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Eight weevils can be controlled with steinernema carpocapsae nematodes

Jan 31

What are weevils?

Like other beetles, weevils belong to an insect order Coleoptera but unlike other beetles, they have a distinct narrow beak or rostrum extending in front of their heads with chewing mouthparts are located at the tip of rostrum. Several species of weevils that have been described so far are considered as the most serious pests of many plant species including field and fruit crops, ornamentals, forest trees and turf grasses. The common names that are assigned to different species of weevils are usually associated with the common names of their host plants (see below). Also, the sizes and colors of both grubs also called as larvae (Photo 1) and adults (Photo 2) generally differ with their species. Following eight species of weevils can be controlled with beneficial entomopathogenic nematode, Steinernema carpocapsae nematodes.   [caption id="attachment_2123" align="aligncenter" width="482"]Weevil Grub Photo 1. Larva of Weevil[/caption] [caption id="attachment_2124" align="aligncenter" width="482"]Weevil adult Photo 2. An adult weevil[/caption]

Alfalfa weevil, Hypera postica

Adult weevils are light brown in color with a broad darker stripe extending down their midline. Young larvae of alfalfa weevils are tiny and yellowish green with black heads whereas their older larvae are greenish in color with black heads and a distinct white line down the center of their backs and white lines along each side of body. Although both adults and larvae of alfalfa weevils cause damage to alfalfa leaves, the damage caused by larvae is more severe than the damage caused by adults. In case of sever infestation, larvae can completely defoliate the plants and reduce the yields. Although all the stages including eggs, larvae, pupae and adults of alfalfa weevil are completed on the plants, sometime their mature larvae can pupate in the leaves that are fallen to the ground. So these mature larvae and pupae that are fallen to the ground can be targeted with beneficial entomopathogenic Steinernema carpocapsae nematodes.

Annual bluegrass weevil, Listronotus maculicollis

The annual bluegrass weevil prefers to feed mainly on annual bluegrass (Poa annua) compared to any other grass species and therefore they are named as annual bluegrass weevils. The larvae of blue grass weevil are creamy white and legless whereas adults are black colored with small hairs and gold scales on their forewings (elytra). Although both adults and larvae (grubs) of annual bluegrass weevil cause damage to turf grass, their larvae cause the most severe damage to annual bluegrass. Overwintering adults become active early in the spring and start feeding on the grass leaves and mate. The eggs laid by female weevils hatch early in the May into young larvae that start feeding by tunneling into grass stems. The symptoms of damage caused by bluegrass weevil include yellowing of grass leaves and subsequently dying of grass.  Matured grubs generally move out of the tunnels and begin feeding externally on the surface of stems, crowns and roots of grass. In case of severe infestation, they can cut off stems from roots causing complete drying and loss of turf grass. Then mature larvae move in the soil for pupation. Both mature larvae and pupae can be easily targeted with Steinernema carpocapsae nematodes.

Billbugs, Sphenophorous parvulus, S. cicastriatus

The billbugs are weevils but they are commonly called as snout beetles. Although billbugs prefer Kentucky bluegrass, they also feed on perennial ryegrass, tall fescue, corn, rye and wheat. Adult weevils are blackish or greyish in color with elbowed antennae and distinct grooves running longitudinally down their forewings. Billbug grubs are legless, cream in color with brown head capsules and slightly curved body. Overwintering billbug adults emerge out from their hiding places in April and start feeding on the grass leaves. Female billbugs then lay eggs inside a hole chewed by them in the stem near the base of the turf plant. After hatching from eggs, young larvae generally move towards crown and roots by burrowing through the grass stem. These grubs will then feed voraciously and completely damage the crowns and eventually kill the entire grass plant. Damage by billbugs generally starts appearing as irregular mottling or thinning patches of the turf in mid-June through August. As both larval and pupal stages of billbugs found in the soil, they can be easily killed with Steinernema carpocapsae nematodes.

Black vine weevil, Otiorhynchus sulcatus

The black vine weevils are considered as one of the most damaging insect pests of many nursery and landscape plants. Adult weevils are black or gray in color with elbowed antennae and a prominent short snout. Fully-grown grubs are C-shaped, legless and whitish in color with brown head capsules. Although both adults and larvae of black vine weevil cause direct damage to host plants, adults mostly feed on foliage and flowers whereas larvae (grubs) feed on the roots. Feeding damage caused by adults can be easily recognized because of a typical notched appearance of damaged leaves or flowers. Adult weevils also feed on the developing young buds and shoots. Larvae of black vine weevils cause a serious damage to roots of different host plants. Heavy infestation of black vine weevil can kill the entire plant. Since all the larval stages and pupae live in the soil, they can be easily found and killed by entomopathogenic nematodes like Steinernema carpocapsae nematodes.

Red palm weevil, Rhynchophorus ferrugineus

The red palm weevil is considered as one of the major pest of palms in the Mediterranean Basin but they can be considered as a serious threat to date production in California. Adult weevils are reddish brown in color with a long curved rostrum (snout) and dark spots on the upper side of the middle part of the body whereas their larvae are white in color with a brown head capsule. Damage to palms is mainly caused by the larvae and only becomes visible long after infestation. By the time the first symptoms of the pest infestation are noticed, the palm plants are already so seriously damaged that they cannot survive. As mature larvae of red palm weevil move in the leaf litter at the base of tree trunk for pupation, both larvae and pupae can be targeted and killed with Steinernema carpocapsae nematodes.

Strawberry root weevils, Otiorhynchus ovatus

The strawberry root weevils are one of the most serious insect pests of strawberry crop.  Adult weevils are reddish brown to black in color whereas their larvae are creamy white in color, C-shaped, legless with tanned heads. Adults of strawberry root weevil feed on the edges of strawberry leaves (notching of leaves) whereas all of their larval stages cause feeding damage by tunneling in the roots and crowns. The main symptoms of damage caused by larvae of strawberry weevils include pruning of roots, weakening, stunting of plant growth and eventually killing of entire plants. As both larvae and pupae of Strawberry root weevils, Otiorhynchus ovatus live in the soil; they are easy target for entomopathogenic nematodes.

Citrus root weevil, Diaprepes abbreviates

The citrus root weevil is one of the major insect pests of citrus and many ornamental plants in California and Florida. Adults of citrus root weevil are black in color with small white, red, orange or yellow colored scales on the forewings (elytra). The larvae (grubs) are white and legless with light head capsule. Adults cause leaf notching by feeding on the margins/ edges of leaves. The larvae of citrus weevil mainly feed on the roots of their hosts. In the severe cases they can girdle main roots that can result into death of citrus trees. As all the larval stages and pupae of the citrus root weevil live in the soil, they can be easily controlled with the application of entomopathogenic nematodes like Steinernema carpocapsae.

Pecan weevil, Curculio caryae

The pecan weevil is considered as one of the most devastating pest of pecans. The adults of pecan weevils are brown to grey in color with a characteristic snout, as its length is equal to the length of weevil’s body. Larvae of the pecan weevil are creamy white in color with reddish head capsule. The adults of the pecan weevil will emerge from pupae in the soil, then they will move out of the soil into the tree canopy either by crawling on the tree trunk or directly flying in August through September. Once in the canopy, adult weevils begin feeding by puncturing young developing nuts that will fall off the trees prematurely within 2-3 days. While feeding adult weevils mate. The mated females then lay eggs inside the nuts by chewing a hole into its hardened shell. After hatching from eggs inside the nut, larvae start feeding immediately on a developing kernel and mature within 2-3 weeks. When the damaged nuts fall to the ground, the mature larvae exit the nuts by cutting a small circular hole in the shell and then burrow into the soil and pupate. Three weeks after pupation, adult weevils emerge in August. It takes about 2-3 years for pecan weevils to complete their life cycle. Since all the mature larval, pupal and adult stages of pecan weevil live in the soil under pecan tree, they can be easily targeted by pre-emergence applications of beneficial entomopathogenic nematodes in early May, June and in late June or in early July.

Biological control of weevils with Steinernema carpocapsae nematodes

Because of cryptic habitats of weevils, their management with chemical insecticides is difficult. During completion of entire life cycle at least one stage (either adult, larva or pupa) of almost all the above stated species of weevils lives in the soil and this soil dwelling stage can be easily targeted and killed with the inundative application of beneficial entomopathogenic nematodes. Entomopathogenic nematodes including Steinernema carpocapsae have been proved to be the best biological control agents to manage weevils that infesting different crops. It has been demonstrated that S. carpocapsae nematodes when applied at the rate of 23000 nematodes per square foot or 1billion nematodes per acre, they can cause between 50 and 100% mortality of many weevil species including alfalfa weevils, annual bluegrass weevils, black vine weevils, citrus root weevils, large pine weevils, pecan weevils, red palm weevils and strawberry root weevils (Abbas, et al., 2001; Ansari and Butt, 2011; Dembilio et al., 2010; Duncan et al., 1996; Lacer et al., 2009; McGraw et al., 2010; Shah et al., 2011; Schroeder, 1992, Simser and Roberts, 1994).

How does Steinernema carpocapsae Nematode Kill Weevils?

