Bugs for Growers — 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"][/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 4. Described Steinernema species and their symbiotic bacterial species in the genus, Xenorhabdus.

Steinernema Species	Associated Xenorhabdus species
Neosteinernema longicurvicauda	Undescribed
Steinernema abbasi	Undescribed
S. aciari	Undescribed
S. affine	Xenorhabdus bovienii (Akhurst 1983) Akhurst and Boemare 1993
S. akhursti	Undescribed
S. anatoliense	Undescribed
S. apuliae	Undescribed
S. arenarium	X. kozodoii Tailliez, Pagès, Ginibre & Boemare, 2006
S. ashiuense	Undescribed
S. asiaticum	Undescribed
S. australe	X. magdalenensis Tailliez, Pages, Edgington, Tymo, and Buddie, 2012
S. backanense	Undescribed
S. beddingi	Undescribed
S. bicornutum	X. budapestensis Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
S. boemarei	Undescribed
S. brazilense	Undescribed
S. carpocapsae	X. nematophila (Poinar and Thomas 1965) Thomas and Poinar 1979
S. caudatum	Undescribed
S. ceratophorum	Undescribed
S. cholashanense	Undescribed
S. colombiense	Undescribed
S. costaricense	Undescribed
S. cubanum	X. poinarii (Akhurst 1983) Akhurst and Boemare 1993
S. cumgarense	Undescribed
S. diaprepesi	Undescribed
S. eapokense	Undescribed
S. ethiopiense	Undescribed
S. feltiae	X. bovienii (Akhurst 1983) Akhurst and Boemare 1993
S. glaseri	X. poinarii (Akhurst 1983) Akhurst and Boemare 1993
S. guangdongense	Undescribed
S. hebeiense	Undescribed
S. hermaphroditum	X. griffiniae Tailliez, Pagès, Ginibre & Boemare, 2006
S. ichnusae	Undescribed
S. intermedium	X. bovienii (Akhurst 1983) Akhurst and Boemare 1993
S. jollieti	Undescribed
S. karii	Undescribed
S. khoisanae	Undescribed
S. kraussei	X. bovienii (Akhurst 1983) Akhurst and Boemare 1993
S. kushidai	X. japonica Nishimura et al. 1995
S. leizhouense	Undescribed
S. litorale	Undescribed
S. loci	Undescribed
S. longicaudum	Undescribed
S. monticolum	Undescribed
S. neocurtillis	Undescribed
S. oregonense	Undescribed
S. pakistanense	Undescribed
S. phyllophagae	Undescribed
S. puertoricense	X. romanii Tailliez, Pagès, Ginibre & Boemare, 2006
S. puntauvense	Undescribed
S. rarum	X. szentirmaii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
S. riobrave	X. cabanillasii Tailliez, Pagès, Ginibre & Boemare, 2006
S. ritteri	Xenorhabdus sp
S. robustispiculum	Undescribed
S. sangi	Undescribed
S. sasonense	Undescribed
S. scapterisci	X. innexi Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
S. scarabaei	X. koppenhoeferi Tailliez, Pagès, Ginibre & Boemare, 2006
S. serratum	X. ehlersii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
S. siamkayai	X. stockiae Tailliez, Pagès, Ginibre & Boemare, 2006
S. sichuanense	X. bovienii (Akhurst 1983) Akhurst and Boemare 1993
S. silvaticum	Undescribed
S. tami	Xenorhabdus sp
S. texanum	Undescribed
S. thanhi	Undescribed
S. thermophilum	X. indica Somvanshi, Lang, Ganguly, Swiderski, Saxena, & Stackebrandt 2006
S. websteri	Undescribed
S. weiseri	Undescribed
S. xinbinense	Undescribed
S. xueshanense	Undescribed
S. yirgalemense	Undescribed
Steinernema sp	X. doucetiae Tailliez, Pagès, Ginibre & Boemare, 2006
Steinernema sp	X. hominickii Tailliez, Pagès, Ginibre & Boemare, 2006
Steinernema sp	X. mauleonii Tailliez, Pagès, Ginibre & Boemare, 2006
Steinernema sp	X. miraniensis Tailliez, Pagès, Ginibre & Boemare, 2006

 

Literature:

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(Rhabditida: Steinernematidae) From Cat Tien Forest, Vietnam. Russian Journal of Nematology 8:33-43. Ma, J., Chen, S., De Clercq, P., Waeyenberge, L., Han, R. and Moens, M. 2012. A new entomopathogenic nematode, Steinernema xinbinense n. sp. (Nematoda: Steinernematidae), from north China. Nematology 14:723-739. Mracek, Z. Qi-Zhi, L. and Nguyen, K.B. 2009. Steinernema xueshanense n. sp. (Rhabditida: Steinernematidae), a new species of entomopathogenic nematode from the province of Yunnan, southeast Tibetan Mts., China. Journal of Invertebrate Pathology 102: 69-78. Nguyen, K. B. and Smart, Jr., G.C.  1990. Steinernema scapterisci n. sp. (Steinernematidae: Nematoda). Journal of nematology 22:187-199. Nguyen, K. B. and Smart, Jr., G.C. 1992. Steinernema neocurtillis n. sp. (Rhabditida: Steinernematidae) and a key to species of the genus Steinernema. Journal of Nematology 24: Nguyen, K. B. and Smart, Jr., G.C. 1994. Neosteinernema longicurvicauda n. gen. n. sp. (Rhabditida: Steinernematidae), a parasite of the termite Reticulitermes flavipes (Koller). 1994. Journal of Nematology 26:162-174. Nguyen, K. B., and Duncan, L.W. 2002. Steinernema diaprepesi n. sp. (Rhabditida: Steinernematidae). a parasite of the root weevil Diaprepes abbreviatus (L) (Coleoptera: Curculionidae). Journal of Nematology 34:159-170. 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. Nguyen, K.B., Ginarte C. M.A, Leite, L.G., dos Santo, J.M. and Harakava, R. 2010. Steinernema brazilense n. sp. (Rhabditida: Steinernematidae) a new entomopathogenic nematode from Moto Grosso, Brazil. Journal of Invertebrate Pathology 103: 8-20. Nguyen, K.B., Malan, A.P. and Gozel, U. 2006. Steinernema khoisanae n. sp. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa. Nematology 8: 157-175. Nguyen, K.B., Puza, V. and Mracek, M. 2008. Steinernema cholashanense n. sp. (Rhabditida: Steinernematidae) a new species of entomopathogenic nematodes from the province of SichuanCholachan mountains, China. Journal of Invertebrate Pathology 97: 251-264. Nguyen, K.B., Qiu, L., Zhou, Y. and Pang, Y. 2006. Steinernema leizhouense sp. n.  (Nematoda: Steinernematidae), a new entomopathogenic nematode from southern China. Russian Journal of Nematology 14:101-118. Nguyen, K.B., Stuart, R.J, Andalo, V., Gozel, U. and Roger, M.E. 2007. Steinernema texanum n. sp. (Rhabditida: Steinernematidae) a new entomopathogenic nematode from Texas, USA. Nematology 9, 379-396. Nguyen, K.B., Tesfamariam, M., Gozel, U., Gaugler, R. and Adams, B.J. 2005. Steinernema yirgalemense n. sp. (Rhabditida: Steinernematidae) from Ethiopia. Nematology 6:839-856. Qiu, L, Fang Y.U., Zhou, Y., Pang, Y. and Nguyen K. B. 2004. Steinernema guangdongense sp. n.  (Nematoda: Steinernematidae), a new entomopathogenic nematode from southern China with a note on S. serratum (nomen nudum). Zootaxa 704:1-20. Qiu, L, Hu, X., Zhou, Y., Mei, S., Nguyen, K. B. and Pang, Y.  2005. Steinernema akhursti sp. n. (Nematoda: Steinernematidae) from Yunan, China. Journal of Invertebrate Pathology 90:151-160. Qiu, L, Hu, X., Zhou, Y., Pang, Y. and Nguyen, K. B. 2005. Steinernema beddingi n. sp. (Nematoda:Steinernematidae), a new entomopathogenic nematodes from Yunan, China. Nematology 7:737-749. Qiu, L., Yan, X., Nguyen, K.B. and Pang, Y. 2005. Steinernema aciari sp. n. (Nematoda: Steinernematidae), a new entomopathogenic nematode from Guangdong, China. Journal of Invertebrate Pathology 88:58-69. Somvanshi, V.S., Lang, E., Ganguly, S., Swiderski, J., Saxena, A.K. and Stackebrandt, E. 2006. A novel species of Xenorhabdus, family Enterobacteriaceae: Xenorhabdus indica sp. nov., symbiotically associated with entomopathogenic nematode Steinernema thermophilum Ganguly and Singh, 2000. Systemic and Applied Microbiology 29: 519-25. 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., Choo, H.Y. and Kaya, H.K. 1996. A new entomopathogenic nematode, Steinernema monticolum sp. n. (Rhabitida: Steinernematidae) from Korea. Nematologica 43: 15-29. Stock, S.P. Griffin, C.T. and Chaenari, R. 2004. Morphological and molecular characterization of Steinernema hermaphroditum n. sp. (Nematoda: Steinernematidae), an entomopathogenic nematode from Indonesia, and its phylogenetic relationship with other closely related taxa.  Nematology 6: 401-412. Stock, S.P. and Koppenhöfer, A.M. 2003. Steinernema scarabaei n. sp. (Rhabditida: Steinernematidae), a natural pathogen of scarab beetle larvae (Coleoptera: Scarabaeidae) from New Jersey. Nematology 5: 191-204. Stock, S.P., Samsook, V. and Reid, A. P. 1998. A new entomopathogenic nematode Steinernema siamkayai sp. n.  (Rhabditida: Steinernematidae) from Thailand.  Systematic Parasitology 41: 105-113. Tamiru, T., Waeyenberge, L., Hailu, T., Ehlers, R.-U., Půža, V., Mráček, Z. 2012.  Steinernema ethiopiense sp. n. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from Ethiopia. Nematology 14: 741- 757. Tarasco, E., Mracek, Z., Nguyen, K.B. and Trigiani, O. 2008. Steinernema ichnusae sp. n. (Nematode: Steinernematidae) a new entomopathogenic nemarode from Sardinia Island (Italy). Journal of Invertebrate Pathology 99: 173-185. Uribe-Lorio, L., Mora, M. and Stock, S. P. 2007. Steinernema costaricense n. sp. and Steinernema puntauvense n.sp. (Rhabditida, Steinernematidae), two new entomopathogenic nematodes from Costa Rica. Systematic Parasitology. 68: 167-172.  

