivo nematode production technique. For more information on in vitro nematode production technology read a book chapter by Ehlers and Shapiro-Ilan (2005). [caption id="attachment_455" align="aligncenter" width="300" caption="Fig. 8. Fourth stage wax worm Galleria melonella larvae used for in vio production of entomopathogenic nematodes."]"The wax worms"[/caption]

    In vivo production of entomopathogenic nematodes:

    Briefly, in this technique insect host larvae are inoculated with infective juveniles of entomopathogenic nematodes in dishes or in trays lined with a filter paper or any other available absorbent substrate (Fig. 9). For effective infection and optimum production, about 100 infective juveniles are used for infection of each wax worm or mealworm larva. The filter papers are generally used in dishes for absorption of excess nematode suspension so that insect larvae are not drowned in the suspension and infective juveniles can easily find moving insect host larvae for infection. Insects will die within 48 hours of infection (Figs. 4 and 5). After 48- 72 hours, the insect larval cadavers are transferred to the White traps (see below Figs. 10 and 11; White 1927). These white traps are then held in an incubator for 10-12 days at optimum temperature ranging from 18 to 28oC (Grewal et al., 1994). After 10-12 days into white traps, infective juveniles of entomopathogenic nematode generally start emerging from cadavers and moving into water. Emerged infective juveniles are then harvested from White traps, cleaned and concentrated by gravity settling (Dutky et al., 1964). These cleaned nematodes are ready for field applications or laboratory use. [caption id="attachment_696" align="aligncenter" width="300" caption="Fig. 9. A Petri dish lined with a filter paper for infecting insects with entomopathogenic nematodes."]"Petri dish lined with a filter paper for infection of insects"[/caption]

    8. How to make a White trap?

    For making White traps, you need one large size dish, a bottom or lid of a small size dish and a filter paper. As shown in Figs. 10 and 11, place a bottom or lid of a small dish inside the large size dish. Cover the bottom or lid of a small dish with a filter paper and then arrange cadavers on the filter paper. Then add enough quantity of water into large dish making sure that the filter paper is touching to water and becoming wet. Replace the lid of large dish and transfer into an incubator for 10-12 days. After 10-12 days, infective juveniles of entomopathogenic nematodes will emerge from cadavers and move into water. [caption id="attachment_476" align="aligncenter" width="300" caption="Fig. 10. A White trap containing entomopathogenic Steinernematid nematode infected wax worm larval cadavers. Click the image for its enlargement"]"White trap for Steinernematid nematode"[/caption] [caption id="attachment_477" align="aligncenter" width="300" caption="Fig. 11. A White trap containing entomopathogenic Heterorhabditis nematode infected wax worm larval cadavers. Click the image for its enlargement"]"White Trap for Heterorhabditis nematode"[/caption]

    9. What are similarities and differences between Steinernematid and Heterorhabditid nematodes?

    Similarities:

     

    Characteristics

    Steinernematid Nematodes

    Heterorhabditid Nematodes

    A single free-living and non-feeding infective/ dauer juvenile stage Present  Present
    Infective juveniles carry several cells of symbiotic bacterial in their guts Yes  Yes
    Infective juveniles enter into insect host’s body cavity through natural openings such as mouth, spiracles and anus Yes  Yes
    Once in the body cavity, symbiotic bacteria released by infective juveniles into insect blood through anus Yes Yes
    In insect blood, symbiotic bacteria quickly multiply, cause a disease and kill insect host within 48 hours of nematode infection (Griffin et al., 2005) Yes Yes
     

    Differences:

     

    Characteristics

     Steinernematid Nematodes

    Heterorhabditid Nematodes

    Taxonomic relationship (Stock and Hunt, 2005) No close relationship  No close relationship
    Type of reproduction (Griffin et al., 2005) Amphimictic reproduction: All generations are produced by a cross fertilization between males and females Both hermaphrodictic and amphimictic reproductions: In hermphrodictic reproduction, first generation individuals are produce by self-fertilization i.e. without males but the second generation individuals are produced by following amphimictic type of reproduction. 
    Number of infective juveniles need to enter into insect host’s body  At least two infective juveniles need to develop into a separate male and female individual for cross-fertilization and colonization  Only one infective juveniles need to develop as a hermaphrodite.
    Type of symbiotic  bacteria carried by infective juveniles Xenorhabdus spp. Photorhabdus spp.
     

    10. Why are entomopathogenic nematodes excellent and safe biological control agents?

    Entomopathogenic nematodes also called as beneficial nematodes belonging to both families, Steinernematidae and Heterorhabditidae are considered as safe and excellent biological control agents against many soil dwelling insect pests (Table 2) of many economically important crops because…..
    • they have a broad host range
    • their ability to search actively for hosts
    • their ability to kill their hosts rapidly within 24-48 hours
    • they have potential to recycle in the soil environment
    • they have no deleterious effects on humans, other vertebrate animals, non-target organisms and plants
    • they have no negative effects on environment
    • they can be easily mass produced using both in vivo and in vitro methods and applied using traditional insecticide spraying equipments
    • they are compatible with many chemical insecticides and biopesticides and therefore,  easily included in IPM programs
    • there is no fear of developing resistance in their insect hosts as these nematodes physically enter into the insect host's body cavity where they release symbiotically associated bacteria and kill insect host within 48 hours.
    • Because of their safety to the environment and human health, they also been exempted from registration and regulation requirement by US Environmental Protection Agency (EPA) and similar agencies in many other countries

    11. How many nematodes do I need to apply for the successful control of target pests?

    For the successful control of most of the soil dwelling insect pests, the optimal rate of 1 billion infective juvenile nematodes in 100 to 260 gallons of water per acre is generally recommended (See Table 1).  

    12. How are entomopathogenic nematodes applied?

    Please read our previous blog for appropriate methods of nematode application.  

    References

    Dutky, S. R., Thompson, J. V. and Cantwell, G. E. 1964.  A technique for the mass propagation of the DD-136 nematode. Journal of Insect Pathology 6, 417- 422. Ehlers, R.-U. and Shapiro-Ilan, D. I. 2005. Mass production. In: Nematodes as biocontrol agents. Grewal, P. S., Ehlers, R.-U. and Shapiro-Ilan, D. I. (Eds.). CABI Publishing,UK. pp. 65-78. Grewal, P.S., Ehlers, R.U. and Shapiro-Ilan, D. I. [Editors]. 2005. Nematodes As Biocontrol Agents, CABI Publishing, Wallingford, UK, pp 1-505. Grewal, P.S., Selvan, S., Gaugler, R., 1994.  Thermal adaptation of entomopathogenic nematodes: Niche breadth for infection, establishment, and reproduction. J. Therm. Biol. 19, 245-53. Griffin, C.T., Boemare, N.E. and Lewis, E.E. Biology and behaviour. In: Nematodes as biocontrol agents. Grewal, P. S., Ehlers, R.-U. and Shapiro-Ilan, D. I. (Eds.). CABI Publishing, UK. pp. 47-64. Griffin, C.T., O'Callaghan, K.M. and Dix, I. 2001. A self-fertile species of Steinernema from Indonesia: further evidence of convergent evolution amongst entomopathogenic nematodes? Parasitology 122: 181-186. Han, R. and Ehlers, R. 2001. Effect of Photorhabdus luminescens phase variants on the in vivo and in vitro development and reproduction of the entomopathogenic nematodes Heterorhabditis bacteriophora and Steinernema carpocapsae. FEMS Microbiological Ecology 35: 239-247. Jagdale, G.B. and Gordon, R. 1997.  Effect of temperature on the composition of fatty acids in total lipids and phospholipids of entomopathogenic nematodes. Journal of Thermal Biology 22: 245-251. Jagdale, G.B. and Grewal, P.S. 2003.  Acclimation of entomopathogenic nematodes to novel temperatures: trehalose accumulation and the acquisition of thermotolerance. International Journal for Parasitology 33: 145-152. Jagdale, G. B. and Grewal, P. S. 2007.  Storage temperature influences desiccation and ultra violet radiation tolerance of entomopathogenic nematodes. Journal of Thermal Biology 32: 20-27. Jagdale, G. B., Grewal, P. S. and Salminen, S. O. 2005.  Both heat-shock and cold-shock influence trehalose metabolism in entomopathogenic nematodes. Journal of Parasitology 91: 988-994. Johnigk, S.-A., and Ehlers, R.-U. 1999. Endotokia matricida in hermaphrodites of Heterorhabditis spp and the effect of the food supply. Nematology 1, 717–726. Stock, S. P. and Hunt, D. J., 2005, Morphology and systematics of nematodes used in biocontrol. In: Nematodes as biocontrol agents. Grewal, P. S., Ehlers, R.-U. and Shapiro-Ilan, D. I. (Eds.). CABI Publishing,UK. pp. 3-43. Stock, S.P., Griffin, C.T., and Haerani, R.C. 2004. Morphological and molecular characterization of Steinernema hermaphroditum n. sp. (Nematoda: Steinernematidae), an entomopathogenic nematode from Indonesia, and its phylogenetic relationships with other members of the genus. Nematology 6: 401- 412. White, G.F., 1927.  A method for obtaining infective nematode larvae from cultures. Science 66, 302-303.

    Economically important insect pests and their susceptibility to major entomopathogenic nematodes

    Table 2. List of species of insect pests that are susceptible to major entomopathogenic nematodes

    Species of insect pests

     Entomopathogenic nematode species

    Publications

    (See below)