Beneficial Steinernema carpocapsae nematodes are parasites of insects and most effective against soil-dwelling stages of many insect pests including weevils at temperatures ranging from 22 to 28°C (72 - 82°F).  Infective juveniles of Steinernema carpocapsae nematode use an "ambush” foraging strategy to find their insect hosts. In this strategy, the infective juveniles of this nematode (Photo 3) “sit-and-wait” to attack passing by and highly mobile larvae/ grubs of insects including weevils. Also, infective juveniles of this nematode can move in the soil and insect host larvae. These nematodes always carry symbiotic bacteria called Xenorhabdus nematophila in their gut and use them as a weapon to kill their insect hosts and as food for their development and reproduction inside the host cadavers. When the infective juveniles S. carpocapsae are applied either to the surface of the soil in the field, thatch layer on golf courses or plant growing medium in the pots, they begin searching for their hosts including weevil grubs and their pupae. Once they locate a grub and/or pupa, the infective juveniles will penetrate into the body cavity of grubs or pupae via natural openings like mouth, anus and spiracles. Once in the body cavity, infective juveniles will release symbiotic bacteria, Xenorhabdus nematophila from their gut in the blood of grubs or pupae.  Multiplying nematode-bacterium complex in the blood causes septicemia and kills the grub usually within 48 h after infection. Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new grubs or pupae in the potting medium/soil. [caption id="attachment_2130" align="aligncenter" width="482"]Steinernema carpocapsae nematodes Photo 3. Infective Juveniles of Steinernema carpocapsae nematodes[/caption]

Published papers

  1. Abbas, M.S.T., Saleh, M.M.E. and Akil, A.M. 2001.  Laboratory and field evaluation of the pathogenicity of entomopathogenic nematodes to the red palm weevil, Rhynchophorus ferrugineus (Oliv.) (Col.: Curculionidae). Anzeiger Fur Schadlingskunde-Journal of Pest Science. 74: 167-168.
  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. Dembilio, O., Karamaouna, F., Kontodimas, D. C., Nomikou, M. and Jacas, J. A. 2011.  Short communication. Susceptibility of Phoenix theophrasti (Palmae: Coryphoideae) to Rhynchophorus ferrugineus (Coleoptera: Curculionidae) and its control using Steinernema carpocapsae in a chitosan formulation. Spanish Journal of Agricultural Research 9: 623-626.
  4. Dembilio, O., Llacer, E., de Altube, M.D.M. and Jacas, J.A. 2010.  Field efficacy of imidacloprid and Steinernema carpocapsae in a chitosan formulation against the red palm weevil Rhynchophorus ferrugineus (Coleoptera: Curculionidae) in Phoenix. Pest Management Science 66: 365-370.
  5. Dembilio, O., Llacer, E., de Altube, M.D.M. and Jacas, J.A. 2010.  Field efficacy of imidacloprid and Steinernema carpocapsae in a chitosan formulation against the red palm weevil Rhynchophorus ferrugineus (Coleoptera: Curculionidae) in Phoenix canariensis. Pest Management Science. 66: 365-370.
  6. 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.
  7. 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.
  8. Llacer, E., de Altube, M.M.M. and Jacas, J.A. 2009.  Evaluation of the efficacy of Steinernema carpocapsae in a chitosan formulation against the red palm weevil, Rhynchophorus ferrugineus, in Phoenix canariensisBiocontrol. 54: 559-565.
  9. 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.
  10. Schroeder, W.J. 1992. Entomopathogenic nematodes for control of root weevils of citrus. Florida Entomologist 75: 563-567.
  11. Shah, N.K., Azmi, M.I. and Tyagi, P.K. 2011. Pathogenicity of Rhabditid nematodes (Nematoda: Heterorhabditidae and Steinernematidae) to the grubs of alfalfa weevil, Hypera postica (Coleoptera: Curculionidae). Range Management and Agroforestry 32: 64-67.
  12. Shapiro-Ilan, D. and Gardner, W.A. 2012. Improved control of Curculio caryae (Coleoptera: Curculionidae) through multi-stage pre-emergence applications of Steinernema carpocapsae. Journal of Entomological Science 47: 27-34.
  13. Shapiro-Ilan, D. and Hall, M.J. 2012. Susceptibility of adult nut Curculio, Curculio hicoriae (Coleoptera: Curculionidae) to entomopathogenic nematodes under laboratory conditions. Journal of Entomological Science 47: 375-378.
  14. Shapiro, D.I. and McCoy, C.W. 2000a. Susceptibility of Diaprepes abbreviatus (Coleoptera : Curculionidae) larvae to different rates of entomopathogenic nematodes in the greenhouse. Florida Entomologist. 83: 1-9.
  15. Shapiro, D.I. and McCoy, C.W. 2000b. Effects of culture method and formulation on the virulence of Steinernema riobrave (Rhabditida: Steinernematidae) to Diaprepes abbreviatus (Coleoptera: Curculionidae). Journal of Nematology 32: 281-288.
  16. 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.
  17. Simser, D. and Roberts, S. 1994. Suppression of strawberry root weevil, Otiorhynchus-ovatus, in cranberries by entomopathogenic nematodes (Nematoda, Steinernematidae and Heterorhabditidae. Nematologica 40: 456-462.

Twelve Important Facts about Beneficial Entomopathogenic Nematodes

Jan 31

1. What are insect-parasitic/entomopathogenic nematodes?

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

Insect-parasitic nematodes:

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

Entomopathogenic nematodes:

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

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

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

3. What is an infective juvenile?

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

4. What is a dauer juvenile?

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

5. Life cycle of entomopathogenic nematodes

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

6. How do entomopathogenic nematodes locate their insect hosts?

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

a. Ambush foraging:

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

b. Cruise foraging:

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

c. Intermediate foraging:

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

7. How are entomopathogenic nematodes produced?

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

In vivo production of entomopathogenic nematodes:

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

8. How to make a White trap?

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

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

Similarities:

 

Characteristics

Steinernematid Nematodes

Heterorhabditid Nematodes

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

Differences:

 

Characteristics

 Steinernematid Nematodes

Heterorhabditid Nematodes

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

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

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

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

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

12. How are entomopathogenic nematodes applied?

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

References

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

Described Steinernema species and their symbiotic bacteria

Jan 31

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:

Edding ton, S., Buddie, A.G., Tymo, L., Hunt, D.J., Nguyen, K.B., France, A.I., Merino, L.M. and Moore, D. 2009. Steinernema australe n. sp. (Panagrolaimorpha: Steinernematidae) a new entomopathogenic nematodefrom Isla Magdalena, Chile. Nematology 11: 699-717. Hazir, S., Stock, S. P. and Keskin, N. 2003. A new entomopathogenic nematode, Steinernema anatoliense. n. sp. (Rhabditida: Steinernematidae), from Turkey. Systematic Parasitology 55: 211-220. Lee, M. M., Sicard, M., Skeie, M, and Stock, S. P. 2009. Steinernema boemarei n. sp. (Nematoda: Steinernematidae), a new entomopathogenic nematode from southern France. Systematic Parasitology 72: 127-141. Lopez Nunez, J.C., Plichta, K., Gongora-Botero, C. and Stock, S.P. 2008. A new entomopathogenic nematode, Steinernema colombiense n. sp. (Nematoda: Steinernematidae) from Colombia. Nematology 10: 561-574. Luc, P. V., Nguyen, K.B., Reid, A.P. and Spiridonov, S.E. 2000. Steinernema tami sp. n. (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.  

Described Heterorhabditis species and their symbiotic bacteria

Jan 31

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.

Economically important insect pests and their susceptibility to major entomopathogenic nematodes

Jan 31

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

Species of insect pests

 Entomopathogenic nematode species

Publications

(See below)

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

Publications:

Apopka weevil, Diaprepes abbreviatus 1. Ali, J.G., Alborn, H.T. and Stelinski, L.L. 2010.  Subterranean herbivore-induced volatiles released by citrus roots upon feeding by Diaprepes abbreviatus recruit entomopathogenic nematodes. Journal of Chemical Ecology. 36: 361-368. 2. Bullock, R.C., Pelosi, R.R. and Killer, E.E. 1999. Management of citrus root weevils (Coleoptera: Curculionidae) on Florida citrus with soil-applied entomopathogenic nematodes (Nematoda: Rhabditida). Florida Entomologist. 82: 1-7. 3. Duncan, L. W., Stuart, R. J., El-Borai, F. E., Campos-Herrera, R., Pathak, E., Giurcanu, M. and Graham, J. H. 2013. Modifying orchard planting sites conserves entomopathogenic nematodes, reduces weevil herbivory and increases citrus tree growth, survival and fruit yield. Biological Control 64: 26-36. 4. Duncan, L.W and McCoy, C.W. 1996. Vertical distribution in soil, persistence, and efficacy against citrus root weevil (Coleoptera: Curculionidae) of two species of entomogenous nematodes (Rhabditida: Steinernematidae; Heterorhabditidae). Environmental Entomology. 25: 174-178. 5. Duncan, L.W. McCoy, C.W. and Terranova, A.C. 1996. Estimating sample size and persistence of entomogenous nematodes in sandy soils and their efficacy against the larvae of Diaprepes abbreviatus in Florida. Journal of Nematology. 28: 56-67. 6. El-Borai, F.E., Stuart, R.J., Campos-Herrera, R., Pathak, E. and Duncan, L.W. 2012.  Entomopathogenic nematodes, root weevil larvae, and dynamic interactions among soil texture, plant growth, herbivory, and predation. Journal of Invertebrate Pathology 109: 134-142. 7. Kaspi, R., Ross, A., Hodson, A.K., Stevens, G.N., Kaya, H.K. and Lewis, E.E. 2010. Foraging efficacy of the entomopathogenic nematode Steinernema riobrave in different soil types from California citrus groves. Applied Soil Ecology 45: 243-253. 8. Schroeder, W.J. 1992. Entomopathogenic nematodes for control of root weevils of citrus. Florida Entomologist 75: 563-567. 9. Shapiro, D.I. and McCoy, C.W. 2000. Susceptibility of Diaprepes abbreviatus (Coleoptera: Curculionidae) larvae to different rates of entomopathogenic nematodes in the greenhouse. Florida Entomologist. 83: 1-9. 10. Shapiro, D.I. and McCoy, C.W. 2000. Effects of culture method and formulation on the virulence of Steinernema riobrave (Rhabditida: Steinernematidae) to Diaprepes abbreviatus (Coleoptera: Curculionidae). Journal of Nematology 32: 281-288. 11. Shapiro, D.I., Cate, J. R., Pena, J., Hunsberger, A. and McCoy, C.W. 1999. Effects of temperature and host age on suppression of Diaprepes abbreviatus (Coleoptera: Curculionidae) by entomopathogenic nematodes. Journal of Economic Entomology. 92: 1086-1092. 12. Shapiro-Ilan, D.I., Mbata, G.N., Nguyen, K.B., Peat, S.M., Blackburn, D. and Adams, B.J. 2009. Characterization of biocontrol traits in the entomopathogenic nematode Heterorhabditis georgiana (Kesha strain), and phylogenetic analysis of the nematode's symbiotic bacteria. Biological Control 51: 377-387. 13. Shapiro-Ilan, D.I., Morales-Ramos, J.A., Rojas, M.G. and Tedders, W.L. 2010.  Effects of a novel entomopathogenic nematode-infected host formulation on cadaver integrity, nematode yield, and suppression of Diaprepes abbreviatus and Aethina tumidaJournal of Invertebrate Pathology. 103: 103-108. Armyworms, Heliothis armiger, Spodoptera exigua, S. frugiperda 14. Andalo, V., Santos, V., Moreira, G.F., Moreira, C., Freire, M. and Moino, A. 2012.   Movement of Heterorhabditis amazonensis and Steinernema arenarium in search of corn fall armyworm larvae in artificial conditions. Scientia Agricola 69: 226-230.  15. Ansari, M.A., Waeyenberge, L. and Moens, M. 2007.  Natural occurrence of Steinernema carpocapsae, Weiser, 1955 (Rhabditida: Steinernematidae) in Belgian turf and its virulence to Spodoptera exigua (Lepidoptera: Noctuidae). Russian Journal of Nematology 15: 21-24. 16. Kim, J. and Kim, Y. 2011. Three metabolites from an entomopathogenic bacterium, Xenorhabdus nematophila, inhibit larval development of Spodoptera exigua (Lepidoptera: Noctuidae) by inhibiting a digestive enzyme, phospholipase A (2). Insect Science 18: 282-288. 17. Negrisoli, A.S., Garcia, M.S., Negrisoli, C.R.C.B., Bernardi, D. and da Silva, A. 2010.  Efficacy of entomopathogenic nematodes (Nematoda: Rhabditida) and insecticide mixtures to control Spodoptera frugiperda (Smith, 1797) (Lepidoptera: Noctuidae) in corn crops. Crop Protection. 29: 677-683. 18. Salvadori, J.D., Defferrari, M.S., Ligabue-Braun, R., Lau, E.Y., Salvadori, J.R. and Carlini, C.R. 2012. Characterization of entomopathogenic nematodes and symbiotic bacteria active against Spodoptera frugiperda (Lepidoptera: Noctuidae) and contribution of bacterial urease to the insecticidal effect. Biological Control 63: 253-263.       Billbugs, Sphenophorus spp. 19. Georgis, R., Koppenhofer, A.M., Lacey, L.A., Belair, G., Duncan, L.W., Grewal, P.S., Samish, M., Tan, L., Torr, P. and van Tol, R.W.H.M. 2006. Successes and failures in the use of parasitic nematodes for pest control. Biological Control 38: 103-123. 20. Giometti, F.H.C., Leite, L.G., Tavares, F.M., Schmit, F.S., Batista, A. and Dell'Acqua, R. 2011.  Virulence of entomopathogenic nematodes (Nematoda: Rhabditida) against Sphenophorus levis (Coleoptera: Curculionidae). Bragantia 70: 81-86.   Black vine weevil, Otiorhynchus sulcatus 21. Ansari, M. A. and Butt, T. M. 2011.  Effect of potting media on the efficacy and dispersal of entomopathogenic nematodes for the control of black vine weevil, Otiorhynchus sulcatus (Coleoptera: Curculionidae). Biological Control 58: 310-318. 22. Ansari, M.A., Shah, F.A. and Butt, T.M. 2008.  Combined use of entomopathogenic nematodes and Metarhizium anisopliae as a new approach for black vine weevil, Otiorhynchus sulcatus control. Entomologia Experimentalis Et Applicata 129: 340-347. 23. Ansari, M.A., Shah, F.A. and Butt, T.M. 2010.  The entomopathogenic nematode Steinernema kraussei and Metarhizium anisopliae work synergistically in controlling overwintering larvae of the black vine weevil, Otiorhynchus sulcatus, in strawberry growbags. Biocontrol Science and Technology. 20: 99-105. 24. Haukeland, S. and Lola-Luz, T. 2010.  Efficacy of the entomopathogenic nematodes, Steinernema kraussei and Heterorhabditis megidis against the black vine weevil Otiorhynchus sulcatus in open field-grown strawberry plants. Agricultural and Forest Entomology.12363-369. 25. Lola-Luz, T. and Downes, M. 2007.  Biological control of black vine weevil Otiorhynchus sulcatus in Ireland using Heterorhabditis megidis. Biological Control 40: 314-319. 26. Susurluk, A. and Ehlers, R.U. 2008.  Sustainable control of black vine weevil larvae, Otiorhynchus sulcatus (Coleoptera: Curculionidae) with Heterorhabditis bacteriophora in strawberry. Biocontrol Science and Technology 18: 635-640. Bluegrass weevil, Listronotus maculicollis 27. McGraw, B.A. and Koppenhofer, A.M.2008.  Evaluation of two endemic and five commercial entomopathogenic nematode species (Rhabditida: Heterorhabditidae and Steinernematidae) against annual bluegrass weevil (Coleoptera: Curculionidae) larvae and adults. Biological Control 46: 467-475. 28. McGraw, B.A. and Koppenhofer, A.M.2009.  Population dynamics and interactions between endemic entomopathogenic nematodes and annual bluegrass weevil populations in golf course turfgrass. Applied Soil Ecology 41: 77-89. 29. McGraw, B.A., Vittumb, P.J. Cowlesc, R.S.and Koppenhoumlfera, A.M. 2010.  Field evaluation of entomopathogenic nematodes for the biological control of the annual bluegrass weevil, Listronotus maculicollis (Coleoptera: Curculionidae), in golf course turfgrass. Journal Biocontrol Science and Technology. 20: 149 – 163. Carpenter worms, Cossus cossus 30. Bazman, I., Ozer, N., and Hazir, S. 2008.  Bionomics of the entomopathogenic nematode, Steinernema weiseri (Rhabditida: Steinernematidae). Nematology 10: 735-742. Carrot weevil, Listronotus oregonensis 31. Belair, G. and Boivin, G.  1995. Evaluation of Steinernema-carpocapsae weiser for control of carrot weevil adults, Listronotus-oregonensis (leconte) (coleopteran: curculionidae), in organically grown carrots. Biocontrol Science and Technology 5: 225-231. 32. Miklasiewicz, T.J., Grewal, P.S., Hoy, C.W. and Malik, V.S. 2002. Evaluation of entomopathogenic nematodes for suppression of carrot weevil. Biocontrol 47: 545-561. Cat fleas, Ctenocephalides felis 33. Henderson, G., Manweiler, S.A., Lawrence, W.J., Tempelman, R.J.and Foil, L.D. 1995.  The effects of Steinernema-carpocapsae (weiser) application to different life stages on adult emergence of the cat flea Ctenocephalides-felis (bouche). Veterinary Dermatology 6: 159-163. 34. Silverman J.S., Platzer, E.G. and M.K. Rust, M.K. 1982. Infection of the cat flea, Ctenocephalides felis (Bouche) by Neoaplectana carpocapsae Weiser. Journal of Nematology 14: 394-397. Chestnut weevil, Curculio elephas 35. Karagoz, M., Gulcu, B., Hazir, S. and Kaya, H.K. 2009.  Laboratory evaluation of Turkish entomopathogenic nematodes for suppression of the chestnut pests, Curculio elephas (Coleoptera: Curculionidae) and Cydia splendana (Lepidoptera: Tortricidae). Biocontrol Science and Technology. 19: 755-768. 36. Kepenekci, I., Gokce, A. and Gaugler, R. 2004.  Virulence of three species of entomopathogenic nematodes to the chestnut weevil, Curculio elephas (Coleoptera: Curculionidae). Nematropica 34: 199-204. 37. Raja, R.K., Sivaramakrishnan, S. and Hazir, S. 2011.   Ecological characterisation of Steinernema siamkayai (Rhabditida: Steinernematidae), a warm-adapted entomopathogenic nematode isolate from India. Biocontrol 56: 789-798. Chinch bugs Bilssus spp. 38. Baxendale, F.P., A.P. Weinhold, and T.P. Riordan. 