Kill leaf beetles (Altica quercetorum, Agelastica alni and Xanthogaleruka luteola) with Entomopathogenic Nematodes / December 21, 2008 by Ganpati Jagdale

The leaf beetles, Altica quercetorum and Agelastica alni are serious pests of urban trees including Quercus sp and Alnus sp, respectively.
The elm leaf beetle Xanthogaleruka luteola is a serious pest that causes defoliation of eml trees (Ulmus spp.) in North America.
Adults of these beetles generally feed on leaves by chewing holes through the leaf tissue.
Larvae skelotonize leaves by feeding on leaf tissues leaving veins and upper epidermis intact.
Entomopathogenice nematodes including Heterorhabditis megidis, Steinernema carpocapsae and S. feltiae can be used as potential biocontrol agents against different species leaf beetles (read Grewal et al., 2005 for more information).
It has been shown that both the pre-pupal and pupal stages of A. quercetorum and A. alni are very susceptible to H. megidis when applied in the soil.
The last instar larvae of X. luteola are highle susceptible to S. carpocapsae when applied to the mulch.

How Entomopathogenic Nematodes kill leaf beetles

When the infective juveniles are applied to the soil surface or mulch, they start searching for their hosts, in this case leaf beetles grubs.
Once a beetle grub has been located, the nematode infective juveniles penetrate into the grub body cavity via natural openings such as mouth, anus and spiracles.
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in grub blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills grubs 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 larvae in the soil.

References: Refer following book to read more about efficacy of entomopathogenic nematodes against leaf beetles

1. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). Nematodes As Biocontrol Agents. CAB publishing, CAB International, Oxon

Kill Japanese beetles (Popillia japonica) with Entomopathogenic Nematodes / December 20, 2008 by Ganpati Jagdale

The Japanese beetle, Popillia japonica, is a most economically important pest of many ornamental plants and turf grasses.
Larvae of these beetles are called white grubs that generally feed on roots of over 300 plants but their primary food source is grass roots. Severe damage caused by these grubs can result in dead patches of turf that can be picked up like a loose carpet.
Adults mostly feed on leaves and flowers by chewing the tissue between the veins, a type of feeding called skeletonizing.
Chemical insecticides including Imidacloprid (Merit), Chlorpyrifos, Isofenphos, and Diazinon are generally used to manage white grubs but due to human health and environment pollution concerns their use is restricted.
Currently, environmentally safe biological control agents including a milky disease causing bacterium Bacillus popilliae (Milky spores) and entomopathogenic nematodes have been used to control this pest.
Three entomopathogenic nematodes including Heterorhabditis bacteriophora GPS11 and TF strains, H. zealandica X1 strain and Steinernema scarabaei have been considered to be the most effective species against Japanese beetle grubs.
It has been demonstrated that the application of H. bacteriophora GPS11 and TF strains, H. zealandica X1 strain and S. scarabaei at rate of 2.5 billion infective juveniles per hectare can cause about 96, 98 and 100%, respectively control of Japanese beetle grubs infesting turfgrass (for more information read Grewal et a., 2005).
Nematodes can be applied using traditional sprayers that are used for the application of insecticides.
Nematodes perform better when they are applied to target small stages of grubs.
Nematodes also survive better and remain efficacious when field/lawns are irrigated before and after nematode applications.

How Entomopathogenic Nematodes kill Japanese beetles

When the infective juveniles are applied to the soil surface or thatch layer, they start searching for their hosts, in this case Japanese beetle grubs.
Once a Japanese beetle 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.
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in grub blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills Japanese beetle grubs 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 larvae in the soil.

References

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.

Why entomopathogenic nematodes are safe to use as biological control agents against insect pests? / December 5, 2008 by Ganpati Jagdale

Because....... 1. Entomopathogenic nematodes and their symbiotic bacterium have no detrimental effects on animals and plants. 2. Both nematodes and their symbiotic bacteria do not cause any harm to the personnel involved in their production and application. 3. Entomopathogenic nematode treated agriculture products are safe to handle and eat. 4. Entomopathogenic nematodes and symbiotic bacteria do not have any pathogenic effects on humans or animals. 5. When applied in the soil, entomopathogenic nematodes have also no negative effect on beneficial nematodes (bacteriovore, fungivore, omnivore and predatory) and other microbial communities. 6. Finally, entomopathogenic nematodes are non-polluting and thus environmentally safe.