    Apopka weevil, Citrus root weevil or Sugarcane borer Diaprepes abbreviatus Heterorhabditis georgiana, H. indica, H. zealandica, Steinernema carpocapsae, S. diaprepesi, S. riobrave 1-13
    Armyworms, Helicoverpa (Heliothis) armigeraSpodoptera exigua, S. frugiperda H. amazonensis, H. indica S. arenarium, S. carpocapsae, S. glaseri 14-18
    Billbugs, Sphenophorus purvulusS. levis H. bacteriophora, S. brazilense, S. carpocapsae 19-20
    Black vine weevil, Otiorhynchus salcatus H. bacteriophora, H. downesi, H. megidi.S. carpocapsaeS. feltiae, S. glaseri, S. kraussei  21-26
    Bluegrass weevil, Listronotus maculicollis H. bacteriophora, S. carpocapsae 27-29
    Carpenter worms, Cossus cossus S. weiseri 30
    Carrot weevil, Listronotus oregonensis H. bacteriophora, H. megidi, S. feltiae, S. carpocapsae, S. riobrave,  feltiae  31-32
    Cat fleas, Ctenocephalides felis S. carpocapsae 33-34
    Chestnut weevil, Curculio elephas H. bacteriophora, S. carpocapsaeS. feltiae,  S. siamkayai, S. weiseri 35-37
    Chinch bugs, Blissus sp. Unknown species 38
    Citrus root weevil, Pachnaeus litus S. carpocapsae 39-41
    Clover root weevil, Sitona hispidulus H. bacteriophora 42-43
    Codling moth, Cydia pomonella H. bacteriophora, H. zealandica, S. carpocapsae, S. feltiae, S. kraussei 44-56
    Crane flies, Tipula paludosa H. marelatus, H. megidis, S. carpocapsae, S. feltiae 57-58
    Cucurbit beetle, Diabrotica speciosa H. amazonensis, S. glaseri 59
    Cutworms, Agrotis ipsilon, A. segetum H. bacteriophora, H. georgiana, H. indica H. Mexicana, S. carpocapsae, S. feltiae, S. riobrave 60-65
    Diamondback moth, Plutella xylostella Heterorhabditis sp., Rhabditis blumi, S. carpocapsae 66-70
    Egyptian cotton leaf worm, Spodoptera littoralis H. bacteriophora, S. glaseri, S. feltiae, S. carpocapsae, S. kraussei, S. riobrave 71-73
    Fall webworms, Hyphantria cunea H. bacteriophora, S. feltiae  74
    Filbertworm, Cydia latiferreana S. carpocapsae, S. kraussei 75-76
    Flea beetles, Phyllotreta striolata, P. cruciferae H. bacteriophora, H. indica, H. megidi, S. carpocapsae, S. feltiae, S. pakistanense 77-79
    Fungus gnats, Bradysis spp.   H. bacteriophora, H. indica, H. zealandica, S. anomali, S. carpocapsae, S. feltiae, S. riobrave 80-84
    House flies, Musca domestica H. bacteriophora,  H. megidi, S. carpocapsae, S. feltiae, S. scapterisci  85-89
    Japanese beetle, Popillia japonica, P. unipuncta H. bacteriophora, H. indica, H. marelata, H. megidis, H. zealandica, S. anomaly, S. carpocapsae, S. feltiae, S. glaseri, S. kushidai, S. minuta, S. scapterisci, S. scarabae,  S. riobrave 90-99
    Leaf minors, Liriomyza bryoniae, L. trifolii, L. huidobrensis S. carpocapsae, S. feltiae 100-107
    Leopard moth, Zeuzera pyrina H. bacteriophora, H. heliothidis, S. carpocapsae 108
    Mediterranean fruit flyCeratitis capitata H. bacteriophora, H. zealandica, S. carpocapsae, S. feltiae, S. khoisanae, S. siamkayai, S. weiseri 109-116
    Mole cricketsScapteriscus vicinus S. carpocapsae, S. riobravis, S. scapterisci 117-131
    Navel orangeworm, Amyelois transitella S. carpocapsae 132
    Peach borer, Synanthedon exitiosa H. bacteriophora, S. carpocapsae, S. riobrave 133-134
    Pecan weevil, Curculio caryae, C. hicoriae H. bacteriophora, H. indica, H. megidis, H. Mexicana, S. carpocapsae, S. riobrave 135-143
    Pine weevil, Hylobius abietis H. downesi, H. megidis, S. carpocapsae, S. feltiae 144-148
    Plum weevil, Conotrachelus nenuphar H. bacteriophora, S. carpocapsae, S. feltiae, S. riobrave  149-154
    Shore flies, Scatella stagnalis, S. tenuicosta H. bacteriophora, H. megidis, S. anomaly, S. arenarium, S. carpocapsae, S. feltiae 155-158
    Sod webworm, Herpetogramma phaeopteralis S. carpocapsae, S. feltiae 159
    Spruce webworm, Cephalcia abietis S. feltiae 160
    Stable fly, Stomoxys calcitrans H. heliothidis, S. glaseri 161
    Stored grain pests: Indian meal moth (Plodia interpunctella), Mediterranean flour moth (Ephestia kuehniella), Sawtoothed grain beetle (Oryzaephilus surinamensis), Mealworm (Tenebrio molitor), Red flour beetle (Tribolium castaneum), Warehouse beetle (Trogoderma variabile) H. bacteriophora, H. megidis, S. carpocapsae, S. feltiae 162-168
    Strawberry root borer, Nemocestes incomptus S. carpocapsae 169
    Strawberry root weevil, Otiorhynchus ovatus, O. dubius strom, Ptiorhynchus ovatus H. bacteriophora, H. marelatus, S. carpocapsae 170-173
    Strawberry crown moth, Synanthedon bibionipennis H. bacteriophora, S. carpocapsae  174
    Tick, Rhipicephalus (Boophilus) microplus H. amazonensis, S. carpocapsae, S. glaseri  175-179
    Western flower thrips, Frankliniella occidentalis, Thrips palmi H. bacteriophoraH. indica, S. arenariumS. bicornutum, S. carpocapsae, S. feltiae, Thripinema nicklewoodi 180-186
    Western corn rootworm, Diabrotica virgifera virgifera H. bacteriophora, S. carpocapsae 187-189
    White flies, Bemisia tabaci, Trialeurodes vaporariorum H. bacteriophora, H. megidis, S. feltiae 190-194
    White grub (Summer Chafer), Amphimallon solstitiale H. bacteriophora 195
    White grub (Oriental beetle), Anomala orientalis, Exomala orientalis, Blitopertha orientalis H. bacteriophoraH. megidis, H. zealandica, S. carpocapsaeS. glaseri, S. longicaudum, S. scarabae 196-216
    White grub, Costelytra zealandica H. bacteriophora, S. glaseri 217
    White grub (June Bettle), Cotinus nitida H. bacteriophora, S. carpocapsae, S. feltiae, S. glaseri, S. scarabae 218-220
    White grub, Cyclocephala borealis, C. hirta, C. lurida, C. pasadenae H. bacteriophoraH. indicaH. marelata, H. megidisH. zealandica, S. carpocapsae,  S. feltiae, S. glaseri, S. kushidai, S. riobrave, S. scarabae 221-227
    White grub, Hoplia philanthus H. bacteriophora, H. indica, H. megidis, S. arenarium, S. carpocapsae, S. feltiae, S. glaseri, S. scarabaei  228-232
    White grub, Melolontha melolontha H. bacteriophoraH. marelata, H. megidisS. arenariaS. feltiaeS. glaseri, S. riobrave 233-235
    White grub, Ataenius spretulus H. bacteriophoraS. glaseri, S. scarabae 236-237
    White grub (Asiatic garden beetle), Maladera castanea H. bacteriophoraS. glaseri, S. scarabae 238-242
    White grubs, Phyllophaga anxia, P. bicolor, P. congrua, P. crinita, P. georgiana, P. hirticula, P. menetriesi H. bacteriophora, H. heliothidis, H. zealandica, S. carpocapsae, S. feltiae, S. glaseri, S. riobrave, S. scarabae  243-250
    White grub, Rhizotrogus majalis H. bacteriophoraH. megidis, H. zealandicaS. carpocapsaeS. feltiae, S. glaseri, S. scarabae 251-255
    Fuller rose beetle, Asynonychus godmani S. carpocapsae 256
    Chive gnat, Bradysia odoriphaga H. bacteriophora, H. indica, H. megidis, S. ceratophorum, S. feltiae, S. hebeiense, S. litorale  257-258
     

    Publications:

    Apopka weevil, Diaprepes abbreviatus 1. Ali, J.G., Alborn, H.T. and Stelinski, L.L. 2010.  Subterranean herbivore-induced volatiles released by citrus roots upon feeding by Diaprepes abbreviatus recruit entomopathogenic nematodes. Journal of Chemical Ecology. 36: 361-368. 2. 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. 3. Duncan, L. W., Stuart, R. J., El-Borai, F. E., Campos-Herrera, R., Pathak, E., Giurcanu, M. and Graham, J. H. 2013. Modifying orchard planting sites conserves entomopathogenic nematodes, reduces weevil herbivory and increases citrus tree growth, survival and fruit yield. Biological Control 64: 26-36. 4. 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. 5. 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. 6. El-Borai, F.E., Stuart, R.J., Campos-Herrera, R., Pathak, E. and Duncan, L.W. 2012.  Entomopathogenic nematodes, root weevil larvae, and dynamic interactions among soil texture, plant growth, herbivory, and predation. Journal of Invertebrate Pathology 109: 134-142. 7. Kaspi, R., Ross, A., Hodson, A.K., Stevens, G.N., Kaya, H.K. and Lewis, E.E. 2010. Foraging efficacy of the entomopathogenic nematode Steinernema riobrave in different soil types from California citrus groves. Applied Soil Ecology 45: 243-253. 8. Schroeder, W.J. 1992. Entomopathogenic nematodes for control of root weevils of citrus. Florida Entomologist 75: 563-567. 9. Shapiro, D.I. and McCoy, C.W. 2000. Susceptibility of Diaprepes abbreviatus (Coleoptera: Curculionidae) larvae to different rates of entomopathogenic nematodes in the greenhouse. Florida Entomologist. 83: 1-9. 10. Shapiro, D.I. and McCoy, C.W. 2000. 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. 11. 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. 12. Shapiro-Ilan, D.I., Mbata, G.N., Nguyen, K.B., Peat, S.M., Blackburn, D. and Adams, B.J. 2009. Characterization of biocontrol traits in the entomopathogenic nematode Heterorhabditis georgiana (Kesha strain), and phylogenetic analysis of the nematode's symbiotic bacteria. Biological Control 51: 377-387. 13. 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 tumidaJournal of Invertebrate Pathology. 103: 103-108. Armyworms, Heliothis armiger, Spodoptera exigua, S. frugiperda 14. Andalo, V., Santos, V., Moreira, G.F., Moreira, C., Freire, M. and Moino, A. 2012.   Movement of Heterorhabditis amazonensis and Steinernema arenarium in search of corn fall armyworm larvae in artificial conditions. Scientia Agricola 69: 226-230.  15. Ansari, M.A., Waeyenberge, L. and Moens, M. 2007.  Natural occurrence of Steinernema carpocapsae, Weiser, 1955 (Rhabditida: Steinernematidae) in Belgian turf and its virulence to Spodoptera exigua (Lepidoptera: Noctuidae). Russian Journal of Nematology 15: 21-24. 16. Kim, J. and Kim, Y. 2011. Three metabolites from an entomopathogenic bacterium, Xenorhabdus nematophila, inhibit larval development of Spodoptera exigua (Lepidoptera: Noctuidae) by inhibiting a digestive enzyme, phospholipase A (2). Insect Science 18: 282-288. 17. Negrisoli, A.S., Garcia, M.S., Negrisoli, C.R.C.B., Bernardi, D. and da Silva, A. 2010.  Efficacy of entomopathogenic nematodes (Nematoda: Rhabditida) and insecticide mixtures to control Spodoptera frugiperda (Smith, 1797) (Lepidoptera: Noctuidae) in corn crops. Crop Protection. 29: 677-683. 18. Salvadori, J.D., Defferrari, M.S., Ligabue-Braun, R., Lau, E.Y., Salvadori, J.R. and Carlini, C.R. 2012. Characterization of entomopathogenic nematodes and symbiotic bacteria active against Spodoptera frugiperda (Lepidoptera: Noctuidae) and contribution of bacterial urease to the insecticidal effect. Biological Control 63: 253-263.       Billbugs, Sphenophorus spp. 19. Georgis, R., Koppenhofer, A.M., Lacey, L.A., Belair, G., Duncan, L.W., Grewal, P.S., Samish, M., Tan, L., Torr, P. and van Tol, R.W.H.M. 2006. Successes and failures in the use of parasitic nematodes for pest control. Biological Control 38: 103-123. 20. Giometti, F.H.C., Leite, L.G., Tavares, F.M., Schmit, F.S., Batista, A. and Dell'Acqua, R. 2011.  Virulence of entomopathogenic nematodes (Nematoda: Rhabditida) against Sphenophorus levis (Coleoptera: Curculionidae). Bragantia 70: 81-86.   Black vine weevil, Otiorhynchus sulcatus 21. Ansari, M. A. and Butt, T. M. 2011.  Effect of potting media on the efficacy and dispersal of entomopathogenic nematodes for the control of black vine weevil, Otiorhynchus sulcatus (Coleoptera: Curculionidae). Biological Control 58: 310-318. 22. Ansari, M.A., Shah, F.A. and Butt, T.M. 2008.  Combined use of entomopathogenic nematodes and Metarhizium anisopliae as a new approach for black vine weevil, Otiorhynchus sulcatus control. Entomologia Experimentalis Et Applicata 129: 340-347. 23. 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. 24. Haukeland, S. and Lola-Luz, T. 2010.  Efficacy of the entomopathogenic nematodes, Steinernema kraussei and Heterorhabditis megidis against the black vine weevil Otiorhynchus sulcatus in open field-grown strawberry plants. Agricultural and Forest Entomology.12363-369. 25. 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. 26. 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. Bluegrass weevil, Listronotus maculicollis 27. McGraw, B.A. and Koppenhofer, A.M.2008.  Evaluation of two endemic and five commercial entomopathogenic nematode species (Rhabditida: Heterorhabditidae and Steinernematidae) against annual bluegrass weevil (Coleoptera: Curculionidae) larvae and adults. Biological Control 46: 467-475. 28. McGraw, B.A. and Koppenhofer, A.M.2009.  Population dynamics and interactions between endemic entomopathogenic nematodes and annual bluegrass weevil populations in golf course turfgrass. Applied Soil Ecology 41: 77-89. 29. McGraw, B.A., Vittumb, P.J. Cowlesc, R.S.and Koppenhoumlfera, A.M. 2010.  Field evaluation of entomopathogenic nematodes for the biological control of the annual bluegrass weevil, Listronotus maculicollis (Coleoptera: Curculionidae), in golf course turfgrass. Journal Biocontrol Science and Technology. 20: 149 – 163. Carpenter worms, Cossus cossus 30. Bazman, I., Ozer, N., and Hazir, S. 2008.  Bionomics of the entomopathogenic nematode, Steinernema weiseri (Rhabditida: Steinernematidae). Nematology 10: 735-742. Carrot weevil, Listronotus oregonensis 31. Belair, G. and Boivin, G.  1995. Evaluation of Steinernema-carpocapsae weiser for control of carrot weevil adults, Listronotus-oregonensis (leconte) (coleopteran: curculionidae), in organically grown carrots. Biocontrol Science and Technology 5: 225-231. 32. Miklasiewicz, T.J., Grewal, P.S., Hoy, C.W. and Malik, V.S. 2002. Evaluation of entomopathogenic nematodes for suppression of carrot weevil. Biocontrol 47: 545-561. Cat fleas, Ctenocephalides felis 33. Henderson, G., Manweiler, S.A., Lawrence, W.J., Tempelman, R.J.and Foil, L.D. 1995.  The effects of Steinernema-carpocapsae (weiser) application to different life stages on adult emergence of the cat flea Ctenocephalides-felis (bouche). Veterinary Dermatology 6: 159-163. 34. Silverman J.S., Platzer, E.G. and M.K. Rust, M.K. 1982. Infection of the cat flea, Ctenocephalides felis (Bouche) by Neoaplectana carpocapsae Weiser. Journal of Nematology 14: 394-397. Chestnut weevil, Curculio elephas 35. Karagoz, M., Gulcu, B., Hazir, S. and Kaya, H.K. 2009.  Laboratory evaluation of Turkish entomopathogenic nematodes for suppression of the chestnut pests, Curculio elephas (Coleoptera: Curculionidae) and Cydia splendana (Lepidoptera: Tortricidae). Biocontrol Science and Technology. 19: 755-768. 36. Kepenekci, I., Gokce, A. and Gaugler, R. 2004.  Virulence of three species of entomopathogenic nematodes to the chestnut weevil, Curculio elephas (Coleoptera: Curculionidae). Nematropica 34: 199-204. 37. Raja, R.K., Sivaramakrishnan, S. and Hazir, S. 2011.   Ecological characterisation of Steinernema siamkayai (Rhabditida: Steinernematidae), a warm-adapted entomopathogenic nematode isolate from India. Biocontrol 56: 789-798. Chinch bugs Bilssus spp. 38. Baxendale, F.P., A.P. Weinhold, and T.P. Riordan. 1994. Control of buffalograss chinch bugs with Beauvaria bassiana and entomopathogenic nematodes, 1993. Nebraska Insect Management and Insecticide Efficacy Reports, Dept. of Entomology Report No. 18, Univ. of Nebr., p. 43. Citrus root weevil, Pachnaeus litus 39. 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.  40. Duncan, L.W., Graham, J.H., Dunn, D.C., Zellers, J., McCoy, C.W. and Nguyen, K. 2003.  Incidence of endemic entomopathogenic nematodes following application of Steinerema riobrave for control of Diaprepes abbreviates. Journal of Nematology 35: 178-186. 41. Schroeder, W.J. 1992. Entomopathogenic nematodes for control of root weevils of citrus. Florida Entomologist 75: 563-567. Clover root weevil, Sitona hispidulus 42. Loya, L.J. and Hower, A.A. 2002.  Population dynamics, persistence, and efficacy of the entomopathogenic nematode Heterorhabditis bacteriophora (Oswego strain) in association with the clover root curculio (Coleoptera: Curculionidae) in Pennsylvania.   Environmental Entomology 31: 1240-1250. 43. Loya, L.J. and Hower, A.A. 2003. Infectivity and reproductive potential of the Oswego strain of Heterorhabditis bacteriophora associated with life stages of the clover root curculio, Sitona hispidulus.   Journal of Invertebrate Pathology 83: 63-72. Codling moth, Cydia pomonella 44. Cossentine, J.E., Jensen, L.B. and Moyls, L. 2002. Fruit bins washed with Steinernema carpocapsae (Rhabditida: Steinernematidae) to control Cydia pomonella (Lepidoptera: Tortricidae). Biocontrol Science and Technology 12: 251-258. 45. de Waal, J.Y., Malan, A.P. and Addison, M.F. 2011.  Evaluating mulches together with Heterorhabditis zealandica (Rhabditida: Heterorhabditidae) for the control of diapausing codling moth larvae, Cydia pomonella (L.) (Lepidoptera: Tortricidae). Biocontrol Science and Technology 21: 255-270. 46. de Waal, J.Y., Malan, A.P., Levings, J. and Addison, M.F. 2010.  Key elements in the successful control of diapausing codling moth, Cydia pomonella (Lepidoptera: Tortricidae) in wooden fruit bins with a South African isolate of Heterorhabditis zealandica (Rhabditida: Heterorhabditidae). Biocontrol Science and Technology. 20: 489-502. 47. Lacey, L.A. and Chauvin, R.L. 1999. Entomopathogenic nematodes for control of diapausing codling moth (Lepidoptera: Tortricidae) in fruit bins. Journal of Economic Entomology 92: 104-109. 48. Lacey, L.A., and Unruh, T.R. 1998. Entomopathogenic nematodes for control of codling moth, Cydia pomonella (Lepidoptera: Tortricidae): Effect of nematode species, concentration, temperature, and humidity.  Biological Control 13: 190-197. 49. Lacey, L.A., Arthurs, S.P., Unruh, T.R., Headrick, H. and Fritts, R. 2006. Entomopathogenic nematodes for control of codling moth (Lepidoptera: Tortricidae) in apple and pear orchards: Effect of nematode species and seasonal temperatures, adjuvants, application equipment, and post-application irrigation. Biological Control 37: 214-223. 50. Lacey, L.A., Granatstein, D., Arthurs, S.P., Headrick, H. and Fritts, R. 2006. Use of entomopathogenic nematodes (Steinernematidae) in conjunction with mulches for control of overwintering codling moth (Lepidoptera: Tortricidae). Journal of Entomological Science 41: 107-119. 51. Lacey, L.A., Neven, L.G., Headrick, H.L. and Fritts, R. 2005.   Factors affecting entomopathogenic nematodes (Steinerneniatidae) for control of overwintering codling moth (Lepidoptera: Tortricidae) in fruit bins. Journal of Economic Entomology 98: 1863-1869. 52. Lacey, L.A., Shapiro-Ilan, D.I. and Glenn, G.M. 2010.   Post-application of anti-desiccant agents improves efficacy of entomopathogenic nematodes in formulated host cadavers or aqueous suspension against diapausing codling moth larvae (Lepidoptera: Tortricidae). Biocontrol Science and Technology. 20: 909-921. 53. Mracek, Z., Becvar, S., Kindlmann, P. and Webster, J.M. 1998.  Infectivity and specificity of Canadian and Czech isolates of Steinernema kraussei (Steiner, 1923) to some insect pests at low temperatures in the laboratory.  Nematologica 44: 437-448. 54. Navaneethan, T., Strauch, O., Besse, S., Bonhomme, A. and Ehlers, R.U. 2010.  Influence of humidity and a surfactant-polymer-formulation on the control potential of the entomopathogenic nematode Steinernema feltiae against diapausing codling moth larvae (Cydia pomonella L.) (Lepidoptera: Tortricidae). Biocontrol 55: 777-788. 55. Unruh, T.R., and Lacey, L.A. 2001. Control of codling moth, Cydia pomonella (Lepidoptera: Tortricidae), with Steinernema carpocapsae: Effects of supplemental wetting and pupation site on infection rate.  Biological Control 20: 48-56. 56. Vega, F.E., Lacey, L.A., Reid, A.P., Herard, F., Pilarska, D., Danova, E., Tomov, R. and Kaya, H.K. 2000.  Infectivity of a Bulgarian and an American strain of Steinernema carpocapsae against codling moth. Biocontrol 45: 337-343.  Crane flies, Tipula paludosa 57. Oestergaard, J., Belau, C., Strauch, O., Ester, A., van Rozen, K. and Ehlers, R.U. 2006.  Biological control of Tipula paludosa (Diptera: Nematocera) using entomopathogenic nematodes (Steinernema spp.) and Bacillus thuringiensis subsp israelensis. Biological Control 39: 525-531. 58. Simard, L., Belair, G., Gosselin, M.E. and Dionne, J. 2006.  Virulence of entomopathogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae) against Tipula paludosa (Diptera: Tipulidae), a turfgrass pest on golf courses. Biocontrol Science and Technology 16: 789-801. Cucurbit beetle, Diabrotica speciosa 59. Santos, V., Moino, A., Andalo, V., Moreira, C.C. and de Olinda, R.A. 2011. Virulence of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) for the control of Diabrotica speciosa Germar (Coleoptera: Chrysomelidae). Ciencia e Agrotecnologia 35: 1149-1156.  