1994. Control of buffalograss chinch bugs with Beauvaria bassiana and entomopathogenic nematodes, 1993. Nebraska Insect Management and Insecticide Efficacy Reports, Dept. of Entomology Report No. 18, Univ. of Nebr., p. 43. Citrus root weevil, Pachnaeus litus 39. Bullock, R.C., Pelosi, R.R. and Killer, E.E. 1999.  Management of citrus root weevils (Coleoptera: Curculionidae) on Florida citrus with soil-applied entomopathogenic nematodes (Nematoda: Rhabditida). Florida Entomologist 82: 1-7.  40. Duncan, L.W., Graham, J.H., Dunn, D.C., Zellers, J., McCoy, C.W. and Nguyen, K. 2003.  Incidence of endemic entomopathogenic nematodes following application of Steinerema riobrave for control of Diaprepes abbreviates. Journal of Nematology 35: 178-186. 41. Schroeder, W.J. 1992. Entomopathogenic nematodes for control of root weevils of citrus. Florida Entomologist 75: 563-567. Clover root weevil, Sitona hispidulus 42. Loya, L.J. and Hower, A.A. 2002.  Population dynamics, persistence, and efficacy of the entomopathogenic nematode Heterorhabditis bacteriophora (Oswego strain) in association with the clover root curculio (Coleoptera: Curculionidae) in Pennsylvania.   Environmental Entomology 31: 1240-1250. 43. Loya, L.J. and Hower, A.A. 2003. Infectivity and reproductive potential of the Oswego strain of Heterorhabditis bacteriophora associated with life stages of the clover root curculio, Sitona hispidulus.   Journal of Invertebrate Pathology 83: 63-72. Codling moth, Cydia pomonella 44. Cossentine, J.E., Jensen, L.B. and Moyls, L. 2002. Fruit bins washed with Steinernema carpocapsae (Rhabditida: Steinernematidae) to control Cydia pomonella (Lepidoptera: Tortricidae). Biocontrol Science and Technology 12: 251-258. 45. de Waal, J.Y., Malan, A.P. and Addison, M.F. 2011.  Evaluating mulches together with Heterorhabditis zealandica (Rhabditida: Heterorhabditidae) for the control of diapausing codling moth larvae, Cydia pomonella (L.) (Lepidoptera: Tortricidae). Biocontrol Science and Technology 21: 255-270. 46. de Waal, J.Y., Malan, A.P., Levings, J. and Addison, M.F. 2010.  Key elements in the successful control of diapausing codling moth, Cydia pomonella (Lepidoptera: Tortricidae) in wooden fruit bins with a South African isolate of Heterorhabditis zealandica (Rhabditida: Heterorhabditidae). Biocontrol Science and Technology. 20: 489-502. 47. Lacey, L.A. and Chauvin, R.L. 1999. Entomopathogenic nematodes for control of diapausing codling moth (Lepidoptera: Tortricidae) in fruit bins. Journal of Economic Entomology 92: 104-109. 48. Lacey, L.A., and Unruh, T.R. 1998. Entomopathogenic nematodes for control of codling moth, Cydia pomonella (Lepidoptera: Tortricidae): Effect of nematode species, concentration, temperature, and humidity.  Biological Control 13: 190-197. 49. Lacey, L.A., Arthurs, S.P., Unruh, T.R., Headrick, H. and Fritts, R. 2006. Entomopathogenic nematodes for control of codling moth (Lepidoptera: Tortricidae) in apple and pear orchards: Effect of nematode species and seasonal temperatures, adjuvants, application equipment, and post-application irrigation. Biological Control 37: 214-223. 50. Lacey, L.A., Granatstein, D., Arthurs, S.P., Headrick, H. and Fritts, R. 2006. Use of entomopathogenic nematodes (Steinernematidae) in conjunction with mulches for control of overwintering codling moth (Lepidoptera: Tortricidae). Journal of Entomological Science 41: 107-119. 51. Lacey, L.A., Neven, L.G., Headrick, H.L. and Fritts, R. 2005.   Factors affecting entomopathogenic nematodes (Steinerneniatidae) for control of overwintering codling moth (Lepidoptera: Tortricidae) in fruit bins. Journal of Economic Entomology 98: 1863-1869. 52. Lacey, L.A., Shapiro-Ilan, D.I. and Glenn, G.M. 2010.   Post-application of anti-desiccant agents improves efficacy of entomopathogenic nematodes in formulated host cadavers or aqueous suspension against diapausing codling moth larvae (Lepidoptera: Tortricidae). Biocontrol Science and Technology. 20: 909-921. 53. Mracek, Z., Becvar, S., Kindlmann, P. and Webster, J.M. 1998.  Infectivity and specificity of Canadian and Czech isolates of Steinernema kraussei (Steiner, 1923) to some insect pests at low temperatures in the laboratory.  Nematologica 44: 437-448. 54. Navaneethan, T., Strauch, O., Besse, S., Bonhomme, A. and Ehlers, R.U. 2010.  Influence of humidity and a surfactant-polymer-formulation on the control potential of the entomopathogenic nematode Steinernema feltiae against diapausing codling moth larvae (Cydia pomonella L.) (Lepidoptera: Tortricidae). Biocontrol 55: 777-788. 55. Unruh, T.R., and Lacey, L.A. 2001. Control of codling moth, Cydia pomonella (Lepidoptera: Tortricidae), with Steinernema carpocapsae: Effects of supplemental wetting and pupation site on infection rate.  Biological Control 20: 48-56. 56. Vega, F.E., Lacey, L.A., Reid, A.P., Herard, F., Pilarska, D., Danova, E., Tomov, R. and Kaya, H.K. 2000.  Infectivity of a Bulgarian and an American strain of Steinernema carpocapsae against codling moth. Biocontrol 45: 337-343.  Crane flies, Tipula paludosa 57. Oestergaard, J., Belau, C., Strauch, O., Ester, A., van Rozen, K. and Ehlers, R.U. 2006.  Biological control of Tipula paludosa (Diptera: Nematocera) using entomopathogenic nematodes (Steinernema spp.) and Bacillus thuringiensis subsp israelensis. Biological Control 39: 525-531. 58. Simard, L., Belair, G., Gosselin, M.E. and Dionne, J. 2006.  Virulence of entomopathogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae) against Tipula paludosa (Diptera: Tipulidae), a turfgrass pest on golf courses. Biocontrol Science and Technology 16: 789-801. Cucurbit beetle, Diabrotica speciosa 59. Santos, V., Moino, A., Andalo, V., Moreira, C.C. and de Olinda, R.A. 2011. Virulence of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) for the control of Diabrotica speciosa Germar (Coleoptera: Chrysomelidae). Ciencia e Agrotecnologia 35: 1149-1156.  Cutworms, Agrotis ipsilon and A. segetum 60. Ebssa, L. and Koppenhofer, A.M.  2011. Efficacy and persistence of entomopathogenic nematodes for black cutworm control in turfgrass.   Biocontrol Science and Technology 21: 779-796. 61. Kunkel, B.A., Grewal, P.S. and Quigley, M.F. 2004.  A mechanism of acquired resistance against an entomopathogenic nematode by Agrotis ipsilon feeding on perennial ryegrass harboring a fungal endophyte.   Biological Control 29: 100-108. 62. Richmond, D.S., and Bigelow, C.A. 2009.  Variation in endophyte-plant associations influence Black Cutworm (Lepidoptera: Noctuidae) performance and susceptibility to the parasitic nematode Steinernema carpocapsae.  Environmental Entomology 38: 996-1004. 63. Shamseldean, M.M., Ibrahim, A.A., Zohdi, N., Shairra, S.A. and Ayaad, T.H. 2008.  Effect of Egyptian entomopathogenic nematode isolates on some economic insect pests.   Egyptian Journal of Biological Pest Control 18: 81-89. 64. Shapiro-Ilan, D.I., Mbata, G.N., Nguyen, K.B. Peat, S.M., Blackburn, D. and Adams, B.J. 2009. Characterization of biocontrol traits in the entomopathogenic nematode Heterorhabditis georgiana (Kesha strain), and phylogenetic analysis of the nematode’s symbiotic bacteria. Biological Control. 51: 377-387. 65. Shapiro-Ilan, D.I., Stuart, R.J. and McCoy, C.W. 2005. Characterization of biological control traits in the entomopathogenic nematode Heterorhabditis mexicana (MX4 strain). Biological Control 32: 97-103. 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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. The effect of nematodes on larvae of Phyllophaga menetriesi and Anomala inconstans (Coleoptera: Melolonthidae).  Revista Colombiana de Entomologia 33: 21-26. 250. Nguyen, K.B., and Buss, E.A. 2011. Steinernema phyllophagae n. sp (Rhabditida: Steinernematidae), a new entomopathogenic nematode from Florida, USA. Nematology 13: 425-442. White grubs, Rhizotrogus majalis 251. An, R. and Grewal, P.S. 2007.  Differences in the virulence of Heterorhabditis bacteriophora and Steinernema scarabaei to three white grub species: The relative contribution of the nematodes and their symbiotic bacteria. Biological Control 43: 310-316. 252. An, R.S., Sreevatsan, S. and Grewal, P.S. 2009.  Comparative in vivo gene expression of the closely related bacteria Photorhabdus temperata and Xenorhabdus koppenhoeferi upon infection of the same insect host, Rhizotrogus majalis. BMC Genomics. 10: 433. 253. Koppenhofer, A.M. and Fuzy, E.M. 2008. Attraction of four entomopathogenic nematodes to four white grub species. Journal of Invertebrate Pathology 99: 227-234. 254. Koppenhofer, A.M., Grewal, P.S. and Fuzy, E.M. 2006. Virulence of the entomopathogenic nematodes Heterorhabditis bacteriophora, Heterorhabditis zealandica, and Steinernema scarabaei against five white grub species (Coleoptera: Scarabaeidae) of economic importance in turfgrass in North America. Biological Control 38: 397-404. 255. Koppenhofer, A.M., Grewal, P.S. and Fuzy, E.M. 2007. Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species. Journal of Invertebrate Pathology 94: 184-195. Fuller rose beetle, Asynonychus godmani 256. Morse, J.G. and Lindegren, J.E. 1996. Suppression of fuller rose beetle (Coleoptera: Curculionidae) on citrus with Steinernema carpocapsae (Rhabditida: Steinernematidae). Florida Entomologist 79: 373-384. Chive gnat, Bradysia odoriphaga 257. Ma, J., Chen, S.L., Moens, M., Han, R.C. and De Clercq, P. 2013. Efficacy of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) against the chive gnat, Bradysia odoriphaga. Journal of Pest Science 86: 551-561. 258. Sun, R. H., A. H. Li, R. C. Han, L. Cao, and X. L. Liu. 2004. Factors affecting the control of Bradysia odoriphaga with entomopathogenic nematode Heterorhabditis indica LN2. Natural Enemies of Insects 26:150–155.