Kill black cutworms (Agrotis ipsilon Hufnagel) with Entomopathogenic Nematodes / November 29, 2008 by Ganpati Jagdale

The black cutworm, Agrotis ipsilon (Hufnagel), is a polyphagous pest, feeding on almost all vegetables, many grain crops, ornamentals, turf grasses and weeds.
The plants damaged by black cutworms include beans, broccoli, cabbage, carrot, Chinese broccoli, Chinese cabbage, Chinese spinach, clover, corn, cotton, eggplant, flowering white cabbage, green beans, head cabbage, lettuce, mustard, potato, spinach, sugarcane, sweet potato, tomato, turnip, alfalfa, rice, sorghum, strawberry, sugarbeet, tobacco, bluegrass (Poa pratensis), curled dock (Rumex crispus); lambsquarters (Chenopodium album), yellow rocket (Barbarea vulgaris) and redroot pigweed (Amaranthus retroflexus).
There are five to nine larval instars that generally feed on seedlings at ground level by cutting off the stem causing a significant damage especially in newly planted fields. They also feed on roots and the below ground stem.
They can damage turfgrass by clipping off their blades and shoots.
The biological control agents including a bacterium Bacillus thuringiensis var. kurstaki, and entomopathogenic nematodes have a great potential against black cutworms.
Bacillus thuringiensis var. kurstaki produces a toxin that paralyzes the gut of the caterpillar. This toxin does not kill the caterpillars quickly, but it does cause the caterpillars to stop feeding, which in turn reducing the intensity of the damage.
Since caterpillars of cutworms are highly mobile insects, the entomopathogenic nematodes with ambush type of foraging strategy can be used very effectively for the management of cutworms.
For example, Steinernema carpocapsae, is an ambusher nematode species that can control black cutworms very effectively if applied at a rate of 1 billion nematodes/acre on golf course greens.

How Entomopathogenic Nematodes Kill Black Cutworms

When the infective juveniles are applied to the soil surface or thatch layer, they start searching for their hosts, in this case caterpillares.
Once a caterpillar has been located, the nematode infective juveniles penetrate into the caterpillar body cavity via natural openings such as mouth, anus and spiracles.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae) from their gut in the caterpillar blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills shore fly larvae 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 larvae in the potting medium/soil.

How entomopathogenic nematodes find their insect hosts (Foraging Strategies) / November 29, 2008 by Ganpati Jagdale

Infective juveniles of entomopathogenic nematodes use three different strategies to find their insect hosts.1. Ambush foraging: Ambushers such as Steinernema carpocapsae and S. scapterisci have adapted "sit and wait" strategy to attack highly mobile insects (billbugs, sod webworms, cutworms, mole-crickets and armyworms) when they come in contact 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. 2. Cruise foraging: Cruiser nematodes such as Heterorhabditis bacteriophora, H. megidis, Steinernema glaseri and S. kraussei generally move actively in search of hosts and therefore, they are distributed throughout the soil profile and more effective against less mobile hosts such as white grubs and black vine weevils. Cruisers never nictate but respond to carbon dioxide released by insects as cues. 3. Intermediate foraging: Some nematode species such as Steinernema feltiae and S.riobrave have adapted a strategy in between ambush and cruise strategies called an intermediate strategy to attack both the mobile and sedentary/less mobile insects at the surface or 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.

Kill leafminers (Liriomyza spp.) with Entomopathogenic Nematodes / November 23, 2008 by Ganpati Jagdale

Leafminers (Liriomyza spp.) are considered as economically important polyphagous pests of many indoor vegetable crops and flowering plants.
Vegetable host crops included beans, beet, carrots, celery, cucumbers, eggplants, lettuce, melons, onions, peas, peppers, potatoes, squash and tomatoes.
Flowering host plants included ageratum, aster, calendula, chrysanthemum, dahlia, gerbera, gypsophila, marigold, petunia, snapdragon, and zinnia.
Leafminer maggots generally feed on leaf parenchyma tissues by tunneling/mining between the upper and lower epidermal leaf surfaces.
Adults generally feed on sap exuding from the punctures caused by maggots during mining.
Infested leaves appear stippled due to the punctures made by leafminers while feeding, mining and oviposition especially at the leaf tip and along the leaf margins.
Widespread mining and stippling on the leaves generally decreases the level of photosynthesis in the plant leading towards the premature leaf drop reducing the amount of shade, which in turn causes sun scalding of fruits.
Injuries caused by maggots on the foliage also allow entry of bacterial and fungal disease causing pathogens.
Life cycle of leafminers contains four stages including egg, maggot, pupa and adult.
Life cycle can be completed within 15-21 days depending upon the host and temperature.
Adult females lay eggs in leaf tissues, eggs hatch within 2-3 days into maggots, hatched maggots starts feeding immediately and become mature within 3-4 days. Mature larvae eventually cut through the leaf epidermis and move to the soil for pupation and adults emerge within 3 weeks of pupation in the summer.
Although, chemical insecticides are generally used to protect foliage from injury caused by leafminers, but development of insecticide resistance among leafminer populations is a major problem.
Insecticides also are highly disruptive to naturally occurring biological control agents, particularly parasitoids.
Therefore, biological control agents including Bacillus thuringiensis var. thuringiensis (Bt), parasitic wasps (Diglyphus begina, D. intermedius, D. pulchripes and Chrysocharis parksi) and entomopathogenic nematodes (Heterorhabditis spp, Steinernema carpocapase and S. feltiae) have been considered as alternatives to chemical pesticides.
For successful control of leafminers, entomopathogenic nematodes can be easily applied in water suspension as spray application on plant foliage.
Entomopathogenice nematodes including S. carpocapase and S. feltiae when applied at the rate of 5.3 X 108 nematodes/ha can cause over 64% mortality of leafminers but need at least 92% relative humidity.