Cutworms, Agrotis ipsilon and A. segetum 60. Ebssa, L. and Koppenhofer, A.M.  2011. Efficacy and persistence of entomopathogenic nematodes for black cutworm control in turfgrass.   Biocontrol Science and Technology 21: 779-796. 61. Kunkel, B.A., Grewal, P.S. and Quigley, M.F. 2004.  A mechanism of acquired resistance against an entomopathogenic nematode by Agrotis ipsilon feeding on perennial ryegrass harboring a fungal endophyte.   Biological Control 29: 100-108. 62. Richmond, D.S., and Bigelow, C.A. 2009.  Variation in endophyte-plant associations influence Black Cutworm (Lepidoptera: Noctuidae) performance and susceptibility to the parasitic nematode Steinernema carpocapsae.  Environmental Entomology 38: 996-1004. 63. Shamseldean, M.M., Ibrahim, A.A., Zohdi, N., Shairra, S.A. and Ayaad, T.H. 2008.  Effect of Egyptian entomopathogenic nematode isolates on some economic insect pests.   Egyptian Journal of Biological Pest Control 18: 81-89. 64. Shapiro-Ilan, D.I., Mbata, G.N., Nguyen, K.B. Peat, S.M., Blackburn, D. and Adams, B.J. 2009. Characterization of biocontrol traits in the entomopathogenic nematode Heterorhabditis georgiana (Kesha strain), and phylogenetic analysis of the nematode’s symbiotic bacteria. Biological Control. 51: 377-387. 65. Shapiro-Ilan, D.I., Stuart, R.J. and McCoy, C.W. 2005. Characterization of biological control traits in the entomopathogenic nematode Heterorhabditis mexicana (MX4 strain). Biological Control 32: 97-103. Diamondback moth, Plutella xylostella 66. Baur M.E., Kaya H.K., Gaugler R. and Tabashnik, B.E. 1997. Effects of adjuvants on entomopathogenic nematode persistence and efficacy against Plutella xylostella, Biocontrol Science and Technology 7: 513–525. 67. Park, H.W., Kim, H.H., Youn, S.H., Shin, T.S., Bilgrami, A.L., Cho, M.R. and Shin, C.S. 2012. Biological control potentials of insect-parasitic nematode Rhabditis blumi (Nematoda: Rhabditida) for major cruciferous vegetable insect pests. Applied Entomology and Zoology 47: 389-397. 68. Schroer S. and Ehlers R.U. 2005.  Foliar application of the entomopathogenic nematode, Steinernema carpocapsae for biological control of diamond black moth larvae (Plutella xylostella). Biological Control 33: 81–86. 69. Schroer S., Sulistyanto D. and Ehlers R.U. 2005. Control of Plutella xylostella using polymerformulated Steinernema carpocapsae and Bacillus thuringiensis in cabbage fields. Journal of Applied Nematology 129:198–204. 70. Schroer S., Ziermann D., Ehlers R.U. 2005. Mode of action of a surfactant-polymer formulation to support performance of the entomopathogenic nematode Steinernema carpocapsae for control of diamondback moth larvae (Plutella xylostella). Biocontrol Science and Technology 15:601–613. Egyptian cotton leaf worm, Spodoptera littoralis 71. Campos-Herrera, R. and Gutierrez, C. 2008.  Comparative study of entomopathogenic nematode isolation using Galleria mellonella (Pyralidae) and Spodoptera littoralis (Noctuidae) as baits. Biocontrol Science and Technology 18: 629-634. 72. Hassan, H.A. and Ibrahim, S.A.M. 2010.  Immune response of the cotton leaf worm Spodoptera littoralis (Biosd.) towards entomopathogenic nematodes. Egyptian Journal of Biological Pest Control 20: 45-53. 73. Ibrahim, A.A. and Shairra, S.A. 2011.  Effect of eicosanoid biosynthesis inhibitors on the immune response of the Cotton Leaf Worm, Spodoptera littoralis (Boisd.) infected with the nematode, Steinernema glaseri (Rhabditida: Steinernematidae). Egyptian Journal of Biological Pest Control 21: 197-202. Fall webworms, Hyphantria cunea 74. Chkhubianishvili, T., Mikaia, N., Malania, I. and Kakhadze, M. 2007. Susceptibility of entomopathogenic nematodes to the fall webworm Hyphantria cunea Drury (Lepidoptera: Arctiidae). Bulletin of the Georgian National Academy of Sciences 175: N2.  Filbertworm, Cydia latiferreana 75. Bruck, D.J. and Walton, V.M. 2007.  Susceptibility of the filbertworm (Cydia latiferreana, Lepidoptera: Tortricidae) and filbert weevil (Curculio occidentalis, Coleoptera: Curculionidae) to entomopathogenic nematodes. Journal of Invertebrate Pathology. 96: 93–96. 76. Chambers, U. Bruck, D.J., Olsen, J. and Walton, V.M. 2010.  Control of overwintering filbertworm (Lepidoptera: Tortricidae) larvae with Steinernema carpocapsae. Journal of Economic Entomology. 103: 416-422.    Flea beetles, Phyllotreta spp. 77. Morris, O.N. 1987.  Evaluation of the nematode, Steinernema-feltiae filipjev, for the control of the crucifer flea beetle, phyllotreta-cruciferae (goeze) (coleoptera, chrysomelidae). Canadian Entomologist 119: 95-101. 78. Trdan, S., Vidrih, M., Valic, N. and Laznik, Z. 2008. Impact of entomopathogenic nematodes on adults of Phyllotreta spp. (Coleoptera: Chrysomelidae) under laboratory conditions. Acta Agriculturae Scandinavica Section B-Soil and Plant Science 58: 169-175. 79. Xu, C.X., De Clercq, P., Moens, M., Chen, S.L and Han, R.C. 2010. Efficacy of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) against the striped flea beetle, Phyllotreta striolata. Biocontrol 55: 789-797. Fungus gnats, Bradysia coprophila 80. 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. Biological Control 5, 412-418. 81. Jagdale, G. B., Casey, M. L., Grewal, P. S. and Lindquist, R. K. 2004.  Application rate and timing, potting medium and host plant on the efficacy of Steinernema feltiae against the fungus gnat, Bradysia coprophila, in floriculture. Biological Control 29: 296-305. 82. Jagdale, G. B., Casey, M. L., Grewal, P. S. and Luis Cañas. 2007.  Effect of entomopathogenic nematode species, split application and potting medium on the control of the fungus gnat, Bradysia difformis (Diptera: Sciaridae), in the greenhouse at alternating cold and warm temperatures. Biological Control 43: 23-30. 83. 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 Science and Technology 14, 171-183. 84. 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. House flies, Musca domestica 85. Renn, N. 1995. Mortality of immature houseflies (Musca-domestica l) in artificial diet and chicken manure after exposure to encapsulated entomopathogenic nematodes (Rhabditida, Steinernematidae, heterorhabditidae).  Biocontrol Science and Technology 5: 349-359. 86. Renn, N. 1998. Routes of penetration of the entomopathogenic nematode Steinernema feltiae attacking larval and adult houseflies (Musca domestica).  Journal of Invertebrate Pathology 72: 281-287. 87. Renn, N. 1998. The efficacy of entomopathogenic nematodes for controlling housefly infestations of intensive pig units. Medical and Veterinary Entomology 12: 46-51. 88. Renn, N. and Wright, E. 2000. The effect of artificial substrates on the pathogenicity of Steinernema feltiae (Rhabditida: Steinernematidae) to adult Musca domestica (Diptera: Muscidae). Nematology 2: 217-222. 89. Taylor, D.B., Szalanski, A.L., Adams, B.J. and Peterson, R.D. 1998. Susceptibility of house fly (Diptera: Muscidae) larvae to entomopathogenic nematodes (Rhabditida: Heterorhabditidae, Steinernematidae). Environmental Entomology 27: 1514-1519. Japanese beetle, Popillia japonica 90. 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. 91. Grewal, P.S., Koppenhofer, A.M., and Choo, H.Y., 2005.  Lawn, turfgrass and Pasture applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166. 92. Hao, Y.J., Montiel, R., Lucena, M.A., Costa, M. and Simoes, N. 2012. Genetic diversity and comparative analysis of gene expression between Heterorhabditis bacteriophora Az29 and Az36 isolates: Uncovering candidate genes involved in insect pathogenicity. Experimental Parasitology 130: 116-125. 93. 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. 94. 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. 95. Maneesakorn, P., An, R., Grewal, P.S.and Chandrapatya, A. 2010. Virulence of our new strains of entomopathogenic nematodes from Thailand against second instar larva of the Japanese beetle, Popillia japonica (Coleoptera: Scarabaeidae). Thai Journal of Agricultural Science.43: 61-66. 96. Morales-Rodriguez, A. and Peck, D.C. 2009.  Synergies between biological and neonicotinoid insecticides for the curative control of the white grubs Amphimallon majale and Popillia japonica. Biological Control. 51: 169-180. 97. Poinar, G. O., Jr., T. Jackson, and M. Klein. 1987. Heterorhabditis megidis sp. n. (Heterorhabditidae: Rhabditida) parasitic in the Japanese beetle, Popillia japonica (Scarabaeidae: Coleoptera), in Ohio. Proceedings of the Helminthological Society of Washington 54:53-59. 98. Power, K.T., An, R. and Grewal, P.S. 2009.  Effectiveness of Heterohabditis bacteriophora strain GPS11 applications targeted against different instars of the Japanese beetle Popillia japonica. Biological Control. 48: 232-236. 99. Yeh, T. and Alm, S.R. 1995. Evaluation of Steinernema glaseri (Nematoda: Steinernematidae) for biological control of Japanese and oriental beetles (Coleoptera, Searabaeidae). Journal of Economic Entomology. 88: 1251-1255. Leaf minors, Liriomyza trifolii and L. huidobrensis 100. Hara, A.H., Kaya, H.K., Gaugler, R., Lebeck, L.M. and Mello, C.L. 1993. Entomopathogenic nematodes for biological control of the leafminer, Liriomyza trifolii (Diptera: Agromyzidae).  Entomophaga 38, 359-369. 101. Head, J. and Walters, K.F.A. 2003.  Augmentation biological control utilising the entomopathogenic nematode, Steinernema feltiae, against the South American Leafminer, Liriomyza huidobrensis. Proceedings of the 1st International Symposium on Biological Control, (Hawaii, USA, 13-18 January 2002). USDA Forest Service, FHTET-03-05, 136-140. 102. Olthof, T.H.A. and Broadbent, A.B. 1992.  Evaluation of steinernematid nematodes for control of a leafminer, Liriomyza trifolii, in greenhouse chrysanthemums. Journal of Nematology 24, 612. 103. Stever, D.D. and Rice, B.   2004. A Comparison between two species of nematode (Steinernema carpocapsae and Steinernema feltiae) applied through a foliar spray to control leafminer (Liriomyza trifolii) in Dendranthema grandiflora. Hortscience 39: 846-846. 104. Tong-Xian Liu, Le Kang, K.M.Heinz, J.Trumble. 2008. Biological control of Liriomyza leafminers: progress and perspective. CAB Reviews: Perspectives in Agriculture, Veterinary Science, Nutrition and Natural Resources, 2009, 4, No. 004, 16 pp. 105. Williams, E.C. and MacDonald, O.C., 1995.  Critical factors required by the nematode Steinernema feltiae for the control of the leafminers Liriomyza huidobrensis, Liriomyza bryoniae and Chromatomyia syngenesiae.  Annals of Applied Biology. 127, 329-341. 106. Williams, E.C. and Walters, K.F.A. 1994. Nematode control of leafminers: Efficacy, temperature and timing. Brighton Crop Protection Conference – Pests and Disease. 1079-1084. 107. Williams, E.C. and Walters, K.F.A. 2000.  Foliar application of the entomopathogenic nematode Steinernema feltiae against leafminers on vegetables. Biocontrol Science and Technology 10, 61-70. Leopard moth, Zeuzera pyrina 108. Abdel-Kawy, A.G.M., El-Bishry, M.H. & El-Kifi, T.A.H. 1992: Controlling the leopard moth borer, Zeuzera pyrina by three entomopathogenic nematode species in the field. Faculty of Agriculture Bulletin of Univercity of Cairo, 43: 769-780. Mediterranean fruit fly Ceratitis capitata 109. Bazman, I., Ozer, N. and Hazir, S. 2008. Bionomics of the entomopathogenic nematode, Steinernema weiseri (Rhabditida: Steinernematidae). Nematology 10: 735-742. 110. Campos-Herrera, R. and Gutierrez, C. 2009. Screening Spanish isolates of steinernematid nematodes for use as biological control agents through laboratory and greenhouse microcosm studies. Journal of Invertebrate Pathology. 100: 100-105. 111. Karagoz, M., Gulcu, B., Hazir, C., Kaya, H.K. and Hazir, S. 2009. Biological control potential of Turkish entomopathogenic nematodes against the Mediterranean fruit fly Ceratitis capitata. Phytoparasitica. 37: 153-159. 112. Malan, A.P. and Manrakhan, A. 2009.  Susceptibility of the Mediterranean fruit fly (Ceratitis capitata) and the natal fruit fly (Ceratitis rosa) to entomopathogenic nematodes. Journal of Invertebrate Pathology. 100: 47-49. 113. Raja, R.K., Sivaramakrishnan, S. and Hazir, S. 2011. Ecological characterisation of Steinernema siamkayai (Rhabditida: Steinernematidae), a warm-adapted entomopathogenic nematode isolate from India. Biocontrol 56: 789-798. 114. Rohde, C., Moino, A., da Silva, M.A.T., Carvalho, F.D.and Ferreira, C.S. 2010. Influence of soil temperature and moisture on the infectivity of entomopathogenic nematodes (Rhabditida: Heterorhabditidae, Steinemematidae) against larvae of Ceratitis capitata (Wiedemann) (Diptera: Tephritidae). NeotropicaL Entomology. 39: 608-611. 115. Soliman, N. A. 2007. Efficacy of the entomopathogenic nematodes, Steinernema riobravis Cabanillas and Heterorhabditis bacteriophora (native strain) against the peach fruit fly, Bactrocera zonata (Saunders) and the Mediterranean fruit fly, Ceratitis capitata (Wiedemann). Egyptian Journal of Biological Pest control. 17: 77-82. 116. Soliman, N.A. 2007. Pathogenicity of three entomopathogenic nematodes to the Peach fruit fly, Bacterocera zonata (Saunders) and the Mediterranean fruit fly, Ceratitis capitata (Wiedemann) (Diptera : Tephritidae). Egyptian Journal of Biological Pest Control 17: 121-124. Mole crickets, Scapteriscus vicinus 117. Adjei, M.B., Smart, Jr. G.C., Frank, J.H. and Leppla, N.C. 2006. Control of pest mole crickets (Orthoptera: Gryllotalpidae) on pasture with the nematode Steinernema scapterisci (Rhabditida: Steinernematidae). Florida Entomologist 89: 532-535. 