Bugs for Growers — Insect Pests

Now is the right time to use ladybugs to control aphids, mealybugs, mites, scales and thrips- Bugsforgrowers.com

Watch YouTube video about using ladybugs for the control of aphids: "

https://youtu.be/UH9jPIUEheo

 Description: Ladybird beetles are also called as ladybugs and scientifically known as Hypodamia convergens. Both larvae and adults of ladybugs are active predators of many soft bodied insects including aphids, mealy bugs, mites and scale insects. Adult ladybugs are hemispherical shaped and bright orange to red in color with 6 black spots on each wing (Fig. 1) whereas larvae have appearance like an alligator with yellow stripes and reddish dark brown color (Fig. 2).  Since adults of ladybugs are very active predators, they can move very fast after their release and find their prey in the garden. Each adult of ladybug is capable of eating over 5000 aphids whereas each larva in its life-span can eat over 400 aphids (Fig. 3).  Use of ladybugs as biological control agents in the organic gardens is a good choice because released ladybug adults can continuously reproduce and maintain their populations as long as they find their food in the garden.  For example, once ladybugs are released in the garden, they immediately start feeding on their prey.  While feeding they mate and lay up to 1,500 tiny yellow eggs on foliage. Eggs hatch within a week into blackish brown alligator like larvae, which are very mobile. After hatching, these larvae immediately start feeding on the available insect hosts in the garden.  While feeding, larvae molts 3 times before pupation.  The pupae are orange to black in color and attached to any substrate in the garden. Young adults emerge from pupae within 1-2 weeks and life cycle continues.  Ladybugs generally complete several generations in a year and they hibernate as adults during winter. [caption id="attachment_1288" align="aligncenter" width="201" caption="Fig. 1. An adult ladybug"][/caption]   [caption id="attachment_1299" align="aligncenter" width="300" caption="Fig.2. A ladybug larva was feeding on okra aphids"][/caption]   [caption id="attachment_1300" align="aligncenter" width="300" caption="Fig. 3. Red aphids feeding on underside of a tomato leaf"][/caption]  

Ladybugs are effective against following insect pests

• Aphids
• Mealy bugs
• Mites
• Scale insects
• Thrips

How ladybugs are applied in the organic garden?

Ladybugs are commercially available and they sold about 1500 adults in a bag and one such bag is enough to treat about 900 square foot area of garden. After purchasing, if it is possible, release adults immediately at dusk in watered garden near plants infested with aphids or other host insect pests. If it is not possible to release ladybugs immediately after arrival, they can be stored in the refrigerator at 38^o F (4^oC) until you are ready to release them in the garden. However, for their highest survival rate in the refrigerator, it is recommended that the bags containing lady bird beetles should be pre-conditioned upon arrival by rinsing bags under cold water and then transfer them into the refrigerator at 4^oC (38^o F).

Are ladybugs safe to use as biological control agent?

Yes, as they are not harmful to humans, children and pets.

Research Papers: Dreistadt, S.H. and Flint, M.L 1996. Melon aphid (Homoptera: Aphididae) control by inundative convergent lady beetle (Coleoptera: Coccinellidae) release on Chrysanthemum. Environmental Entomology 25:688-697. Eigenbrode, S.D., White, C., Rohde, M. and Simon, C.J. 1998. Behavior and effectiveness of adult Hippodamia convergens (Coleoptera: Coccinellidae) as a predator of Acyrthosiphon pisum (Homoptera: Aphididae) on a Wax Mutant of Pisum sativum. Environmental Entomology 27: 902-909.

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"][/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."][/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"][/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"][/caption] [caption id="attachment_691" align="aligncenter" width="300" caption="Fig. 5. Red colored 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"][/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 30^oC (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."][/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."][/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 28^oC (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."][/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"][/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"][/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.

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) armigera, Spodoptera exigua, S. frugiperda	H. amazonensis, H. indica S. arenarium, S. carpocapsae, S. glaseri	14-18
Billbugs, Sphenophorus purvulus, S. levis	H. bacteriophora, S. brazilense, S. carpocapsae	19-20
Black vine weevil, Otiorhynchus salcatus	H. bacteriophora, H. downesi, H. megidi.S. carpocapsae, S. 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. carpocapsae, S. 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 fly, Ceratitis capitata	H. bacteriophora, H. zealandica, S. carpocapsae, S. feltiae, S. khoisanae, S. siamkayai, S. weiseri	109-116
Mole crickets, Scapteriscus 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. bacteriophora, H. indica, S. arenarium, S. 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. bacteriophora, H. megidis, H. zealandica, S. carpocapsae, S. 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. bacteriophora, H. indica, H. marelata, H. megidis, H. 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. bacteriophora, H. marelata, H. megidis, S. arenaria, S. feltiae, S. glaseri, S. riobrave	233-235
White grub, Ataenius spretulus	H. bacteriophora, S. glaseri, S. scarabae	236-237
White grub (Asiatic garden beetle), Maladera castanea	H. bacteriophora, S. 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. bacteriophora, H. megidis, H. zealandica, S. carpocapsae, S. 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 tumida. Journal 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. 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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. 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