How Entomopathogenic Nematodes kill leafminers

When the infective juveniles are applied as spray to plant foliage, they enter the leaf mines through the leaf miner feeding punctures or exit holes made by the adults.
Once inside the mine the nematodes swim to find a leafminer maggot, nematodes then penetrate into the maggot body cavity via natural openings such as mouth, anus and spiracles.
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the larval cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the maggot blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills maggots usually within 48 h after infection.

For more information on the interaction between entomopathogenic nematodes and leafminers, please read following research and extension publications.

Hara, A.H., Kaya, H.K., Gaugler, R., Lebeck, L.M. and Mello, C.L. 1993. Entomopathogenic nematodes for biological control of the leafminer, Liriomyza trifolii (Dipt.: Agromyzidae). Entomophaga 38, 359-369.
Head, J. and Walters, K.F.A. 2003. Augmentation biological control utilising the entomopathogenic nematode, Steinernema feltiae, against the South American Leafminer, Liriomyza huidobrensis. Proceedings of the 1st International Symposium on Biological Control, (Hawaii, USA, 13-18 January 2002). USDA Forest Service, FHTET-03-05, 136-140.
Olthof, T.H.A. and Broadbent, A.B. 1992. Evaluation of steinernematid nematodes for control of a leafminer, Liriomyza trifolii, in greenhouse chrysanthemums. Journal of Nematology 24, 612.
Tong-Xian Liu, Le Kang, K.M.Heinz, J.Trumble. 2008. Biological control of Liriomyza leafminers: progress and perspective. CAB Reviews: Perspectives in Agriculture, Veterinary Science, Nutrition and Natural Resources, 2009, 4, No. 004, 16 pp.
Williams, E.C. and Walters, K.F.A. 1994. Nematode control of leafminers: Efficacy, temperature and timing. Brighton Crop Protection Conference - Pests and Disease. 1079-1084.
Williams, E.C. and MacDonald, O.C., 1995. Critical factors required by the nematode Steinernema feltiae for the control of the leafminers Liriomyza huidobrensis, Liriomyza bryoniae and Chromatomyia syngenesiae. Annals of Applied Biology. 127, 329-341.
Williams, E.C. and Walters, K.F.A. 2000. Foliar application of the entomopathogenic nematode Steinernema feltiae against leafminers on vegetables. Biocontrol Science and Technology 10, 61-70.

Kill Shore flies (Scatella stagnalis) with Entomopathogenic Nematodes / November 22, 2008 by Ganpati Jagdale

The shore fly, Scatella stagnalis (Fallén) (Diptera: Ephydridae) is an important insect pest of greenhouse plants.
Larvae of these flies mainly feed on blue-green algae grown on the surface of plant growing media, walls, floors, benches, and pots.
But larvae can also cause a serious damage to tender plant tissues thus reducing quality and productivity of plants.
The adults are not considered as plant feeders but they are nuisance to people and disseminate pathogens such as Fusarium and Pythium from plant to plant as they disperse through the greenhouse.
Currently, most growers rely on chemicals that kill host plants such as blue-green algae to reduce the incidence of shore flies. However, this method has not been proved effective in reducing shore fly incidence.
Biological control agents including Bacillus thuringiensis var. thuringiensis (Bt) and entomopathogenic nematodes have been considered as alternatives to chemical pesticides.
For successful control of shore flies, entomopathogenic nematodes can be easily applied in water suspension as spray application to the surface of plant growing medium.
Entomopathogenice nematodes including Heterorhabditis megidis, Steinernema arenarium and Steinernema feltiae when applied at the rate of 50 nematodes/cm2 can cause 94- 100% mortality of shore flies.