118. Barbara, K.A. and Buss, E.A. 2004. Survival and infectivity of Steinernema scapterisci (Nematoda: Steinernematidae) after contact with soil drench solutions. Florida Entomologist 87: 300-305. 119. Barbara, K.A. and Buss, E.A. 2005. Integration of insect parasitic nematodes (Rhabditida Steinernematidae) with insecticides for control of pest mole crickets (Orthoptera: Gryllotalpidae: Scapteriscus spp.). Journal of Economic Entomology 98: 689-693. 120. Barbara, K.A. and Buss, E.A. 2006. Augmentative applications of Steinernema scapterisci (Nematoda: Steinernematidae) for mole cricket (Orthoptera: Gryllotalpidae) control on golf courses. Florida Entomologist 89: 257-262. 121. Frank, J.H. 1999. Beneficial nematode killing mole crickets in Florida pastures, still spreading. Florida Turf Digest 16: 32, 34-35. 122. Frank, J.H., Grissom, C., Williams, C., Jennings. E., Lippi, C. and Zerba, E. 1999. A beneficial nematode is killing pest mole crickets in some Florida pastures and is spreading. Florida Cattleman 63: 31-32. 123. Georgis, R., Poinar Jr., G.O., 1994. Nematodes as biopesticides in turf and ornamentals. In: Leslie, A. (Ed.), Integrated Pest Management for Turf and Ornamentals. CRC Press, Boca Raton, FL, USA, pp. 477–489. 124. Nguyen, K.B. and Smart, G.C. 1990. Steinernema scapterisci n. sp. (Rhabditida: Steinernematidae). Journal of Nematology 22: 187-199. 125. Nguyen, K.B. and Smart, G.C. 1991. Mode of entry and site of development of Steinernema scapterisci in mole crickets. Journal of Nematology 23: 267-268. 126. Nguyen, K.B. and Smart, G.C. 1991. Pathogenicity of Steinernema scapterisci to selected invertebrates. Journal of Nematology 23: 7-11. 127. Nguyen, K.B. and Smart, G.C. 1991. Vertical distribution of Steinernema scapterisci. Journal of Nematology 23: 574-578. 128. Nguyen, K.B. and Smart, G.C. 1992. Addendum to the morphology of Steinernema scapterisci. Journal of Nematology 24: 478-481. 129. Nguyen, K.B. and Smart, G.C. 1992. Life cycle of Steinernema scapterisci Nguyen & Smart, 1990. Journal of Nematology 24: 160-169. 130. Parkman, J.P., Frank, J.H., Nguyen, K.B. and Smart, G.C. 1993. Dispersal of Steinernema scapterisci (Rhabditida: Steinernematidae) after inoculative applications for mole cricket (Orthoptera: Gryllotalpidae) control in pastures. Biological Control 3: 226-232. 131. Parkman, J.P., Frank, J.H., Nguyen, K.B. and Smart, G.C. 1994. Inoculative release of Steinernema scapterisci (Rhabditida: Steinernematidae) to suppress pest mole crickets (Orthoptera: Gryllotalpidae) on golf courses. Environmental Entomology 23: 1331-1337. Navel orangeworm, Amyelois transitella 132. Hodson, A.K., Siegel, J.P. and Lewis, E.E. 2012. Ecological influence of the entomopathogenic nematode, Steinernema carpocapsae, on pistachio orchard soil arthropods. Pedobiologia 55: 51-58. Peach borer, Synanthedon exitiosa 133. Cottrell, T.E. and Shapiro-Ilan, D.I. 2006. Susceptibility of the peachtree borer, Synanthedon exitiosa, to Steinernema carpocapsae and Steinernema riobrave in laboratory and field trials. Journal of Invertebrate Pathology 92: 85-88. 134. Shapiro-Ilan, D.I., Cottrell, T.E., Mizell, R.F., Horton, D.L. and Davis, J. 2009.  A novel approach to biological control with entomopathogenic nematodes: prophylactic control of the peachtree borer, Synanthedon exitiosa. Biological Control. 48: 259-263. Pecan weevil, Curculio caryae and C. hicoriae 135. Shapiro-Ilan, D. and Gardner, W.A. 2012. Improved control of Curculio caryae (Coleoptera: Curculionidae) through multi-stage pre-emergence applications of Steinernema carpocapsae. Journal of Entomological Science 47: 27-34. 136. Shapiro-Ilan, D. and Hall, M.J. 2012. Susceptibility of adult nut Curculio, Curculio hicoriae (Coleoptera: Curculionidae) to entomopathogenic nematodes under laboratory conditions. Journal of Entomological Science 47: 375-378. 137. Shapiro-Ilan, D.I., Cottrell, T.E. and Wood, B.W. 2011.  Effects of combining microbial and chemical insecticides on mortality of the Pecan weevil (Coleoptera: Curculionidae).  Journal of Economic Entomology 104: 14-20. 138. Shapiro-Ilan, D.I., Cottrell, T.E., Brown, I., Gardner, W.A., Hubbard, R.K. and Wood, B.W. 2006. Effect of soil moisture and a surfactant on entomopathogenic nematode suppression of the pecan weevil, Curculio caryae. Journal of Nematology 38: 474-482. 139. Shapiro-Ilan, D.I., Cottrell, T.E., Gardner, W.A., Leland, J. and Behles, R.W. 2009.  Laboratory mortality and mycosis of adult Curculio caryae (Coleoptera: Curculionidae) following application of Metarhizium anisopliae in the laboratory or field. Journal of Entomological Science. 44: 24-36. 140. Shapiro-Ilan, D.I., Dutcher, J.D. and Hatab, M. 2005.  Recycling potential and fitness of steinernematid nematodes cultured in Curculio caryae and Gaileria mellonella. Journal of Nematology 37: 12-17. 141. Shapiro-Ilan, D.I., Jackson, M., Reilly, C.C. and Hotchkiss, M.W. 2004.  Effects of combining an entomopathogenic fungi or bacterium with entomopathogenic nematodes on mortality of Curculio caryae (Coleoptera: Curculionidae). Biological Control 30: 119-126. 142. Shapiro-Ilan, D.I., Mizell, R.F., Cottrell, T.E. and Horton, D.L. 2008.  Control of plum curculio, Conotrachelus nenuphar, with entomopathogenic nematodes: Effects of application timing, alternate host plant, and nematode strain. Biological Control 44: 207-215. 143. Shapiro-Ilan, D.I., Stuart, R.J. and McCoy, C.W.  2005.  Characterization of biological control traits in the entomopathogenic nematode Heterorhabditis mexicana (MX4 strain).  Biological Control 32: 97-103. Pine weevil, Hylobius abietis 144. Dillon, A.B., Moore, C.P., Downes, M.J. and Griffin, C.T. 2008.  Evict or infect? Managing populations of the large pine weevil, Hylobius abietis, using a bottom-up and top-down approach. Forest Ecology and Management 255: 2634-2642. 145. Ennis, D.E., Dillon, A.B. and Griffin, C.T.  2010.  Pine weevils modulate defensive behaviour in response to parasites of differing virulence. Animal Behaviour 80: 283-288. 146. Everard, A., Griffin, C.T. and Dillon, A.B. 2009.  Competition and intraguild predation between the braconid parasitoid Bracon hylobii and the entomopathogenic nematode Heterorhabditis downesi, natural enemies of the large pine weevil, Hylobius abietis. Bulletin of Entomological Research. 99: 151-161. 147. Girling, R.D., Ennis, D., Dillon, A.B. and Griffin, C.T. 2010.  The lethal and sub-lethal consequences of entomopathogenic nematode infestation and exposure for adult pine weevils, Hylobius abietis (Coleoptera: Curculionidae). Journal of Invertebrate Pathology 104: 195-202. 148. Kruitbos, L.M., Heritage, S. and Wilson, M.J. 2009.  Phoretic dispersal of entomopathogenic nematodes by Hylobius abietis. Nematology. 11: 419-427. Plum weevil, Conotrachelus nenuphar 149. Alston, D.G., Rangel, D.E.N., Lacey, L.A., Golez, H.G., Kim, J.J. and Roberts, D.W. 2005.  Evaluation of novel fungal and nematode isolates for control of Conotrachelus nenuphar (Coleoptera: Curculionidae) larvae. Biological Control 35: 163-171. 150. Kim, H.G. and Alston, D.G. 2008. Potential of two entomopathogenic nematodes for suppression of plum Curculio (Conotrachelus nenuphar, Coleoptera: Curculionidae) life stages in northern climates.  Environmental Entomology 37: 1272-1279. 151. Pereault, R.J., Whalon, M.E. and Alston, D.G. 2009.  Field efficacy of entomopathogenic fungi and nematodes targeting caged last-instar plum Curculio (Coleoptera: Curculionidae) in Michigan cherry and apple Orchards. Environmental Entomology. 38: 1126-1134. 152. Shapiro-Ilan, D.I., Mizell, R.F. and Campbell, J.F. 2002. Susceptibility of the plum curculio, Conotrachelus nenuphar to entomopathogenic nematodes. Journal of Nematology 34: 246-249. 153. Shapiro-Ilan, D.I., Mizell, R.F., Cottrell, T.E and Horton, D.L. 2004. Measuring field efficacy of Steinernema feltiae and Steinernema riobrave for suppression of plum curculio, Conotrachelus nenuphar larvae. Biological Control 30: 496–503. 154. Shapiro-Ilan, D.I., Mizell, R.F., Cottrell, T.E., Horton, D.L. 2008. Control of plum curculio, Conotrachelus nenuphar with entomopathogenic nematodes: Effects of application timing, alternate host plant, and nematode strain. Biological Control. 44: 207-215. Shore flies, Scatella stagnalis 155.  Lindquist, R., Buxton, J. and Piatkowski, J. 1994.  Biological control of sciarid flies and shore flies in glasshouses. Brighton Crop Protection Conference, Pests and Diseases, BCPC Publications 3, 1067-1072. 156.  Morton, A. and Garcia del Pino, F. 2003. Potential of entomopathogenic nematodes for the control of shore flies (Scatella stagnalis). Growing Biocontrol Markets Challenge Research and Development. 9th European Meeting IOBC/WPRS Working Group “Insect Pathogens and Entomopathogenic Nematodes”, Abstracts, 67. 157.  Morton, A., Garcia del Pino, F., 2007.  Susceptibility of shore fly Scatella stagnalis to five entomopathogenic nematode strains in bioassays. Biocontrol 52: 533-545. 158.  Vanninen, I., Koskula, H. 2000. Biological control of the shore fly (Scatella tenuicosta) with steinernematid nematodes and Bacillus thuringiensis var. thuringiensis in peat and rockwool. Biocontrol Science and Technology 13: 47-63. Sod webworm, Herpetogramma phaeopteralis 159.  Nastaran Tofangsazie, N., Arthurs, S.P. and Cherry, R.  https://edis.ifas.ufl.edu/in968 Spruce webworm, Cephalcia abietis 160.  Fischer, P. 1996. The entomoparasitic nematode species Steinernema feltiae (Nematoda: Steinernematidae) as a mortality factor of the spruce webworm Cephalcia abietis (Hymenoptera: Pamphiliidae). Entomologia Generalis 21: 107-115. Stable fly, Stomoxys calcitrans 161.  Pierce, L. 2008. Efficacy of entomopathogenic nematodes used for control of stable flies (Stomoxys calcitrans) at round bale feeding sites in pastures. Abstract, Annual Meeting of Entomological Society of America. Stored grain pests 162.  Athanassiou, C.G., Kavallieratos, N.C., Menti, H. and Karanastasi, E. 2010. Mortality of four stored product pests in stored wheat when exposed to doses of three entomopathogenic nematodes. Journal of Economic Entomology 103: 977-984. 163.  Athanassiou, C.G., Palyvos, N.E. and 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. 164.  Fayyaz S. and Javed, S. 2009.  Laboratory evaluation of seven Pakistani strains of entomopathogenic nematodes against a stored grain insect pest, pulse beetle Callosobruchus chinensis (L.).  Journal of Nematology 41: 255-260. 165.  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. 166.  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. 167.  Ramos-Rodriguez, O., Campbell, J.F. and Ramaswamy, S.B. 2006. Pathogenicity of three species of entomopathogenic nematodes to some major stored-product insect pests. Journal of Stored Products Research 42: 241-252. 168.  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. Strawberry root borer, Nemocestes incomptus 169.  Georgis, R and Poinar, GO. 1984. Field control of the strawberry root weevil, Nemocestes-incomptus, by Neoaplectanid nematodes (Steinernematidae, Nematoda). Journal of Invertebrate Pathology 43: 130-131. Strawberry root weevil, Otiorhynchus ovatus and Ptiorhynchus ovatus 170. 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. 171. Booth, S.R., Tanigoshi, L.K. and 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. 172. 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. 173. 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. Strawberry crown moth, Synanthedon bibionipennis 174. 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. Tick, Rhipicephalus (Boophilus) microplus 175. de Carvalho, L.B., Furlong, J., Prata, M.C.D., dos Reis, E.S., Batista, E.S.D., Faza, A.P. and Leite R.C. 2010.  Evaluation in vitro of the infection times of engorged females of Rhipicephalus (Boophilus) microplus by the entomopathogenic nematode Steinernema glaseri CCA strain. Ciencia Rural. 40: 939-943. 176. 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. 177. 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. 178. 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. 179. 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. Western flower thrips, Frankliniella occidentalis 180. Arthurs, S. and Heinz, K.M.  2006. Evaluation of the nematodes Steinernema feltiae and Thripinema nicklewoodi as biological control agents of western flower thrips Frankliniella occidentalis infesting chrysanthemum. Biocontrol Science and Technology 16: 141-155. 181. Ebssa, L., Borgemeister, C. and Poehling, H.M. 2004. Effects of post-application irrigation and substrate moisture on the efficacy of entomopathogenic nematodes against western flower thrips, Frankliniella occidentalis. Entomologia Experimentalis et Applicata 112: 65-72. 182. Ebssa, L., Borgemeister, C. and Poehling, H.M. 2006. Simultaneous application of entomopathogenic nematodes and predatory mites to control western flower thrips Frankliniella occidentalis. Biological Control 39: 66-74. 183. Ebssa, L., Borgemeister, C., Berndt, O. and Poehling, H.M.  2001. Efficacy of entomopathogenic nematodes against soil-dwelling life stages of western flower thrips, Frankliniella occidentalis (Thysanoptera: Thripidae). Journal of Invertebrate Pathology 78: 119-127. 184. North, J.P., Cuthbertson, A.G.S. and Walters, K.F.A. 2006. The efficacy of two entomopathogenic biocontrol agents against adult Thrips palmi (Thysanoptera: Thripidae).  Journal of Invertebrate Pathology 92: 89-92. 185. Premachandra, D.W.T.S., Borgemeister, C., Berndt, O., Ehlers, R.U. and Poehling, H.M. 2003.  