Employ three biological control agents to manage imported cabbageworms

Jan 31

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"]"The Damage by imported cabbageworms"[/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"]"The imported Cabbageworm"[/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:
Bélair, G., Fournier, C.Y. and Dauphinais, N. 2003. Efficacy of Steinernematid nematodes against three insect pests of Crucifers in Quebec.  Journal of Nematology 35: 259–265. Cai, J., Ye, G.Y. and Hu, C. 2004.  Parasitism of Pieris rapae (Lepidoptera: Pieridae) by a pupal endoparasitoid, Pteromalus puparum (Hymenoptera: Pteromalidae): effects of parasitization and venom on host hemocytes. Journal of Insect Physiology 50:315-322. Cameron, P.J. and Walker, G.P. 1997.  Host specificity of Cotesia rubecula and Cotesia plutellae, parasitoids of white butterfly and diamondback moth. Proceedings of 50th N.Z. Plant Protection Conference: 236-241 Herlihy, M.V., Van Driesche, R.G., Abney, M.R., Brodeur, J., Bryant, A.B., Casagrande, R.A., Delaney, D.A., Elkner, T.E., Fleischer, S J., Groves, R.L., Gruner, D.S., Harmon, J.P., Heimpel, G.E., Hemady, K., Kuhar,T.P., Maund, C.M., Shelton, A.M., Seaman, A.J., Skinner, M., Weinzierl, R., Yeargan, KV. And Szendrei, Z. 2012. Distribution of Cotesia rubecula (Hymenoptera: Braconidae) and its displacement of Cotesia glomerata in Eastern North America.  Florida Entomologist, 95:461-467. Mahar, A.N., Jan, N.D., Chachar, Q.I., Markhand, G.S., Munir M. and Mahar, A.Q. 2005. Production and infectivity of some entomopathogenic nematodes against larvae and pupae of Cabbage Butterfly, Pieris brassicae L. (Lepidoptera:Pieridae). Journal of Entomology 2: 86-91. Oatman, E. R.; Platner, G. R.; Greany, P. D. 1968. Parasitization of imported cabbageworm and cabbage looper eggs on cabbage in Southern California, with notes on the colonization of Trichogramma evanescens. Journal of Economic Entomology 61: 724-730. 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.

Five Simple Steps to Grow Organic Garlic

Jan 31

Organic Garlic Production

1. Preparation of Land for Garlic Planting:

Garlic is generally planted in late fall and harvested in late spring. Garlic can be grown in partially sunny areas but it thrives best in full sun. Clean the selected area by removing all the previous plant and weed debris. This can be used to make compost (Fig. 1). [caption id="attachment_610" align="aligncenter" width="300"]"Plant and weed debris for compost" Fig. 1. A pile of previous plant and weed debris used for making compost[/caption] Then loosen the soil with a tiller or a shovel and level it with a rake (Fig. 2). [caption id="attachment_600" align="aligncenter" width="300"]"Leveling of garlic filed" Fig. 2. The selected area should be leveled before planting.[/caption] After leveling the soil, cover the whole area with old newspapers. This will block out sunlight and help prevent germination of weed seeds, which in turn will save the trouble of weeding when the garlic is growing (Fig. 3) [caption id="attachment_601" align="aligncenter" width="300"]"Newspaper and organic weed control" Fig. 3. Old newspapers are easily available and can be used as a organic weed control in the garlic field.[/caption] Secure the newspapers with 6" wooden planks (Figs. 4 and 5) or any other available heavy material so that the papers do not fly away with the wind (Figs. 3 and 4). Using 6" planks allows a desirable distance of 6" to be maintained between two planting rows. While laying the planks on the newspaper, leave 3- 4 inch gap between the planks. In this way, planting rows will be formed automatically between the planks (Fig. 4). [caption id="attachment_608" align="aligncenter" width="300"]"Wooden planks for securing Newspapers" Fig. 4. Wooden planks used to secure newspapers and for forming rows for planting garlic.[/caption] After arranging wooden planks in each row, make holes with a metal rod or a wooden stick in the soil by ripping newspaper in a circular fashion at every 6 inches for planting garlic cloves (Fig. 5). [caption id="attachment_602" align="aligncenter" width="242"]" Hole in the soil for planting garlic clove" Fig. 5. A hole was made in the soil using a small metal rod to plant a garlic seed clove[/caption] In this way a standard distance of 6 inches would be maintained between two garlic plants and between two rows.  Note that the process described above (wooden planks and newspapers) is only feasible for a small garden and not for large acreage.

2. Planting Garlic:

After the above preparations are completed, break a garlic bulb into individual cloves (Fig. 6). Select comparatively large sized cloves for planting as large sized cloves will produce large sized garlic bulbs (Fig. 7). [caption id="attachment_604" align="aligncenter" width="278"]"Sigle clove can produce a garlic bulb" Fig. 6. Garlic cloves are separated from bulbs and individual clove used as garlic seeds. Each clove produces a garlic bulb.[/caption] [caption id="attachment_605" align="aligncenter" width="253"]"Select large size garlic cloves for planting" Fig. 7. Large size garlic cloves were selected for planting because they generally produce large size garlic bulbs.[/caption] Store these cloves in a cool place until you are ready tor plant, generally the beginning of November to the end of December is an ideal time here in Georgia. In the colder parts of the country, garlic should be planted 3-4 weeks before the ground freezes. The timing of planting is important to let the garlic cloves produce a good root system before winter sets in. When optimum moisture (at field capacity) is present in the soil, plant a single selected large sized clove with pointed end up in the hole at least 1.5 to 2.0 inches deep and cover it with a thin layer of soil (Fig. 8). [caption id="attachment_595" align="aligncenter" width="237"]"Planting of garlic seed clove" Fig. 8. As shown in the picture insert a single clove, with pointed end up, in the hole and cover it with a thin layer of soil.[/caption] Depending on the moisture in the ground and environmental temperature, garlic cloves will sprout within 7- 10 days of planting (Fig. 9). [caption id="attachment_606" align="aligncenter" width="260"]"The sprouted garlic clove" Fig. 9. Sprouted garlic clove 10 days after planting.[/caption] Allow garlic plants to grow until garlic plants show typical symptoms of their readiness to harvest. Fig. 10. shows growth of garlic 3 months after planting. [caption id="attachment_607" align="aligncenter" width="244"]"The garlic crop" Fig. 10. The growth of garlic crop three months after planting[/caption] Generally garlic bulbs are ready when the lower leaves turn yellowish or brownish in color and top leaves are still greenish in color (Fig. 11).  However, if you wait until all the leaves turn brown or become dry then it is too late to harvest the garlic. You will notice that all bulbs are divided (split) into separate cloves and they will not have enough leaf sheathes to wrap all the cloves together into an intact bulb, which in turn affects the storage life of garlic bulbs.  Also, there is a possibility that these divided bulbs can become targets for infection by fungus or any other disease causing organisms.

3. Harvesting of Garlic:

Harvesting garlic is a very easy process but you still need to take care to avoid bruising or injuring the bulbs so that their storage life will be enhanced.  Garlic bulbs sunburn easily and some varieties' flavor will change when exposed to the sun so select a cloudy day to harvest garlic. Th ground should also be soft (i.e. a few days after moderate amount of rain) for easy uprooting.  It is always advisable to loosen the soil beside the plant with a shovel or fork and then lift the plant (Fig. 11). This way the bulbs are not injured and remain intact with the stems/leaves, which are required for the process of proper curing. [caption id="attachment_592" align="aligncenter" width="300"]"Harvesting of Garlic" Fig. 11. For easy uprooting, first loosen the soil beside the plant with a shovel or fork and then lift the plant. along with garlic bulb[/caption] Then leave the harvested plants on the ground (if day is cloudy) for a couple of hours to dry the soil attached to the bulbs (Fig. 12). [caption id="attachment_596" align="aligncenter" width="300"]"Harvested garlic kept outside for a few hours for drying of attached soil" Fig. 12. Harvested garlic laid to dry out soil[/caption]

4. Curing of Garlic:

Shake the garlic plants to remove any extra dry soil from the bulbs and take them to the curing barn. For better curing, the curing barn should be a well-ventilated and warm but not hot.  For curing purposes, tie the stems/leaves of four plants together (Fig. 13) and then hang them on a stick or on the rope to form a single layer for easy drying/curing (Fig. 14). [caption id="attachment_598" align="aligncenter" width="195"]"The garlic plants for curing" Fig. 13. Several bunches of four harvested garlic plants that tied together for hanging on a stick for proper curing[/caption] [caption id="attachment_594" align="aligncenter" width="300"]"Method of curing of garlic" Fig. 14. For better curing, hang garlic plants in a single layer on a stick in the curing barn[/caption]

5. Storage of Garlic:

This process of curing generally takes several weeks. After curing, brush any remaining soil and loose leaf sheathes off of the bulbs, clip the roots, remove the stems and store the bulbs in mesh or paper bags (Fig. 15) in a well-ventilated and cool place. [caption id="attachment_593" align="aligncenter" width="179"]"Storage of cured garlic" Fig. 15. Store properly cured garlic in a mesh or paper bags in a well-ventilated and cool place.[/caption]    

Five beneficial insects that control the Squash Bug

Jan 31

Biological control of squash bug

The squash bugs (Anasa tristis) are the economically important pests of many plants in the Cucurbitae family. Adult bugs are grayish in color and about 5/8 inch long. [caption id="attachment_457" align="aligncenter" width="300" caption="The adult squash bug found on zucchini leaf"]"An adult squash bug"[/caption] Female bugs lay yellowish orange or reddish colored eggs on the underside of leaves or on stems. [caption id="attachment_458" align="aligncenter" width="300" caption="Squash bugs generally lay eggs in a group of 20- 25 on the underside of leaves but they can also lay eggs on the uppersurface of leaf"]"Eggs of squash bug"[/caption] Immediately after hatching from eggs, nymphs start feeding on the leaves, leaf stalks and stems, and become mature by going through five nymphal stages. [caption id="attachment_456" align="aligncenter" width="300" caption="Single Squash bug nymph feeding on a zucchini leaf"]"The squash bug nymph"[/caption] [caption id="attachment_551" align="aligncenter" width="300" caption="Squash bug nymphs feeding on Zucchini leaf stalks"]"The squash bug nymphs"[/caption] Both adults and nymphs suck cell sap from leaves and leaf stalks using their sucking piercing types of mouth parts. Heavy infestation causes wilting of leaves and eventually killing the entire plant. There are several species of predatory and parasitic insects that feed on the both mature and immature stages, and eggs of squash bugs. For example, predatory insects including the bigeyed bug (Geocoris punctipes), Pagasa fusca and the damsel bug (Nabis sp) directly munch on the all the stages of squash bugs whereas the feather-legged fly (Trichopoda pennipes) adults parasitizes both the nymphal and adult squash bugs but adults of the scelionid wasp (Gyron pennsylvanicum)parasitizes eggs of squash bugs. Thus these beneficial insects have a potential to keep populations of squash bugs below economic threshold level in your garden. Therefore, if you want to have presence of more of these beneficial insects in your organic garden, you need to plant specific types of attractive plants that will serve as food source for their adults to hang around in your garden. For example, the adult feather-legged flies that parasitize and kill the squash bugs in your organic garden are attracted to plants such as carrot, dill and parsley. [caption id="attachment_552" align="aligncenter" width="300" caption="Dill flowers can attract both predatory and parasitic insects to your organic garden"]"The dill plant"[/caption] The predatory bigeyed bugs are attracted to sunflowers whereas damsel bugs are attracted alfalfa, clover and radish flowers.