How Entomopathogenic Nematodes kill Shore flies

When the infective juveniles are applied to the surface of plant growing substrate, they start searching for their hosts, in this case shore fly larvae.
Once a larva has been located, the nematode infective juveniles penetrate into the larval body cavity via natural openings such as mouth, anus and spiracles.
Infective juveniles of Heterorhabditis spp also enter through the intersegmental members of the larval cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the larval blood.
In the blood, multiplying nematode-bacterium complex causes septicemia and kills shore fly larvae 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 larvae in the potting medium/soil.

For more information on the interaction between entomopathogenic nematodes and leafminers, please read following research and extension publications.

Foote, B.A. 1977. Utilization of blue-breen algae by larvae of shore flies. Environmental Entomology 6, 812-814.
Goldberg, N.P. and Stanghellini, M.E. 1990. Ingestion-egestion and aerial transmission of Pythium aphanidermatum by shore flies (Ephydrinae: Scatella stagnalis). Phytopathology 80, 1244-1246.
Lindquist, R., Buxton, J. and Piatkowski, J. 1994. Biological control of sciarid flies and shore flies in glasshouses. Brighton Crop Protection Conference, Pests and Diseases, BCPC Publications 3, 1067-1072.
Morton, A., Garcia del Pino, F., 2007. Susceptibility of shore fly Scatella stagnalis to five entomopathogenic nematode strains in bioassays. Biocontrol 52: 533-545.
Morton, A. and Garcia del Pino, F. 2003. Potential of entomopathogenic nematodes for the control of shore flies (Scatella stagnalis). Growing Biocontrol Markets Challenge Research and Development. 9th European Meeting IOBC/WPRS Working Group "Insect Pathogens and Entomopathogenic Nematodes", Abstracts, 67.
Vanninen, I., Koskula, H. 2000. Biological control of the shore fly (Scatella tenuicosta) with steinernematid nematodes and Bacillus thuringiensis var. thuringiensis in peat and rockwool. Biocontrol Sci. Technol.. 13: 47-63.
Zack, R.S. and Foote, B.A. 1978. Utilization of algal monoculture by larvae of Scatella stagnalis. Environmental Entomology 7, 509-511.

Kill Western Flower Thrips with Entomopathogenic Nematodes / November 18, 2008 by Ganpati Jagdale

The Western flower thrips, Frankliniella occidentalis is a most economically important pest of many field- and glasshouse-grown vegetables and ornamentals.
Adults lay eggs in the parenchyma tissue and there are two larval stages (first and second instars), prepupal and pupal stages are present in the life cycle of thrips.
Adult thrips generally feed by piercing and scraping of the stem, leaf, flower and fruit tissues.
Both instars also feed on all the aerial plant parts including leaves, flowers and fruits.
Piercing and scraping of the plant tissues leads to discoloration and drying of the damaged area, in some cases, abortion of flower/leaf buds or distortion of emerging leaves, thus reducing field crop yield and aesthetic value of ornamental plants.
Thrips are also capable of transmitting tospoviruses such as tomato spotted wilt virus (TSWV) and impatiens necrotic spot virus (INSV) during feeding, thus causing a tremendous loss to agricultural and horticultural greenhouse industries.
Controlling western flower thrips is difficult because of their small size and cryptic behavior.
Western flower thrips are commonly eradicated using endosulfan, chlorpyrifos, bendiocarb, and synthetic pyrethrinoids but use of these insecticides is restricted due to their environmental pollution and human health concerns, development of resistance to pesticides and removal of some of the most effective products from the market.
Biological control agents including predacious mites (Neoseilus cucumeris and Neoseilus degenerans), predacious bugs (Orius insidiosus), entomopathogenic fungi (Beauveria bassiana, Metarhizium anisopliae) and entomopathogenic nematodes (see below) have been used as alternatives to chemical pesticides.
The entomopathogenic nematodes species including Heterorhabditis bacteriophora, H. indica, H. marelata and Steinernema abassi, S. carpocapase, and S. feltiae have been found to be effective alternatives to chemical insecticides in controlling western flower thrips.
The entomopathogenic nematodes specifically attack soil-dwelling second instar larval, prepupal and pupal stages.
Generally, Heterorhabditis species are more effective than Steinernema species nematodes in controlling western flower thrips.
The insect- parasitic nematodes such as Thripinema nicklewoodii also have a potential to use as a biological control agent against western flower thrips.
Application of entomopathgenic nematodes at the rate of 400 infective juveniles/ cm2 of soil surface can cause over 50% mortality of thrip population.
Nematodes can be easily applied in water suspension as spray applications to the surface of plant growing medium or on the plant foliage infested with western flower thrips.
Although larval stages, prepupae and pupae are susceptible to entomopathogenic nematodes, H. bacteriophora HK3 strain can cause higher mortality of larval and prepupal stages than pupal stages

How Entomopathogenic Nematodes kill Western Flower Thrips

When the infective juveniles are applied to the surface of plant growing medium or injected in the potting medium, they start searching for their hosts, in this case Western Flower Thrip larvae, prepupae and pupae.
Once a larvae, prepupae and pupae has been located, the nematode infective juveniles penetrate into the larvae, prepupae and pupae body cavity via natural openings (mouth, anus and spiracles).
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub/pupa cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the larvae, prepupal and pupal blood.
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 larvae, prepupae and pupae in the potting medium/soil.