Laboratory bioassays of virulence of entomopathogenic nematodes against soil-inhabiting stages of Frankliniella occidentalis Pergande (Thysanoptera: Thripidae). Nematology 5: 539-547. 186. Trdan, S., Znidarcic, D. and Vidrih, M. 2007.  Control of Frankliniella occidentalis on glasshouse-grown cucumbers: an efficacy comparison of foliar application of Steinernema feltiae and spraying with abamectin.  Russian Journal of Nematology 15: 25-34. Western corn rootworm Diabrotica virgifera virgifera 187. Hiltpold, I., Hibbard, B.E., French, B.W. and Turlings, T.C.J. 2012. Capsules containing entomopathogenic nematodes as a Trojan horse approach to control the western corn rootworm. Plant and Soil 358: 10-24. 188. Petzold-Maxwell, J.L., Cibils-Stewart, X., French, B.W. and Gassmann, A.J. 2012.  Adaptation by western corn rootworm (Coleoptera: Chrysomelidae) to Bt Maize: inheritance, fitness costs, and feeding preference. Journal of Economic Entomology 105: 1407-1418. 189.  Petzold-Maxwell, J.L., Jaronski, T. and Gassmann, A.J.  2012.  Tritrophic interactions among Bt maize, an insect pest and entomopathogens: effects on development and survival of western corn rootworm. Annals of Applied Biology 160: 43-55. Whiteflies, Bemisia tabaci and Trialeurodes vaporariorum 190.  Cuthbertson, A.G.S., Blackburn, L.F., Eyre, D.P., Cannon, R.J.C., Miller, J. and Northing, P. 2011. Bemisia tabaci: The current situation in the UK and the prospect of developing strategies for eradication using entomopathogens. Insect Science 18: 1-10. 191. Cuthbertson, A.G.S., Mathers, J.J., Northing, P., Prickett, A.J. and Walters, K.F.A. 2008. The integrated use of chemical insecticides and the entomopathogenic nematode, Steinernema carpocapsae (Nematoda: Steinernematidae), for the control of sweetpotato whitefly, Bemisia tabaci (Hemiptera: Aleyrodidae). Insect Science 15: 447-453. 192. Head J., Lawrence A.J. and Walters K.F.A. 2004. Efficacy of the entomopathogenic nematode, Steinernema feltiae against Bemisia tabaci in relation to plant species, Journal of Applied Entomology. 128:543–547. 193.  Laznik, Z., Znidarcic, D. and Trdan, S. 2011. Control of Trialeurodes vaporariorum (Westwood) adults on glasshouse-grown cucumbers in four different growth substrates: an efficacy comparison of foliar application of Steinernema feltiae (Filipjev) and spraying with thiamethoxamn. Turkish Journal of Agriculture and Forestry 35: 631-640. 194. Shrestha, Y.K. and Lee, K.Y. 2012. Oral toxicity of Photorhabdus culture media on gene expression of the adult sweetpotato whitefly, Bemisia tabaci. Journal of Invertebrate Pathology 109: 91-96. White grubs (Summer Chafer), Amphimallon solstitiale 195. Glare T.R., Jackson T.A., Zimmermann G. 1993. Occurrence of Bacillus popilliae and two nematode pathogens in populations of Amphimallon solstitialis (Col.: Scarabaeidae) near Darmstadt, Germany.  Entomophaga. 38: 441-450. White grubs (Oriental beetle), Anomala orientalis, Exomala orientalis and Blitopertha orientalis 196. 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. 197. 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. 198. Grewal, P.S., Grewal, S.K., Malik, V.S. and Klein, M.G. 2007. Differences in susceptibility of introduced and native white grub species to entomopathogenic nematodes from various geographic localities. Biological Control 24: 230-237. 199. 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. 200. Koppenhofer, A.M. and Fuzy, E.M. 2003. Steinernema scarabaei for the control of white grubs. Biological Control 28: 47-59. 201. Koppenhofer, A.M. and Fuzy, E.M. 2006. Effect of soil type on infectivity and persistence of the entomopathogenic nematodes Steinernema scarabaei, Steinernema glaseri, Heterorhabditis zealandica, and Heterorhabditis bacteriophora. Journal of Invertebrate Pathology. 92: 11-22. 202. Koppenhofer, A.M. and Fuzy, E.M. 2008. Attraction of four entomopathogenic nematodes to four white grub species.  Journal of Invertebrate Pathology 99: 227-234. 203. Koppenhofer, A.M. and Fuzy, E.M. 2008. Earl timing and new combinations to increase the efficacy of neonicotinoid-entomopathogenic nematode (Rhabditida: Heterorhabditidae) combinations against white grubs (Coleoptera: Scarabaeidae.  Pest Management Science 64: 725-735. 204. Koppenhofer, A.M. and Fuzy, E.M. 2008c. Effect of the anthranilic diamide insecticide, chlorantraniliprole, on Heterorhabditis bacteriophora (Rhabditida: Heterorhabditidae) efficacy against white grubs (Coleoptera: Scarabaeldae). Biological Control. 45: 93-102. 205. 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. 206. Koppenhofer, A.M., Brown, I.M., Gaugler, R., Grewal, P.S., Kaya, H.K. and Klein M.G. 2000. Synergism of entomopathogenic nematodes and imidacloprid against white grubs: Greenhouse and field evaluation. Biological Control 19: 245-251. 207. 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. 208. 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. 209. 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. 210. Koppenhofer, A.M., Grewal, P.S. and Fuzy, E.M. 2007. Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species. Journal of Invertebrate Pathology 94: 184-195. 211. 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. 212. 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. 213. 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. 214. 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. 215. 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. 216. 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. White grubs, Costelytra zealandica 217. Kain, W.M. Bedding, R.A. and Vandermespel, C.J.  1982. Preliminary evaluations of parasitic nematodes for grass grub (Costelytra-zealandica (white)) control in central hawkes bay of new-Zealand. New Zealand Journal of Experimental Agriculture 10: 447-450. White grubs, Cotinus nitida 218. 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. 219. Townsend, M.L., Johnson, D.T. and Steinkraus, D.C. 1998. Laboratory studies of the interactions of environmental conditions on the susceptibility of green June beetle (Coleoptera: Scarabaeidae) grubs to entomopathogenic nematodes. Journal of Entomological Science 33: 40-48. 220. Townsend, M.L., Steinkraus, D.C. and Johnson, D.T. 1994. Mortality response of green June beetle (Coleoptera, Scarabaeidae) to 4 species of entomopathogenic nematodes. Journal of Entomological Science 29: 268-275. White grubs, Cyclocephala borealis, C. pasadenae and C. hirta 221. An, R. and Grewal, P.S. 2007. Differences in the virulence of Heterorhabditis bacteriophora and Steinernema scarabaei to three white grub species: The relative contribution of the nematodes and their symbiotic bacteria. Biological Control 43: 310-316. 222. Converse, V. and Grewal, P.S, 1998. Virulence of entomopathogenic nematodes to the western masked chafer Cyclocephala hirta (Coleoptera: Scarabaeidae). Journal of Economic Entomology 91: 428-432. 223. Koppenhofer, A.M. and Fuzy, E.M. 2008. Attraction of four entomopathogenic nematodes to four white grub species. Journal of Invertebrate Pathology 99: 227-234. 224. Koppenhofer, A.M. and Fuzy, E.M. 2008. Earl timing and new combinations to increase the efficacy of neonicotinoid-entomopathogenic nematode (Rhabditida: Heterorhabditidae) combinations against white grubs (Coleoptera: Scarabaeidae).  Pest Management Science 64: 725-735. 225. 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. 226. Koppenhofer, A.M., Choo, H.Y., Kaya, H.K., Lee, D.W. and Gelernter, W.D.  1999. Increased field and greenhouse efficacy against scarab grubs with a combination of an entomopathogenic nematode and Bacillus thuringiensis. Biological Control 14: 37-44 227. Koppenhofer, A.M., Grewal, P.S. and Fuzy, E.M. 2007. Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species. Journal of Invertebrate Pathology 94: 184-195. White grubs, Hoplia philanthus 228. Ansari, M.A., Adhikari, B.N. and Moens, M.  2008. Susceptibility of Hoplia philanthus (Coleoptera: Scarabaeidae) larvae and pupae to entomopathogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae). Biological Control 47: 315-321. 229. Ansari, M.A., Ali, F. and Moens, M.   2006. Compared virulence of the Belgian isolate of Steinernema glaseri (Rhabditida: Steinernematidae) and the type population of S-scarabaei to white grub species (Coleoptera: Scarabaeidae). Nematology 8: 787-791. 230. Ansari, M.A., Hussain, M. and Moens, M. 2009.  Formulation and application of entomopathogenic nematode-infected cadavers for control of Hoplia philanthus in turf grass. Pest Management Science. 65: 367-374. 231. Ansari, M.A., Shah, F.A., Tirry, L. and Moens, M. 2006.  Field trials against Hoplia philanthus (Coleoptera: Scarabaeidae) with a combination of an entomopathogenic nematode and the fungus Metarhizium anisopliae CLO 53. Biological Control 39: 453-459. 232. Ansari, M.A., Waeyenberge, L. and Moens, M. 2005.  First record of Steinernema glaseri Steiner, 1929 (Rhabditida: Steinernematidae) from Belgium: a natural pathogen of Hoplia philanthus (Coleoptera: Scarabaeidae). Nematology 7: 953-956. White grubs, Melolontha melolontha 233. Ansari, M.A., Ali, F. and Moens, M.   2006. Compared virulence of the Belgian isolate of Steinernema glaseri (Rhabditida: Steinernematidae) and the type population of S-scarabaei to white grub species (Coleoptera: Scarabaeidae). Nematology 8: 787-791. 234. Malinowski, H. 2011. Possibility of forest protection against insects damaging root systems with the use of biological method based on entomopathogenic nematodes and bacteria. Sylwan 155: 104-111. 235. Steiner, W.A. 1996. Dispersal and host-finding ability of entomopathogenic nematodes at low temperatures. Nematologica 42: 243-261. White grubs, Ataenius spretulus 236. 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. 237. 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. White grubs (Asiatic garden beetle), Maladera castanea 238. Koppenhofer, A.M. and Fuzy E.M. 2003. Biological and chemical control of the Asiatic garden beetle, Maladera castanea (Coleoptera: Scarabaeidae). Journal of Economic Entomology 96: 1076-1082. 239. 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. 240. 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. 241. 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. 242. 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. White grubs, Phyllophaga Spp. 243. Forschler, B.T. and Gardner, W.A.  1991.  Concentration mortality response of Phyllophaga-hirticula (Coleoptera, Scarabaeidae) to 3 entomogenous nematodes. Journal of Economic Entomology 84: 841-843. 244. Forschler, B.T. and Gardner, W.A.  1991. Parasitism of Phyllophaga-hirticula (coleoptera, scarabaeidae) by Heterorhabditis-heliothidis and Steinernema-carpocapsae.  Journal of Invertebrate Pathology 58: 396-407. 245. 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. 246. Koppenhofer, A.M., Rodriguez-Saona, C.R., Polavarapu, S. and Holdcraft, R.J. 2008. Entomopathogenic nematodes for control of Phyllophaga georgiana (Coleoptera: Scarabaeidae) in cranberries. Biocontrol Science and Technology 18: 21-31. 247. Liesch, P.J. and Williamson, R.C. 2010.  Evaluation of chemical controls and entomopathogenic nematodes for control of Phyllophaga white grubs in a Fraser Fir production field. Journal of Economic Entomology 103: 1979-1987. 248. Melo, E.L., Ortega, C.A., Gaigl, A. and Bellotti, A. 2010. Evaluation of entomopathogenic nematodes for the management of Phyllophaga bicolor (Coleoptera: Melolonthidae). Revista Colombiana de Entomologia 36: 207-212. 249. Melo-Molina, E.L., Ortega-Ojeda, C.A. and Gaigl, A. 2007. The effect of nematodes on larvae of Phyllophaga menetriesi and Anomala inconstans (Coleoptera: Melolonthidae).  Revista Colombiana de Entomologia 33: 21-26. 250. Nguyen, K.B., and Buss, E.A. 2011. Steinernema phyllophagae n. sp (Rhabditida: Steinernematidae), a new entomopathogenic nematode from Florida, USA. Nematology 13: 425-442. White grubs, Rhizotrogus majalis 251. An, R. and Grewal, P.S. 2007.  Differences in the virulence of Heterorhabditis bacteriophora and Steinernema scarabaei to three white grub species: The relative contribution of the nematodes and their symbiotic bacteria. Biological Control 43: 310-316. 252. An, R.S., Sreevatsan, S. and Grewal, P.S. 2009.  Comparative in vivo gene expression of the closely related bacteria Photorhabdus temperata and Xenorhabdus koppenhoeferi upon infection of the same insect host, Rhizotrogus majalis. BMC Genomics. 10: 433. 253. Koppenhofer, A.M. and Fuzy, E.M. 2008. Attraction of four entomopathogenic nematodes to four white grub species. Journal of Invertebrate Pathology 99: 227-234. 254. 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. 255. Koppenhofer, A.M., Grewal, P.S. and Fuzy, E.M. 2007. Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species. Journal of Invertebrate Pathology 94: 184-195. Fuller rose beetle, Asynonychus godmani 256. Morse, J.G. and Lindegren, J.E. 1996. Suppression of fuller rose beetle (Coleoptera: Curculionidae) on citrus with Steinernema carpocapsae (Rhabditida: Steinernematidae). Florida Entomologist 79: 373-384. Chive gnat, Bradysia odoriphaga 257. Ma, J., Chen, S.L., Moens, M., Han, R.C. and De Clercq, P. 2013. Efficacy of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) against the chive gnat, Bradysia odoriphaga. Journal of Pest Science 86: 551-561. 258. Sun, R. H., A. H. Li, R. C. Han, L. Cao, and X. L. Liu. 2004. Factors affecting the control of Bradysia odoriphaga with entomopathogenic nematode Heterorhabditis indica LN2. Natural Enemies of Insects 26:150–155.