Literature:

Decker, K.B. and Yeargan, K.V. 2008. Seasonal phenology and natural enemies of the squash bug (Hemiptera : Coreidae) in Kentucky. Environmental Entomology 37: 670-678. Olson, D.L., Nechols, J.R. and Schurle, B.W. 1996.   Comparative evaluation of population effect and economic potential of biological suppression tactics versus chemical control for squash bug (Heteroptera: Coreidae) management on pumpkins. Journal of Economic Entomology 89: 631-639. Pickett, C.H. Schoenig, S.E. and Hoffmann, M.P.  1996. Establishment of the squash bug parasitoid, Trichopoda pennipes Fabr (Diptera: Tachnidae), in northern California. Pan-pacific Entomologist 72: 220-226. Vogt, E.A. and Nechols, J.R. 1993. Responses of the squash bug (Hemiptera, Coreidae) and its egg parasitoid, Gryon-pennsylvanicum (Hymenoptera, Scelionidae) to 3 cucurbita cultivars. Environmental Entomology 22: 238-245.

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

Jan 31
Interaction between flea beetles and entomopathogenic nematodes
  1. Flea beetles are called as flea beetles because they jump like fleas. Flea beetles are metallic black, blue, bronze or brown in color and about 1/16-1/8th inch long.
  2. Life cycle of flea beetles is very simple containing egg, larval and adult stages.  Flea beetles overwinter as adults by hiding under shelters including dry debris of plants (leaves and stems) left over from your garden crops or weeds. Early in the spring when temperature rises to about 50 F, the overwintering beetles become active and start feeding on the leaves of young plants. While feeding, they mate and lay eggs in the soil cracks around the root system of host plants or weeds in your garden or surrounding areas . Eggs hatch within 1-2 weeks and immediately larvae starts feeding on the roots of young host plants (see below) or weed hosts until they become mature. Then mature larvae pupate in the soil for 1-2 weeks; then emerge as adults and the life cycle continues. Generally this insect completes 2-3 generations in a year.
  3. Flea beetles are known to cause economic damage to many different vegetable crops including beans, broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, collards, corn, eggplant, kale, lettuce, melons, mustard,  peppers, potatoes, radishes, red Russian kale, rutabaga, spinach, squash, sunflowers, tomatoes, turnips and several species of weeds.
  4. Adults are the most damaging stage of flea beetles. They generally feed on foliage by chewing small holes through leaves and their heavy infestation gives a sieve-like appearance to the plant leaves thus reducing their marketable value especially leafy vegetables. Also, the heavy infestation of flea beetles can kill young seedlings.
  5. Flea beetle larvae feed on the plant roots but they do not cause a considerable economic damage to crop.
  6.  As temperature starts declining in the fall, adult flea beetles start looking for a shelter to hide and overwinter. Therefore, the process of management of flea beetles should begin in the fall to target overwintering beetles to reduce their incidence and outbreak in the next spring. The management of flea beetles should include both cultural and biological methods. Although the chemical insecticides could be more effective than other methods in controlling flea beetles, their use in the organic gardens should be avoided due to their detrimental effects on the human/animal health and environmental pollution.
  7. As a cultural control practice, keep your garden and its surrounding clean in the fall by removing all the plant debris including dry leaves and stems of harvested crops, weeds and other trash that may serve as the possible shelter for overwintering beetles.
  8.  Biological control method includes use of entomopathogenic nematodes (also called as insect-parasitic or beneficial nematodes) to target and kill larval and pupal stages of flea beetles in the spring.  Entomopathogenic nematodes can also attack and kill flea beetle adults if they come in contact in the soil.  Application of entomopathogenic nematodes such as Steinernema carpocapsae, Heterorhabditis bacteriophora and Heterorhabditis indica in the mid-late spring in your garden can kill both larval and pupal stages of flea beetles and thus reduce the emergence second generation adults, which are the most damaging to your crop.
  9. For the optimal rates and appropriate methods of application of entomopathogenic nematodes, read our blog at http://blog.bugsforgrowers.com/natural-predators/entomopathogenic-nematodes/beneficial-nematodes/how-to-deploy-your-nematode-army-and-kill-insect-pests/

Target Japanese beetle larvae with entomopathogenic nematodes in the fall

Jan 31

What are Japanese beetles?

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

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

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

Life cycle of Japanese beetle:

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

What are entomopathogenic nematodes?

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

Which species of entomopathogenic nematodes are effective against Japanese beetles?

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

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

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

What stages of Japanese Beetles can be targeted?

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

How can Entomopathogenic Nematodes kill Japanese beetle larvae?

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

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

For details read our blog

Literature:

Grewal, P.S., Koppenhofer, A.M., and Choo, H.Y., 2005.  Lawn, turfgrass and Pasture applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166. Koppenhofer, A.M., Fuzy, E.M., Crocker, R.L., Gelernter, W.D. and Polavarapu, S. 2004. Pathogenicity of Heterorhabditis bacteriophora, Steinernema glaseri, and S. scarabaei (Rhabditida : Heterorhabditidae, Steinernematidae) against 12 white grub species (Coleoptera : Scarabaeidae). Biocontrol Science and Technology. 14: 87-92. Maneesakorn, P., An, R., Grewal, P.S.and Chandrapatya, A. 2010. Virulence of our new strains of entomopathogenic nematodes from Thailand against second instar larva of the Japanese Beetle, Popillia japonica (Coleoptera: Scarabaeidae). Thai Journal of Agricultural Science.43: 61-66. Mannion, C.M., McLane, W., Klein, M.G., Moyseenko, J., Oliver, J.B. and Cowan D. 2001. Management of early-instar Japanese beetle (Coleoptera : Scarabaeidae) in field-grown nursery crops. Journal of Economic Entomology. 94: 1151-1161.

Control fleas using entomopathogenic nematodes

Jan 31
Fleas are one of the medically important pests of both animals and humans as they are capable of transmitting different kinds of disease causing organisms. Fleas are wingless insects but they can jump on their hosts including cats, dogs, humans and rats. Fleas have piercing and sucking type of mouthparts with that they suck blood of their hosts.  Like other insect, fleas also develop through the four different developmental stages including eggs, larva, pupa and adult.  Only adult fleas feed on blood but larval stage feeds on organic matter.  Pupa is a non-feeding stage. Fleas generally lay eggs on host’s body but they fall off on the ground where their host usually rests or sleeps.  Eggs hatch within 1-2 weeks and larvae immediately starts feeding on the organic matter that present at the resting place of animal hosts. Larvae develop through three larval stages and pupate in soil inside the silken cocoons. After 1-2 weeks, adult fleas emerge from cocoons but generally they use different kinds of host cues such as carbon dioxide, heat and vibration to emerge from pupae.  Fleas generally overwinter as larval and pupal stages, which can be easily targeted and killed by using biological control agents such as entomopathogenic nematodes.

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

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

Which species of entomopathogenic nematodes are effective against fleas?

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

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

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

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

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

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

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

How to apply entomopathogenic nematodes?

  • Entomopathogenic nematodes that you receive in sponge as liquid formulation are thoroughly mixed in water and can be easily sprayed directly on the area where animal hosts rests/sleeps using traditional Knapsack/backpack sprayers or watering cans.
  • However, at time of spraying care should be taken that the nematodes should not be allowed to settle at the bottom of sprayer or watering can to avoid their uneven distribution.
  • To avoid the settling of nematodes at the bottom of sprayer or watering can, nematode suspension should be constantly agitated.
  • However, nematodes will be easily damaged, if they are agitated through excessive recirculation of spray mix or if the temperature in the tank increases beyond 86oF.
  • Nematodes can also be applied through different types of irrigation systems but pumps should have proper pressure to avoid damage to nematodes and screen sizes should be larger than 50 mesh so that nematodes will pass through them live.
  • Nematodes received in granular formulation can be directly applied by broadcasting with hand or for larger area by using traditional spreaders that are used for application of granular or pallet pesticides or synthetic fertilizers.
  • Also, nematodes need about 20% moisture in the ground for survival. So please make sure nematode treated area should be watered immediately after the application of nematodes and continue to spray the area with water every few days.
[caption id="attachment_80" align="aligncenter" width="300" caption="Watering can for application of entomopathogenic nematodes on a small area"]Entomopathogenic nematodes can be applied with a watering can[/caption]

How Steinernema carpocapsae nematodes infect and kill fleas?