Biological Control of Black Vine Weevil using Insect Parasitic Nematodes / November 18, 2008 by Ganpati Jagdale

Black vine weevil, Otiorhynchus sulcatus is a common insect pest of over 150 plant species that grown in the greenhouses and nurseries.
Some of the plant species damaged by black vine weevils include Azalea, Cyclamen, Euonymus, Fuxia, Rosa, Rhododendron and Taxus.
Grubs (Larvae) of these weevils generally girdle the main stem, and feed and damage roots leading to nutrient deficiencies.
Adults feed on leaves and flowers by notching their edges thus reducing aesthetic value of plants.
The entomopathogenic nematodes species including Heterorhabditis bacteriophora, H. megidis and Steinernema carpocapase, S. feltiae and S. glaseri have been found to be effective alternatives to chemical insecticides such as chlorpyrifos (Dursban) in controlling black vine weevils.
Susceptibility of black vine weevil to nematodes is species and strain specific.
The rate of application of the nematode species/strains that tested against black vine weevil varies (5,000- 60,000 infective juveniles/pot) among different studies but nematodes applied at the rate of 5000- 20,000 infective juveniles/pot can cause up to 100% grub mortality.
Nematodes can be easily applied in water suspension as spray applications to the surface of plant growing medium but if nematodes are injected at depths deeper than 5 cm i.e. near to grubs they can cause highest mortality of grubs (70-93%) than those nematodes applied to the surface.
All the four larval stages (instars) and pupae of black vine weevil are susceptible to all entomopathogenic nematode species.
However, Heterorhabdtis bacteriophora can cause higher mortality of first and second instars than S. carpocapase and S. glaseri.
Also, all the three nematodes species are equally effective against third and fourth instars of black vine weevil.

How Entomopathogenic Nematodes Kill Black Vine Weevil

When the infective juveniles are applied to the surface of plant growing medium or injected in the potting medium, they start searching for their hosts, in this case black vine weevil grubs and pupae.
Once a grub/pupa has been located, the nematode infective juveniles penetrate into the grub or pupa body cavity via natural openings (mouth, anus and spiracles).
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the grub/pupa cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the grub blood.
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.

Kill fungus gnats using biological control agents: Insect-parasitic nematodes / November 14, 2008 by Ganpati Jagdale

Several fungus gnat species including Bradysia coprophila, B. impatiens and B. difformis are considered economically important indoor and greenhouse pests in Europe and the US.
Fungus gnat flies are black or gray in color with clear wings, relatively small (3-4 mm) in size and commonly associated with compost and natural soils with high organic contents.
You can see these hopping flies when you water your plants.
Fungus gnat maggots (larvae) are white-bodied with black heads and can be found just under the surface of the potting medium/soil.
These maggots primarily feed on fungi and organic matter but they can also cause a serious damage to many ornamental plants.
Maggots often chew or strip plant roots and tunnel stems affecting water and nutrient absorption in severely injured plants resulting in lost vigor, turn off-color and eventually death.
Maggots are also capable of transmitting fungal pathogens (Fusarium, Phoma, Pythium and Verticillium) during feeding.
Adult flies are nuisance to people and disseminate fungal spores from plant to plant as they disperse through the greenhouse.
Females often laying over 1000 eggs in a lifetime on the media surface and completing egg-to-egg life cycle within 20-25 days at 20-25oC.
Continuous and overlapping generations of fungus gnats in the greenhouse have made most control strategies difficult.
Currently, most growers rely on insecticides to manage fungus gnats in floriculture.
However, use of these insecticides is restricted due to their environmental pollution and human health concerns, development of resistance to pesticides and removal of some of the most effective products from the market.
Biological control agents including Bacillus thuringiensis (Bt), the predatory mite, Hypoaspis miles and entomopathogenic nematodes have been used as alternatives to chemical pesticides.
The entomopathogenic nematodes species including Heterorhabditis bacteriophora GPS11 strain, H. indica LN2 strain and Steinernema feltiae UK strain have a potential to use as biocontrol agents against fungus gnats.
These nematodes kill both maggots (larvae) and pupae, but the second and fourth stages are most susceptible than pupae.
Nematodes are generally applied in water suspension as spray applications to the surface of plant growing medium to target larval and pupal stages.
The potting medium (Ball-mix, Nursery-mix or Pro-mix) can influence the survival, persistence and efficacy of entomopathogenic nematodes in greenhouse production.
In the Nursery-mix, H. bacteriophora can survive longer and perform better than H. indica, H. marelatus Oregon, H. zealandica X1 and Steinernema feltiae against fungus gnats.
In the Pro-mix, only H. indica have performed better than all other nematode species that tested against fungus gnats.
Application of S. feltiae can cause 40% reduction in fungus gnat population in Ball-mix, 50% in Metro-mix and 56% in Pro-mix, but only 27% in the Nursery-mix.
In the greenhouse, temperature can influence efficacy of nematodes. For example, H. bacteriophora and H. indica can survive and cause very high mortality of fungus gnats at warmer (above 25oC) temperatures whereas S. feltiae is generally effective against fungus gnats at cooler (below 25oC) temperatures.
Application of an appropriate concentration of nematodes is a crucial step in the cost effective control of fungus gnats in greenhouse production.
Generally, application of one billion infective juveniles of H. bacteriophora, H. indica or S. feltiae per acre can kill over 50% fungus gnats in greenhouse productions.