    Nine important things about the damage caused by flea beetles and their control

    Interaction between flea beetles and entomopathogenic nematodes
    1. Flea beetles are called as flea beetles because they jump like fleas. Flea beetles are metallic black, blue, bronze or brown in color and about 1/16-1/8th inch long.
    2. Life cycle of flea beetles is very simple containing egg, larval and adult stages.  Flea beetles overwinter as adults by hiding under shelters including dry debris of plants (leaves and stems) left over from your garden crops or weeds. Early in the spring when temperature rises to about 50 F, the overwintering beetles become active and start feeding on the leaves of young plants. While feeding, they mate and lay eggs in the soil cracks around the root system of host plants or weeds in your garden or surrounding areas . Eggs hatch within 1-2 weeks and immediately larvae starts feeding on the roots of young host plants (see below) or weed hosts until they become mature. Then mature larvae pupate in the soil for 1-2 weeks; then emerge as adults and the life cycle continues. Generally this insect completes 2-3 generations in a year.
    3. Flea beetles are known to cause economic damage to many different vegetable crops including beans, broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, collards, corn, eggplant, kale, lettuce, melons, mustard,  peppers, potatoes, radishes, red Russian kale, rutabaga, spinach, squash, sunflowers, tomatoes, turnips and several species of weeds.
    4. Adults are the most damaging stage of flea beetles. They generally feed on foliage by chewing small holes through leaves and their heavy infestation gives a sieve-like appearance to the plant leaves thus reducing their marketable value especially leafy vegetables. Also, the heavy infestation of flea beetles can kill young seedlings.
    5. Flea beetle larvae feed on the plant roots but they do not cause a considerable economic damage to crop.
    6.  As temperature starts declining in the fall, adult flea beetles start looking for a shelter to hide and overwinter. Therefore, the process of management of flea beetles should begin in the fall to target overwintering beetles to reduce their incidence and outbreak in the next spring. The management of flea beetles should include both cultural and biological methods. Although the chemical insecticides could be more effective than other methods in controlling flea beetles, their use in the organic gardens should be avoided due to their detrimental effects on the human/animal health and environmental pollution.
    7. As a cultural control practice, keep your garden and its surrounding clean in the fall by removing all the plant debris including dry leaves and stems of harvested crops, weeds and other trash that may serve as the possible shelter for overwintering beetles.
    8.  Biological control method includes use of entomopathogenic nematodes (also called as insect-parasitic or beneficial nematodes) to target and kill larval and pupal stages of flea beetles in the spring.  Entomopathogenic nematodes can also attack and kill flea beetle adults if they come in contact in the soil.  Application of entomopathogenic nematodes such as Steinernema carpocapsae, Heterorhabditis bacteriophora and Heterorhabditis indica in the mid-late spring in your garden can kill both larval and pupal stages of flea beetles and thus reduce the emergence second generation adults, which are the most damaging to your crop.
    9. For the optimal rates and appropriate methods of application of entomopathogenic nematodes, read our blog at http://blog.bugsforgrowers.com/natural-predators/entomopathogenic-nematodes/beneficial-nematodes/how-to-deploy-your-nematode-army-and-kill-insect-pests/

    Target Japanese beetle larvae with entomopathogenic nematodes in the fall

    What are Japanese beetles?