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

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

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

 Literature:

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

A Symposium on Entomopathogenic Nematodes and Multitrophic interactions in the Rhizosphere

Jan 31
A Symposium entitled “Entomopathogenic Nematodes and Multitrophic interactions in the Rhizosphere” has been organized by Raquel Campos-Herrera, Claudia Dolinski and Ganpati B. Jagdale at the Society of Nematologists 51st Annual meeting, which would be held in Savannah, Georgia from August 12th to 15th 2012. Following topics by various authors/speakers on interactions among entomopathogenic nematodes and multitrophic groups in the rhizosphere will be discussed between 8.0- 10.0 am on Tuesday August 14, 2012 in Marriot Riverfront hotel, Savanna, GA.   Topics with authors:
  1. Multitrophic interactions involving entomopathogenic nematodes applied against pine weevils in a forest ecosystem by Christine T. Griffin, A.M. Dillon, C.D. Harvey and C.D. Williams.
  2. Entomophathogenic nematodes: Effects of the soil agroecosystem on biological control potential by David I. Shapiro-Ilan, T.C. Leskey, S.E. Wright, I. Brown, and L. Fall.
  3. Interactions among entomopathogenic nematodes and other nematode trophic groups and plants in agroecosystems by Somasekhar Nethi, G.B. Jagdale and P.S. Grewal.
  4. Herbivore induced plants volatiles and entomopathogenic nematodes as agents of plant indirect defense by Jared G. Ali, H.T. Alborn, R. Campos-Herrera, F. Kaplan, L.W. Duncan, C. Rodriguez-Saona, A.M. Koppenhöfer, and L.L. Stelinski.

Heavy infestation of organically grown Chinese long beans by Kudzu bugs

Jan 31

Incidence of Kudzu bugs on Chinese long beans- bugsforgrowers

For the last couple of years, I have been growing Chinese long beans (Vigna unguiculata subsp. sesquipedalis) in my organic garden. Although this year (2012) I did not plant these beans in my garden, I noticed early in the spring that a few number of Chinese long bean seeds were voluntarily germinated. This means these seeds from last years crop were overwintered in the soil and germinated early in the spring when there was enough moisture in the ground and optimum temperature for their germination. So I let these voluntarily germinated vines/plants to grow in my organic garden. Over the growing season, these plants grew very well and healthy without the infestation of any insect pests. For the last few weeks these plants are producing a lot of pods that we have been harvesting at their maturity and shelling from them beans (grains) and preparing delicious curries. However, two weeks ago, I noticed that the vines of my Chinese long beans were heavily infested with Kudzu bug, Megacopta cribraria (Fig. 1). [caption id="attachment_387" align="aligncenter" width="300" caption="Fig.1. Chinese long bean vine is heavily infested with Kudzu bugs- Click on the image for its enlargement"]"Chinese long beans infested by Kudzu bugs"[/caption] Last year, I did not see any infestation of Kudzu bugs on long beans but as shown in pictures, they are now feeding on the pods (Fig. 2), leaves and stem (Fig. 3) of my Chinese long beans.   [caption id="attachment_388" align="aligncenter" width="300" caption="Fig. 2. Several adult of kudzu bugs are feeding and causing severe damage to my organically grown Chinese long bean pods- Click on the image for its enlargement"]"Kudzu bugs on Chinese long bean pods"[/caption] [caption id="attachment_389" align="aligncenter" width="300" caption="Fig. 3. Several nymphs of kudzu bugs are congregated and feeding on leaves and stem of Chinese long bean in my organic garden- Click on the image for its enlargement"]"Several greenish looking Kudzu bug nymphs"[/caption] The immature and mature stages of Kudzu bugs are dark brown (Fig. 2) and greenish brown (Fig. 3) in color, respectively. Both mature and immature stages of Kudzu bugs feed on the long bean vine/plant. Since these Kudzu bugs have sucking type of mouthparts, they suck cell sap from leaves, stem and pods. The intensive feeding by these bugs can cause leaves, stem and pods to dry and eventually cause the death of vines/plants.

Possible organic control measures

  • Although I don’t know how to control these Kudzu bugs organically, I have decided to spray garlic extract (prepared by grinding garlic in water) as a repellent. I will let you know the effects of garlic extract on the Kudzu bugs in my next blog.
  • Meanwhile, I have also exposed both nymphs and adults of Kudzu bugs to entomopathogenic Steinernema carpocapsae nematodes in a petri dish. I will also communicate with you the results of this experiment in my next blog.

Apply Heterorhabditis indica nematodes to kill small hive beetles

Jan 31

Why entomopathoegnic Heterorhabditis indica nematodes should be used to kill small hive beetles?

  • They are not harmful to honeybees and honeybee brood but can kill larvae or pupae of honeybee hive insect pest called small hive beetle within 48 hours of application.
  • They are commercially available and easy to apply using water cans or traditional knapsack sprayes.
  • They are not harmful to children, dogs, cats and personnel involved in its application.
  • Since they are exempted by EPA, no special permission need to apply them around honeybee hives against small hive beetles.

How do entomopathogenic nematodes kill small hive beetles?

When the infective juveniles of entomopathogenic nematodes are applied to the soil surface around bee hives, they start searching for their insect hosts such as larvae (grubs) or pupae of small hive beetles that are already present in the soil. Once larva and/or pupa has been located, the nematode infective juveniles penetrate into the body cavity of larva or pupa via natural openings such as mouth, anus and spiracles (breathing pores). Infective juveniles of Heterorhabditis nematodes can also enter by puncturing the inter-segmental membranes of the host cuticle. Once in the body cavity, infective juveniles of Steinernematid and Heterorhabditid nematodes release symbiotic bacteria, Xenorhabdus spp. and Photorhabdus spp., respectively from their gut in the blood of small hive beetle larva/pupae. In the blood, multiplying nematode-bacterium complex causes septicemia and kill mature larvae and/or pupae of small hive beetles usually within 48 hours after infection. Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the small hive beetle larval or pupal cadavers to seek new larvae small hive beetle that are already moved from bee hives in the soil for pupation.    

Protect honey bee hives from small hive beetle with Heterorhabditis indica

Jan 31

Honeybees:

Honeybees are the insects that are members of the genus Aphis belonging to an insect order, Hymenoptera, family, Aphidae and class, Insecta. These bees are well known for collection (Fig. 1) and storage of honey in the combs constructed by bees out of wax. Since honeybees visit hundreds of different kinds beautiful flowers to collect honey and pollen (Fig. 2), they are also very good pollinators of different plant species including crops grown in organic vegetable gardens. [caption id="attachment_384" align="aligncenter" width="179" caption="Fig. 1. A honey bee collecting honey"]"Honeybee"[/caption] [caption id="attachment_385" align="aligncenter" width="179" caption="Fig. 2. Honeybee visiting flowers for honey and pollen"]"A honeybee on flowers"[/caption]

What is a beekeeping?

Beekeeping is the practice in which beekeepers who raise honey bee colonies for the production of honey for their personal use or commercial purpose (i.e. selling honey, bee wax, royal jelly etc). Beekeepers also use these bees to pollinate crops.

What are bee hives?

Bee hives are the enclosed structures that are generally made out of wood by beekeepers for raising colonies of honey bees for the production and storage of honey. The active honey bee hives are also placed in the fields or near to organic gardens to use bees for the pollination of crops.

What are small hive beetles?

Small hive beetlesAethina tumida Murray (Coleoptera: Nitidulidae) are a destructive pest of honey bees, Aphis mellifera L. (Ellis and Delaplane, 2008). Although this insect pest is native to South Africa, its been around in North America since 1996 and responsible for causing serious economic damage to Apiculture (Beekeeping) industry in the United Sates.

How to identify small hive beetles?

Newly emerged young adults are light brown to red in color whereas mature adults are dark black in color and very active. Fully grown larvae (also called grubs) of small hive beetles are about 9-10 mm in length, 1.5- 2.0 mm wide and whitish in color. Pre-pupal stages are creamy white in color but mature pupae are dark brown in color. Both larval and adult stages found in active bee hives but pupal stages found about 3 feet away from bee hive and 10 - 20 cm deep in the soil.

Life cycle of small hive beetles

Under optimal climatic conditions, small hive beetles can complete their life cycle from egg to adult stage within 4-6 weeks and go through 5-6 generations in a year. Briefly, adult beetles immediately after emerging from pupae invade active honey bee colonies.  They find honey bee hives by using their olfactory system to detect smells released from their favorite foods such as honey, pollen or honey bee alarm pheromones. After locating hives, beetles enter into the bee colony and hide from bee attacks in the cracks and crevices of colony boxes.  In these cracks beetles mate and females lay over 1000 eggs in their life-time. Depending on the temperature, eggs hatch within 2-3 days after laying. Right after hatching from eggs, grubs enter into the comb and start feeding on honey, pollen and broods and matures in a couple of weeks. After maturation, larvae move into soil for pupation.  Generally, pupal stage lasts for 3-4 weeks and and life cycle begins again when new adults emerged from pupae.

How do small hive beetle cause damage to honey bee hives?

Immediately after hatching from eggs, small hive beetle larvae begin feeding on honey, pollen collected by bees and especially brood. During this feeding process they destroy honey combs. In case of heavy infestation of small hive beetles, bees will leave the colony. In addition, both adults and larvae of small hive beetles carry yeast (Scientific name: Kodamaea ohmeri) on their bodies into the colony (Benda et al., 2008). This yeast grows on honey combs and that ferments all honey in the comb and the compounds secreted by this yeast also attracts beetles to bee hives. This fermented honey becomes useless as food for honey bees as well as for human consumption. Small hive beetles can also damage stored honey as described above.

What options are available for the control of small hive beetles?

Following options are available for beekeepers to control small hive beetles. Chemical control: Although there are two chemical insecticides (including GuardStar and Checkmite) available to kill pupae and adults, respectively, extra care is needed as they may be toxic to humans, pets and the environment. Biological Control: Entomopathogenic nematodes can be used as biological control agents against small hive beetles (see below). These nematodes are commercially available and not harmful to animals, honeybees and humans. They are easy to apply using water cans or any traditional sprayers. According to USDA scientist Dr. Shapiri-Ilan and his colleagues, Heterorhabditis indica nematodes have a potential to suppress the population of small hive beetles (Shapiro-Ilan et al., 2010). These researchers also reported that H. indica can cause over 78% mortality of small hive beetles.

What stages of small hive beetles can be targeted with entomopathogenic nematodes?