How entomopathogenic nematodes kill fungus gnats

When the infective juveniles are applied to the surface of plant growing medium, they start searching for hosts, in this case fungus gnat maggots (larvae) and pupae.
Once a maggot/pupa has been located, the nematode infective juveniles penetrate into the maggot body cavity via natural openings such as mouth, anus and breathing pores called spiracles.
Infective juveniles of Heterorhabditis spp also enter through the intersegmental members of the maggot/pupal cuticle.
Once in the body cavity, infective juveniles release symbiotic bacteria (Xenorhabdus spp. for Steinernematidae and Photorhabdus spp. for Heterorhabditidae) from their gut in the fungus gnat blood.
Multiplying nematode-bacterium complex causes septicemia and kills the host usually within 48 h after infection.
Nematodes feed on multiplying bacteria, mature into adults, reproduce and then emerge as infective juveniles from the cadaver to seek new maggots in the potting medium/soil.

Nematodes are now commercially available from many suppliers distributed throughout in the USA.

For more information on biological control of fungus gnats, please read following research papers/book chapters:

Binns, E.S., 1973. Fungus gnats (Diptera: Mycetophilidae, Sciaridae) and the role of mycophagy in soil: a review. Rev. Ecol. Biol. Sol. 18, 77-90.
Chambers, R.J., Wright, E.M., Lind, R.J., 1993. Biological control of glasshouse sciarid larvae (Bradysia spp.) with the predatory mite, Hypoaspis miles on Cyclamen and Poinsettia. Biocontrol Sci. Technol. 3, 285-293.
Ecke, P.Jr., Faust, J.E., Williams, J., Higgins, A., 2004. The Poinsettia Manual. Ball Publishing, The Paul Ecke Ranch, Encinitas, California, USA.
Freeman, P., 1983. Sciarid flies, Diptera; Sciaridae. Handbooks for the identification of British insects 9, Part 6. London, Royal Entomol. Soc. pp 68.
Gillespie, D.R., Menzies, J.G., 1993. Fungus gnat vector Fusarium oxysporum f. sp. radicislycopersici. Ann. Appl. Biol. 123, 539-544.
Gouge, D.H., Hague, N.G.M., 1994. Control of sciarids in glass and propagation houses with Steinernema feltiae. Brighton Crop Protection Conference: Pest Dis. 3, 1073-1078.
Gouge, D.H., Hague, N.G.M., 1995. Glasshouse control of fungus gnats, Bradysia paupera, on fuchsias by Steinernema feltiae. Fundam. Appl. Nematol. 18, 77-80.
Grewal, P.S., Richardson, P.N., 1993. Effects of application rates of Steinernema feltiae (Nematoda: Steinernematidae) on control of the mushroom sciarid fly, Lycoriella auripila (Diptera: Sciaridae). Biocontrol Sci. Technol. 3, 29-40.
Grewal, P.S., Tomalak, M., Keil, C.B.O., Gaugler, R., 1993. Evaluation of a genetically selected strain of Steinernema feltiae against the mushroom sciarid fly, Lycoriella mali. Ann. Appl. Biol. 123, 695-702.
Harris, M.A., Oetting, R.D., Gardner, W.A., 1995. Use of entomopathogenic nematodes and new monitoring technique for control of fungus gnats, Bradysia coprophila (Diptera: Sciaridae), in floriculture. Biol. Control 5, 412-418.
Jagdale, G. B., Casey, M. L., Grewal, P. S. and Lindquist, R. K. 2004. Application rate and timing, potting medium and host plant on the efficacy of Steinernema feltiae against the fungus gnat, Bradysia coprophila, in floriculture. Biol. Contrl. 29: 296-305.
Jagdale, G. B., Casey, M. L., Grewal, P. S. and Luis Cañas. 2007. Effect of entomopathogenic nematode species, split application and potting medium on the control of the fungus gnat, Bradysia difformis (Diptera: Sciaridae), in the greenhouse at alternating cold and warm temperatures. Biol. Control. 43: 23-30.
Kim, H.H., Choo, H.Y., Kaya, H.K., Lee, D.W., Lee, S.M., Jeon, H.Y., 2004. Steinernema carpocapsae (Rhabditida: Steinernematidae) as a biological control agent against the fungus gnat Bradysia agrestis (Diptera: Sciaridae) in propogation houses. Biocontrol Sci. Technol. 14, 171-183.
Lindquist R., Piatkowski J. 1993. Evaluation of entomopathogenic nematodes for control of fungus gnat larvae. Bull. Int. Organiz. Biol. Integr. Control Noxious Animals and Plants. 16, 97-100.
Lindquist, R.K., Faber, W.R., Casey, M.L., 1985. Effect of various soilless root media and insecticides on fungus gnats. HortScience. 20, 358-360.
Menzel, F., Smith, J.E., Colauto, N.B., 2003. Bradysia difformis Frey and Bradysia ocellaris (Comstock): two additional neotropical species of black fungus gnats (Diptera : Sciaridae) of economic importance: a redescription and review. Ann. Entomol. Soc. Am. 96, 448-457.
Nielsen, G. R., 2003. Fungus gnats. http://www.uvm.edu/extension/publications/el/el50.htm
Oetting, R.D., Latimer, J.G., 1991. An entomogenous nematode Steinernema carpocapsae is compatible with potting media environments created by horticultural practices. J. Entomol. Sci. 26, 390-394.
Olson, D.L., Oetting, R.D., van Iersel, M.W., 2002. Effect of soilless media and water management on development of fungus gnats (Diptera: Sciaridae) and plant growth. HortScience. 37: 919-923.
Richardson, P.N., Grewal, P.S., 1991. Comparative assessment of biological (Nematoda: Steinernema feltiae) and chemical methods of control of mushroom fly, Lycoriella auripila (Diptera: Sciaridae). Biocontrol Sci. Technol. 1, 217-228.
Tomalak, M., Piggott, S. and Jagdale, G. B. 2005. Glasshouse applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166.
Wilkinson, J.D., Daugherty, D.M., 1970. Comparative development of Bradysia impatiens (Diptera: Sciaridae) under constant and variable temperatures. Ann. Entomol. Soc. Am. 63, 1079-1083.

Published Books and Related Literature on Beneficial Nematodes: Insect- and Slug- parasitc nematodes / May 2, 2008 by Ganpati Jagdale