Table 3. Described species of Heterorhabditis nematodes and their symbiotic bacteria species in the genus, Photorhabdus.

Heterorhabditis Species  Associated Photorhabdus species Heterorhabditis amazonensis Undescribed H. argentinensis Photorhabdus temperata H. atacamensis Undescribed H. bacteriophora  P. luminescens subspecies including. laumondii TT01,  kayaii, thracensis H. baujardi P. luminescens H. beicherriana Undescribed H. brevicaudis P. luminescens subsp. akhurstii H. downesi  Photorhabdus sp H. floridensis Undescribed H. georgiana P. luminescens subsp. akhurstii H. gerrardi P. asymbiotica  H. hambletoni Undescribed H. hawaiiensis P. luminescens H. heliothidis Undescribed H. hepialius P. luminescens H. hoptha  Undescribed H. indica P. luminescens H. marelata  P. luminescens H. megidis P. temperata subsp. temperata Xl Nach H. mexicana Undescribed H. noenieputensis Undescribed H. poinari  Photorhabdus sp H. safricana Undescribed H. sonorensis P. luminescens subsp. sonorensis H. taysearae Undescribed H. zealandica  P. temperata

 

Literature:

Andalo, V., Nguyen, K. B. and Moino, Jr., A. 2006. Heterorhabditis amazonensis n. sp. (Rhabditida: Heterorhabditidae) from Amazonas, Brazil. Nematology 8, 853-867. Edgington, S., Buddie, A. G., Moore, D., France, A., Merino, L. and Hunt, D. J. 2011. Heterorhabditis atacamensis n. sp (Nematoda: Heterorhabditidae), a new entomopathogenic nematode from the Atacama Desert, Chile. Journal of Helminthology 85: 381-394. Hsieh, F.C., Tzeng, C.Y., Tseng, J.T., Tsai, Y.S., Meng, M.H. and Kao, S.S. 2009.  Isolation and Characterization of the Native Entomopathogenic Nematode, Heterorhabditis brevicaudis, and its Symbiotic Bacteria from Taiwan.  Current Microbiology. 58: 564-570. Li, X.Y., Liu, Q.Z., Nermut, J., Puza, V. and Mracek, Z. 2012. Heterorhabditis beicherriana n. sp (Nematoda: Heterorhabditidae), a new entomopathogenic nematode from the Shunyi district of Beijing, China. Zootaxa  Issue: 3569: 25-40. Malan, A.P., Knoetze, R. and Tiedt, L. 2012. Heterorhabditis noenieputensis n. sp. (Rhabditida: Heterorhabditidae), a new entomopathogenic nematode from South Africa. Journal of Helminthology 12:1-13. Malan, A.P., Nguyen, K.B., de Waal, J.Y. and Tiedt, L. 2008. Heterorhabditis safricana n. sp (Rhabditida : Heterorhabditidae), a new entomopathogenic nematode from South Africa. Nematology 10: 381-396. Nguyen, K.B., Gozel, U., Koppenhofer, H. S. and Adams, B. J. 2006. Heterorhabditis floridensis n. sp. (Rhabditida: Heterorhabditidae) from Florida. Zootaxa 1177: 1-19. Nguyen, K.B., Shapiro-Ilan, D. and Mbata, G. 2008. Heterorhabditis georgiana n. sp. (Rhabditida: Steinernematidae) from Georgia, USA. Nematology10, 433-448. Nguyen, K.B., Shapiro-Ilan, D. I., Stuart, R.J., MCCoy, C.W., James, R.R. and Adams, B.J. 2004. Heterorhabditis mexicana n. sp. (Heterorhabditidae: Rhabditida) from Tamaulipas, Mexico with morphological studies of bursa of Heterorhabditis spp. Nematology 6:231-244. Orozco, R.A., Hill, T. and Stock, S.P. 2013.  Characterization and phylogenetic relationships of Photorhabdus luminescens subsp. sonorensis (gamma-Proteobacteria: Enterobacteriaceae), the bacterial symbiont of the entomopathogenic nematode Heterorhabditis sonorensis (Nematoda: Heterorhabditidae). Current Microbiology 66: 30-39. Phan, K.L., Subbotin, S.A., Nguyen, N.C. and Moens, M. 2003. Heterorhabditis baujardi sp n. (Rhabditida : Heterorhabditidae) from Vietnam and morphometric data for H-indica populations.  