  • Both mature larvae (grubs) and pupae are the best targets of entomopathogenic nematodes.
  • When mature grubs of small hive beetle moves in the soil for pupation, entomopathogenic nematodes should be applied to the soil surface within 3 feet area around honey bee hives to target and kill both mature grubs and pupae.

When to apply nematodes

  • As mature larvae of small beetles move away from bee hives and enter the soil to pupate, entomopathogenic nematodes should be applied on the soil surface.
  • Since entomopathogenic nematodes are very sensitive to UV light, they will die within a minute or two when exposed to direct sunlight. Therefore, nematodes should be applied early in the morning or late in the evening to avoid exposure to UV light.
  • Another advantage of applying entomopathogenic nematodes late in the evening around the bee hives is that these nematodes will be ready to attack the mature grubs/larvae of small hive beetles that generally move during night time to the soil to pupate.
  • In addition, these moving grubs will be easily found by cruiser entomopathogenic nematode like Heterorhabditis indica to attack mature larvae that are already entered in the soil (at 10-20 cm depth) to pupae and those larvae already pupated.

How many nematodes should be applied to obtain good control of small hive beetles?

See our table for the right dosages of each entomopathogenic nematode species to be applied for optimum control of small hive beetles.   Read following papers for detailed information on effect of entomopathogenic nematodes on the small hive beetles. Cabanillas, H.E. and Elzen, P.J. 2006.  Infectivity of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) against the small hive beetle Aethina tumida (Coleoptera : Nitidulidae) . Journal of Apicultural Research 45: 49-50. Ellis, J.D., Spiewok, S., Delaplane, K.S., Buchholz, S., Neumann, P. and Tedders, W.L. 2010.  Susceptibility of Aethina tumida (Coleoptera: Nitidulidae) larvae and pupae to entomopathogenic nematodes. Journal of Economic Entomology. 103: 1-9. Shapiro-Ilan, D.I., Morales-Ramos, J.A., Rojas, M.G. and Tedders, W.L. 2010.  Effects of a novel entomopathogenic nematode-infected host formulation on cadaver integrity, nematode yield, and suppression of Diaprepes abbreviatus and Aethina tumida. Journal of Invertebrate Pathology. 103: 103-108.

Susceptibility of black cutworms to beneficial nematodes

Jan 31
In my last blog post, I demonstrated the cutworms were susceptible to beneficial entomopathogenic Steinernema carpocapsae nematodes. These results are confirmed by a recent finding of Ebssa and Koppenhofer (2012), who also demonstrated that the Steinernema carpocapsae nematodes were highly effective against cutworm Agrotis ipsilon.  These researchers also demonstrated that the other species of beneficial nematodes including Heterorhabditis bacteriophora, Heterorhabditis megidis and Steinernema riobrave were effective in killing larval stages of cutworms.

Literature

Ebssa, L. and Koppenhofer, A.M. 2012. Entomopathogenic nematodes for the management of Agrotis ipsilon: effect of instar, nematode species and nematode production method.Pest Management Science 68: 947-957.

Naturally grown raspberries from our organic garden

Jan 31
Yesterday, we harvested fresh raspberries (see photo below) from the raspberry plants that are naturally growing at the borderline of our organic garden. [caption id="attachment_312" align="aligncenter" width="300" caption="Wild raspberries picked from naturally grown plants in our organic garden"]"Wild raspberries"[/caption] These raspberries are delicious and can be considered 100% organic because they are grown on the border of our organic garden in which we do not apply any chemical fertilizers as a food source for our vegetable plants and chemical insecticides for the control of insect pests. In our organic vegetable garden we use only compost as a nutrient source for plants. As an alternative to chemical pesticides, we use entomopathogenic nematodes as biological control agents to manage insect pest problem in our organic garden.

Tiny flea beetles causing a serious damage to my organic radish crop

Jan 31
Flea beetles are very small insects but they can cause a very serious damage to many [caption id="attachment_303" align="aligncenter" width="300" caption="A flea beetle feeding on a radish leaf (Click to enlarge)"]"Flea beetles are pests of radish"[/caption] vegetable crops including radish. Last month, I have planted 15-20 radish plants in my organic vegetable garden and they are growing very well because we had a good amount of rain. Last week, when I went to harvest some of these radish plants from my organic, I saw several holes on the leaves of radish. After careful examination, I found out that several adult flea beetles were hiding in the radish foliage and realized that these holes were caused by flea beetle damage. [caption id="attachment_304" align="aligncenter" width="300" caption="Severely damaged leaves of radish leaves by flea beetles (Click to enlarge)"]"Radish leaves damaged by flea beetles"[/caption] Since I have only a few plants, I thought I can catch them and kill them. However, my efforts failed because these tiny beetles are very active and able to runaway very fast. I am hoping that the natural enemies such as braconid wasps (I see sometime in my garden) may suppress the population of flea beetles. Application of environment friendly beneficial nematodes to target larvae of flea beetles would be an another option to manage flea beetles. Beneficial nematodes are not harmful to humans and pets.

Tent worms are susceptible to entomopathogenic nematodes

Jan 31
Last Monday, I read an article about eastern tent caterpillars on the website of Crossville Chronicle (http://crossville-chronicle.com/features/x1221402699/PLATEAU-GARDENING-Reader-inquires-about-Eastern-tent-caterpillars) and thought that I should share the results of my small experiment that I conducted about an interaction between tent worms and [caption id="attachment_292" align="aligncenter" width="300" caption="Tent worm caterpillar- bugsforgrowers"]"Tent worm caterpillar"[/caption] entomopathogenic nematodes. I hand picked four tent worms, which were crawling on my driveway and tested their susceptibility to an entomopathogenic Steinernema carpocapsae nematodes. In this experiment, I transferred 400 infective juveniles of Steinernema carpocapsae nematodes (100 juveniles/larva of tent worms) in 1 ml of water on a filter paper placed in a plastic dish (9 cm diameter) and then four tent worm larvae were transferred in the same plastic dish. [caption id="attachment_294" align="aligncenter" width="300" caption="Entomopathogenic Steinernema carpocapsae nematode infected Tent worm caterpillars"]"Entomopathogenic nematode infected Tent worms"[/caption] These plates were then incubated at room temperature for 48 hours. After 48 hours of incubation, I found that all the four tent worm larvae were dead.  This means entomopathogenic Steinernema carpocapsae nematodes infected and killed tent worms within 48 hours of infection. This showed me that the tent worms were susceptible to Steinernema carpocapsae nematodes. In order to confirm the infection by entomopathogenic nematodes, tent worm cadavers were transferred in a White trap for the emergence infective juveniles of entomopathogenic Steinernema carpocapsae nematodes. [caption id="attachment_295" align="aligncenter" width="300" caption="Entomopathogenic nematode Steinernema carpocapsae infected cadavers of Tent worm caterpillars in a White trap"]"Entomopathogenic nematode and Tent worms"[/caption] After 12 days, I saw under microscope that the thousands of infective juveniles of Steinernema carpocapsae were emerged from tent worm cadavers in a White trap . Thus these results suggest that the entomopathogenic nematodes can be used to kill tent caterpillars. However, for better I believe that you have to apply nematodes when tent worms are crawling are on the ground.

Colorado potato beetles on my organic potato plants

Jan 31
Yesterday (Saturday May 12, 2012), I found a bunch of Colorado potato beetle grubs (also called as larvae) were feeding on all of my four potato plants (Photo 1) that I had planted in December of 2011. This is the first time I have planted potatoes in my organic garden but did not expect that they will be attacked by Colorado potato beetles or any other insect pests.   [caption id="attachment_273" align="aligncenter" width="300" caption="Photo 1. Potatoes were planted in a organic garden in December 2012"]"Organically grown Potato plants"[/caption] I found that all the different stages of Colorado potato beetle grubs feeding at the same time on the potato leaves (Photo 2) but I did not come across the presence of their adult stages on plants. This means that these grubs are not from a same generation but they were from several different generations of beetles.   [caption id="attachment_274" align="aligncenter" width="300" caption="Photo 2. Different stages of Colorado potato beetle grubs found on organic potato plants"]"The different instars of Colorado potato beetle grubs"[/caption] I saw that all the stages of grubs were voraciously feeding (I wish, I had a video to record feeding) on the leaves as seen in photo number 3 these grubs have completely skeletonized the potato leaves.   [caption id="attachment_275" align="aligncenter" width="300" caption="Photo 3. Severe damage caused by Colorado potato beetle grubs to potato plants"]"The severe damage to potato leaves by Colorado potato beetle grubs"[/caption] I have collected almost all the mixed stages of grubs of Colorado potato beetle with my hand from my all potato plants in a container (photo 4).   [caption id="attachment_276" align="aligncenter" width="300" caption="Photo 4. Mixed stages of Colorado potato beetle grubs are collected for exposing to entomopathogenic Steinernema carpocapsae nematode"]"The grub stages of Colorado potato beetles"[/caption] I am going to expose these collected larvae of Colorado potato beetle to an entomopathogenic nematode, Steinernema carpocapsae. In my next blog post, I will share results of efficacy of Steinernema carpocapsae against these Colorado potato beetle larvae. I also noticed several masses of yellow colored eggs laid by Colorado potato beetles on the under side of several different potato leaves (Photo 5).   [caption id="attachment_277" align="aligncenter" width="200" caption="Photo number 5 showing a yellow egg mass laid by Colorado potato beetle female on the underside of a leaf of potato plant"]"The egg mass of Colorado potato beetle is yellow in color"[/caption] This means when these eggs will hatch there will be a heavy infestation of Colorado potato beetles on my potato plants. Therefore, to reduce future infestation of Colorado potato beetles I need to destroy all the egg masses on the leaves. I have destroyed the eggs by plucking the leaves with eggs and crushed them. This type of mechanical control of Colorado potato beetle is possible because I have only four potato plants in my garden. However, if you have large acreage of potato crop then you cannot destroy eggs this way but you have to let grubs hatch from eggs and then use any chemical pesticides or biological control agents such as entomopathogenic nematodes to manage a high population of hatched larvae Colorado potato beetles.