Acarology IX. Edited by Roger, M., Horn, D.J., Needham, G.R., Welbourn, W.C., Ohio Biological Survey, Columbus, Ohio (1996).
A Handbook of Biology and Techniques. By Woodring, J.L., Kaya, H.K., Sothern Cooperative Bulletin 331, Arkansas Agricultural Experiment Station, Fayettville, Arkansas (1988).
Annual Report, CSIRO Division of Entomology (1981).
A Taxonomic Review of the Suborder Rhabditina (Nematoda: Secernentia) by Andrassy, I. ORSTROM, Paris (1983).
A worldwide guide to beneficial animals (insects, mites, nematodes) used for pest control purposes. By Thomson, W.T, Thomson Publications; Fresno, CA (1992).
Bioassays of Entomopathogenic Microbes and Nematodes. Edited by Navon, A., Ascher, K.R.S., CAB International, Wallingford, UK (2000).
Biological Control: Benefits and Risks. Edited by Hokkanen, H.M.T., Lynch, J.M., Cambridge University Press, UK (1995).
Biological Control of Mosquitoes. Edited by Chapman, H.C., American Mosquito Control Association Bulletin 6 (1985).
Biorational Pest Control Agents: Formulation and Delivery. Edited by Hall, F.R., Barry, J.W., American Chemical Society, Maryland (1995).
Cranberry Research Compilation. Progress and Final Reports on Cranberry Research Conducted in 1998. Edited by Deziel,G., Hogan, M., Cranberry Institute, Warenham, Massachsetts (1999).
Conservation Biological Control. Edited by Barbosa, P., Academic Press, San Diego, California (1998).
Control of Insect Pests with Entomopathogenic Nematodes. edited by Smith, K.A., Hatsukade, M., Food and Fertilizer Technology Center, Republic of China in Taiwan (1994).
Cost Action 819 Developments in Entomopathogenic Nematode/Bacterial Research. European Commission, Luxembourg (2001).
Cost 819 Entomopathogenic Nematode: Application and Pesistence of Entomopathogenic nematodes. European Communities, Luxembourg (1999).
Cost 819 Biotechnology-Ecology and Transmission Strategies of Entomopathogenic Nematodes. EOfficial Publication of EC, Luxembourg, EN 16269 EN (1995).
Cost 819. Entomopathogenic Nematodes- Pathogenicity of Entomopathogenic Nematodes Versus Insect Defence Mechanisms: Impact on Selection of Virulent Strains. Edited by Simoes, N. Boemare, N & Ehlers, R., Office for Official Publications of the European Communities (1988).
Ecology and Biology of Soil Organisms. Edited by Bhandari, S.C., Somani, L.L., Agrotech Publishing Academy, Udaipur (1994).
Ecology and Classification of North American Freshwater Invertebrates, 2nd edn., Edited by Thorp, J.H., Covich, A.P., Academic Press, San Diego, California (2001).
Ecology and Transmission Strategies of Etomopathogenic Nematodes. Edited by Griffin, C.T., Gwynn, R.L., Masson, J.P., European Commission, Luxembourg (1995).
Entomogenous Nematodes, A Manual and Host List Insect- Nematode Associations. By Poinar, G.O.Jr., E.J. Brill, Leiden, The Netherlands (1975).