    As name implies Japanese beetles, Popillia japonica are native to Japan but in the United States, they were first accidentally introduced into New Jersey in 1916. Currently, Japanese beetles have been distributed throughout the United State and causing economic loss to many agricultural and horticultural crops, and reducing aesthetic values of many ornamental plants. Japanese beetle adults are shiny and attractive metallic-green in color, oval shaped and about 1.5 inch long (Fig. 1.). These beetles cause a severe damage to leaves (Fig. 1), flowers (Fig.2) and ripening fruits of different plant species.  In case of severe infestation, adult Japanese beetles can completely skeletonize all the leaves (Fig. 3) and eventually defoliate the whole plants.  Larvae (also called grubs) of Japanese beetle make C- shape when they are disturbed (Fig. 4) and they possess three pairs of thoracic legs. They are whitish in color with yellowish-brown colored head capsule. Japanese beetle grubs generally feed on the roots of turf grass and many ornamental plants. The damage caused by Japanese beetle grubs to turf grass is easily recognized.   [caption id="attachment_483" align="aligncenter" width="179" caption="Fig.1. Japanese Beetles feeding on rose leaves"]"The Japanese beetles"[/caption] [caption id="attachment_485" align="aligncenter" width="179" caption="Fig. 2. Adult Japanese beetles are feeding on the rose flowers"]"The Japanese beetles feeding on roses"[/caption] [caption id="attachment_484" align="aligncenter" width="179" caption="Fig.3. Rose leaves are completely skeletonized by Japanese beetle adults"]"The severely skeletonized rose leaves"[/caption] [caption id="attachment_486" align="aligncenter" width="300" caption="Fig. 4. Japanese beetle larvae or grubs feed on the turfgrass roots."]"The Japanese beetle larvae or grub"[/caption]

    Signs of Japanese beetle infestation and damage to lawns and golf courses.

    • At the beginning of infestation in your lawn, you will notice localized patches of dead turf grass, which is always confused with the symptoms of water stress.
    • As the feeding activity of grubs on turf roots increases, small patches of dead turf are enlarged and joined together to form the large areas of dead turf.
    • This dead turf is generally loose and can be easily picked up with hand like a piece of carpet.
    • The most important sign of presence of Japanese beetle grubs in your lawn is that the infested areas of lawn is destroyed by digging animals such as raccoons and skunks or by birds that are looking for grubs to feast on them.

    Life cycle of Japanese beetle:

    For Japanese beetles, it takes about one year to complete egg to egg life cycle.  For example, adults of Japanese beetles emerge from pupae in the late June through July and start feeding on leaves, flowers and fruits. While feeding they mate and lay eggs in the soil near grass root zone at the depth of 1-2 inches. The eggs hatch within 1-2 weeks (i.e. in August) and first instar grub immediately starts feeding on grass roots and organic matter.  Grubs develop into two more instars August through October by continuously feeding on grass roots. In September and October they start moving deep into soil for overwintering.  When weather warms in April, grubs move back into the turf root-zone, start feeding on turf roots again and continue to develop and early in the June they pupate into the soil.  Then adults of Japanese beetles emerge from pupae in the late June, then they mate, lay eggs and life cycle continues.

    What are entomopathogenic nematodes?

    Entomopathogenic nematode are also called as insect-parasitic nematodes, which are defined as thread-like microscopic, colorless and un-segmented round worms. These round worms are the members of both Steinernematidae and Heterorhabditidae families and currently used as an excellent biological control agents against many soil dwelling insect pests of many economically important insect pests including Japanese beetles.  Entomopathogenic nematodes are sold when they are in the infective juvenile stage that also called as dauer juveniles. These infective juveniles always carry mutualistically associated symbiotic bacterial cells in their gut. Since these bacteria are pathogenic and capable of causing a disease to a variety of insect hosts, they are called as entomopathogenic nematodes.

    Which species of entomopathogenic nematodes are effective against Japanese beetles?

    Following species of entomopathogenic nematodes have been considered to be the most effective species against Japanese beetle grubs (see below for the optimum rates of nematodes).
    • Heterorhabditis bacteriophora nematodes
    • Heterorhabditis zealandica
    • Heterorhabditis indica nematodes
    • Steinernema scarabaei
    • Steinernema carpocapsae nematodes
    • Steinernema rivobrave

    Why fall is the time to apply nematodes and reduce existing populations to prevent future outbreaks of Japanese beetles.

    As we know that Japanese beetles overwinter in their larval stages. To do this, they will start moving deep into the soil in September and October (depending on the temperature). In some places the temperature has already started declining, which is an important cue for Japanese beetle larvae to get ready for winter weather.  Therefore, it is time to apply entompopathogenic nematodes which can target the Japanese beetle larvae that start going deep into the soil for overwintering.

    What stages of Japanese Beetles can be targeted?

    • All the immature stages of Japanese beetles are susceptible to entomopathogenic nematodes.
    • Adults of Japanese beetles are also susceptible to entomopathogenic nematodes.

    How can Entomopathogenic Nematodes kill Japanese beetle larvae?

    When the infective juveniles of entomopathogenic nematodes are applied to the soil surface or thatch layer, they start looking for their hosts including Japanese beetle grubs. Once a grub has been located, the nematode infective juveniles penetrate into the Japanese beetle grub body cavity via natural openings such as mouth, anus and spiracles. Then these infective juveniles enter grub’s body cavity where they release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in grub blood. When in the grub’s blood, multiplying nematode-bacterium complex causes septicemia and kills Japanese beetle grubs usually within 48 h after infection.  Nematodes generally feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new Japanese beetle grubs or other insect host that present in the soil.

    When, how and how many entomopathogenic nematodes should be applied for the effective control of Japanese beetles?

    For details read our blog

    Literature:

    Grewal, P.S., Koppenhofer, A.M., and Choo, H.Y., 2005.  Lawn, turfgrass and Pasture applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166. 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. Maneesakorn, P., An, R., Grewal, P.S.and Chandrapatya, A. 2010. Virulence of our new strains of entomopathogenic nematodes from Thailand against second instar larva of the Japanese Beetle, Popillia japonica (Coleoptera: Scarabaeidae). Thai Journal of Agricultural Science.43: 61-66. 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.

    Control fleas using entomopathogenic nematodes

    Fleas are one of the medically important pests of both animals and humans as they are capable of transmitting different kinds of disease causing organisms. Fleas are wingless insects but they can jump on their hosts including cats, dogs, humans and rats. Fleas have piercing and sucking type of mouthparts with that they suck blood of their hosts.  Like other insect, fleas also develop through the four different developmental stages including eggs, larva, pupa and adult.  Only adult fleas feed on blood but larval stage feeds on organic matter.  Pupa is a non-feeding stage. Fleas generally lay eggs on host’s body but they fall off on the ground where their host usually rests or sleeps.  Eggs hatch within 1-2 weeks and larvae immediately starts feeding on the organic matter that present at the resting place of animal hosts. Larvae develop through three larval stages and pupate in soil inside the silken cocoons. After 1-2 weeks, adult fleas emerge from cocoons but generally they use different kinds of host cues such as carbon dioxide, heat and vibration to emerge from pupae.  Fleas generally overwinter as larval and pupal stages, which can be easily targeted and killed by using biological control agents such as entomopathogenic nematodes.

    Why now it’s time to apply entomopathogenic nematodes and reduce the existing populations and future outbreaks of fleas.

    As we know that fleas overwinter as larval and pupal stages. In some places now temperature is already started declining, which is an important cue for fleas to get ready for winter weather.  This means both larval and pupal stages are ready for overwintering in the areas where temperatures are cooling down. Therefore, it is now time to apply entompopathogenic nematodes and target the overwintering stages of fleas.

    Which species of entomopathogenic nematodes are effective against fleas?

    • Steinernema carpocapsae nematodes are effective against fleas. It has been reported that when Steinernema carpocapsae nematodes applied in the potting medium, sand and gravel infested with larval and pupal stages of fleas, they reduced over 70% emergence of adults of cat fleas (Henderson et al., 1995).

    Where to apply entomopathogenic nematodes for the effective control of outdoor and indoor fleas?

    As stated above entomopathogenic nematodes can kill only larval and pupal stages but not adults of fleas. These stages are generally present on a large scale on the ground where host animals rests, sleeps or spends lot of time. These areas are generally located outdoors. These outdoor areas also serve as a source of indoor infestation of adult fleas. Therefore, it is important to treat all the outdoor animal resting/ sleeping areas or any other suspected areas where fleas are breeding with entomopathogenic nematodes.

    When to apply entomopathogenic nematodes for the effective control of outdoor and indoor fleas?

    • To target larvae and pupae of fleas, entomopathogenic nematodes should be applied starting from early spring through late fall i.e. when overwintering larval stages of fleas are becoming active and before emergence of adults from the pupae (spring and summer) or in the fall (September to November) when both larvae and pupae are getting ready for overwintering.
    • 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.
    • Another advantage of applying nematodes late in the evening is that the larval stages of fleas can be easily targeted because they are blind and do not like sunlight and therefore, they are generally active during night searching for food and easily found by entomopathogenic nematodes like Steinernema carpocapsae that uses sit and wait (ambush) strategy to attack its passing by host.  These nematodes can also find larvae and pupae that are hiding under organic matter during day time.

    How many entomopathogenic nematodes should be applied for the effective control of fleas?

    • See our Table for the exact quantity of Steinernema carpocapsae nematodes required to treat different square foot/meter areas.

    How to apply entomopathogenic nematodes?

    • Entomopathogenic nematodes that you receive in sponge as liquid formulation are thoroughly mixed in water and can be easily sprayed directly on the area where animal hosts rests/sleeps using traditional Knapsack/backpack sprayers or watering cans.
    • However, at time of spraying care should be taken that the nematodes should not be allowed to settle at the bottom of sprayer or watering can to avoid their uneven distribution.
    • To avoid the settling of nematodes at the bottom of sprayer or watering can, nematode suspension should be constantly agitated.
    • 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 86oF.
    • 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.
    • Nematodes received in granular formulation can be directly applied by broadcasting with hand or for larger area by using traditional spreaders that are used for application of granular or pallet pesticides or synthetic fertilizers.
    • Also, nematodes need about 20% moisture in the ground for survival. So please make sure nematode treated area should be watered immediately after the application of nematodes and continue to spray the area with water every few days.
    [caption id="attachment_80" align="aligncenter" width="300" caption="Watering can for application of entomopathogenic nematodes on a small area"]Entomopathogenic nematodes can be applied with a watering can[/caption]

    How Steinernema carpocapsae nematodes infect and kill fleas?

    Infective juveniles of Steinernema carpocapsae enter their insect host through natural openings such as mouth, anus and spiracles and eventually reach in the insect body cavity, which is filled with the blood that is technically called as hemolymph.  The infective juveniles of Steinernema spp. carry in their gut species specific symbiotic bacteria of the genus, Xenorhabdus. Once infective juveniles of Steinernema spp. are in the insect body cavity, they release several cells of symbiotic bacteria, Xenorhabdus spp. from their gut via anus in the insect blood. Insect blood is conducive for the multiplication of symbiotic bacteria. In the blood, multiplying nematode-bacterium complex causes septicemia and kill their insect host usually within 48 h after infection.

    Are entomopathogenic nematodes harmful to dogs, cats, chickens, birds, wild animals and humans?

    • Entomopathogenic nematodes are absolutely not harmful to humans and any pet animals (dogs, cats, chickens and birds) and wild animals/birds, and even to beneficial insects like honeybees.

     Literature:

    1. Henderson, G., Manweiler, S.A., Lawrence, W.J., Templeman, R.J. and Foil, L.D. 1995. The effects of Steinernema carpocapsae (Weiser) application to different life stages on adult emergence of the cat flea Ctenocephalides felis (Bouche). Vet. Dermatol. 6:159-163.
    2. Smith, C.A. 1995: Current concepts: Searching for safe methods of flea control. JAVMA: 1137-1143.