Nematology 5: 367-382. Plichta, K.L., Joyce, S.A., Clarke, D., Waterfield, N. and Stock, S.P. 2009.  Heterorhabditis gerrardi n. sp (Nematoda: Heterorhabditidae): the hidden host of Photorhabdus asymbiotica (Enterobacteriaceae: gamma-Proteobacteria). Journal of Helminthology.83: 309-320. Poinar, G.O. 1975. Description and biology of a new insect parasitic rhabditoid, Heterorhabditis-bacteriophora n-gen, n-sp (rhabditida, heterorhabditidae n-fam). Nematologica 21: 463-470. 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. 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., Rivera-Orduno, B. and Flores-Lara, Y. 2009. Heterorhabditis sonorensis n. sp (Nematoda: Heterorhabditidae), a natural pathogen of the seasonal cicada Diceroprocta ornea (Walker) (Homoptera: Cicadidae) in the Sonoran desert. Journal of Invertebrate Pathology 100: 175-184.

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. 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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. 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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. 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Biological and Cultural methods to control imported cabbageworms

The cabbage butterfly is commonly called as imported cabbageworm, Artogeia rapae (Pieris rapae). This is one of the most important pests of many Cole crops including broccoli, cabbage, collard greens, cauliflower, kale and turnip.  Butterflies are easy to identify as they have whitish colored fore-and hind-wings with one and two black spots on the top of each of fore wings of males and females, respectively.  Also, both males and females have a black spot on the outer front margin of each hind wing.  Females lay singly yellow colored and oblong eggs on the either side of the leaves and depending on the temperature eggs hatch within a week. Mature larvae are velvety green in color with a narrow orange stripe down the middle of the back and a yellowish stripe along each side of the body (Fig. 1.)  The pupae are green to light brown in color, attached to bottom leaves and adults generally emerge from these pupae within 2 weeks of pupation. Cabbage butterflies overwinter as pupae in previous crop plant debris in the garden. [caption id="attachment_644" align="aligncenter" width="300" caption="Fig. 1. Severe damage caused by Imported Cabbageworms near growing point of a collard green plant"][/caption] Generally cabbage butterfly larvae feed voraciously near to the growing point of the host plants (Fig. 2) but they can also feed indiscriminately by chewing large irregular holes on both young and mature leaves of different host plants including broccoli, cabbage, cauliflower, collard greens, kale and turnip (Figs. 1 and 2). [caption id="attachment_645" align="aligncenter" width="200" caption="Fig. 2. Larva of an imported cabbageworm feeding by chewing large irregular holes on a collard green mature leaf"][/caption] Since chemical insecticides cannot be used in organic vegetable gardens, growers have to rely on the cultural and biological methods to manage populations of imported cabbageworm.

Cultural Methods

For small or large vegetable gardens, best cultural practice is hand picking and killing of all the larval stages of imported cabbageworm. Although this practice is laborious and time consuming, it works and reduces the damage caused by this economically important insect pest. Also, at the end of the fall season remove all the previous crop plant debris so that there will be less protected areas available for overwintering imported cabbage worms, which in turn will reduce the populations of adults in the next spring.  This low number of adult emergence means there will be less numbers of eggs to hatch into larvae meaning there will be less larval incidence to cause the damage to the crop in the spring.

Biological Methods

Biological methods include use of natural enemies/biological control agents to control cabbage butterflies. Three well known biological control agents including Bacillus thuringiensis (Bt), entomopathogenic nematodes and wasps have a potential to manage imported cabbageworm population in the vegetable gardens.

Bacillus thuringiensis kurstaki (Bt):

This bacterium is recognized as a bacterial insecticide but it is not harmful to the humans, animals or the environment. This is a very effective biopesticide on young larval stages as compared to the mature larval stages of cabbage butterflies.  This microbial biocontrol agent is commercially available and can be applied using traditional sprayers. For the effective control of imported cabbageworms, Bacillus thuringiensis kurstaki should be applied at every seven day interval after noticing the first incidence of pest.

Entomopathogenic nematodes:

Currently, entomopathogenic nematodes are used as effective biological control agents against many different kinds of soil-dwelling insect pests of many economically important crops and turfgrasses. These nematodes are commercially available and are not harmful to humans, animals and even beneficial insects like honeybees. Canadian researchers have demonstrated that the entomopathogenic nematodes including Steinernema carpocapsae, S. feltiae and S. riobrave can cause 76 to 100% mortality of imported cabbageworms Artogeia rapae if applied at temperatures ranging from 25 to 30 °C and their LC50 values were ranged from 4 to 18 infective juveniles (Bélair et al., 2003). Mahar et al (2005) also reported that in addition to the above stated species of entomopathogenic nematodes, Heterorhabditis bacteriophora and H. indica nematodes can infect and kill both larvae and pupae of cabbage butterflies. Recently, another insect-parasitic nematode, Rhabditis blumi also been shown to be effective against imported cabbageworm (Park et al., 2012).

Wasps:

Following four species of parasitic wasps can serves as effective biological control agents against imported cabbage worm.

1. The egg parasitic wasp, Trichogramma spp.: This is a very tiny parasitic wasp known for parasitizing eggs of imported cabbageworms. These wasps are commercially available and can be mass released when lots of adult butterflies are present in the garden and already started laying eggs on the leaves.  This will prevent the hatching of eggs into larvae thus preventing damage caused by imported cabbageworm larvae to Cole crops (Oatman et al., 1968).
2. The brachonid wasp, Cotesia glomerata: This gregarious wasp parasitizes the larvae of the imported cabbageworms. This wasp species is not commercially available but it can naturally occur (Herlihy et al., 2012) and capable of suppressing the populations of cabbage butterflies in the vegetable gardens and fields. This wasp lays eggs inside the young caterpillars of imported cabbageworms. The eggs hatch and the larvae develop inside the developing imported cabbageworm larvae, then emerge as mature larvae and pupate in yellow silken cocoons outside the host, which dies during the process of the emergence of wasp larvae. If this wasp is present in the fields, which are infested with imported cabbageworms or other insect hosts, it can parasitize and kill over 60% of their insect host larvae.
3. The solitary wasp, Cotesia rubecula:  This naturally occurring parasitic wasp is known to its specificity to the members of genus Pieris especially imported cabbageworms. Although C. rubecula wasp parasitizes all the stages of imported cabbageworms, it prefers last instar of imported cabbageworms, which is the most damaging stage. This is the most studied parasitic wasp of imported cabbageworms and found to be distributed throughout the US (Herlihy et al., 2012).
4. The pteromalid wasp, Pteromalus puparum: This tiny wasp specifically parasitizes pupae of imported cabbageworms and other lepidopterous insects.  Since this wasp parasitoid kills only pupae of its insect host, it does not reduce the larval feeding damage caused before pupation but it certainly reduces the emergence of the next generation of adults. This means there are less number of egg laying females that results in the less number of eggs and therefore, less larval incidence to cause severe damage to the crop.

 

Literature:

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