Entomopathogenic Nematodes in Biological Control. Edited by Gaugler, R., Kaya, H.K., CRC Press, Boca Raton, Florida (1990).
Entomopathogenic Nematodes: Systematics, Phylogeny and Bacterial Symbionts. Edited by Nguyen, K.B. & Hunt, D. Brill, The Neitherland, (2007).
Entopathogenic Nematology. Edited by Gaugler, R., CAB International, Wallingford, UK (2002).
Environmental Persistance of Entomopathogens and Nematodes. Edited by Baur, M.E., Kaya, H.K., Fuxa, J., Southern Cooperative Series Bulletin 398, Oklahoma Agricultural Experiment Station, Stillwater, Oklahoma (2001).
Genetics of Entomopathogenic Nematode Bacterium-Complexes. European Commission, Luxembourg City, Luxembourg (1994).
Field Manual of Techniques in Invertebrate Pathology. Edited by Lacy, L.A., Kaya, H.K., Kluwer Academic Publishers, The Netherlands (2000).
Formulation of Microbial Biopesticides - Beneficial Microorganisms, Nematodes, Seed Treatments. Edited by Burges, H.D. 1998.Kluwer Academic Pub. Boston and Dordrecht 496p (1998).
Helminths of Insects. Edited by Sonin, M.D., Amerind Publishing, New Delhi (1987).
Integrated Pest Management for Turfgrass and Ornamentals. Edited by Leslie, A.R., CRC Press, Boca Raton, Florida (1994).
Manual of Agricultural Nematology. Edited by Nickle, W.R., Marcel Dekker, New York (1991).
Manual of Techniques in Insect Pathology. Biological Techniques Series. Edited by Lacey, L.A., Academic Press, San Diego, California (1997).
Manual of Techniques in Invertebrate Pathology. Edited by Lacey, L.A., Kaya, H.K., Kluwer Academic Publishers, Dordrecht, The Netherlands (2000).
Methods in Biotechnology: Biopesticides: Use and Delivery. Edited by Hall, F.R., Menon, J. Humanan Press Inc, Totowa, New Jersey (1998).
Microorganismos patógenos empleados en el control microbiano de insectos plaga. Edited by Lecuona, R., Talleres Gráficos Mariano Mas, Buenos Aires (1996).
Natural Enemies of Terrestrial Molluscs. Edited by Barker, G.M., CAB International, Wallingford, UK (2003).
Nematodes and the Biological Control of Insect Pests. Edited by Bedding, R.A., Akhurst, R., Kaya, H.K., CSIRO Publications, East Melbourne, Australia (1993).
Nematodes as Biocontrol Agents. Edited by Grewal P. S., Ehlers, R.-U., Shapiro-Ilan, D.I., CABI Publishing, CAB International, Wallingford, UK (2005).
Nematodes for Biological Control of Insects. Edited by Poinar, G.O.Jr., CRC Press, Boca Raton, Florida (1979).
Nematology, Advances and Perspective. Edited by Chen, Z.X., Chen, S.Y. and Dickson, D.W.,Tsinghua University Press, TUP, China (2001).
New Directions in Biological Control. Edited by Baker, R.P., Dunn, P.E., Liss, New York (1990).
Plant and Insect Nematodes. Edited by Nickle, W. R., Marcel Dekker, New York (1984).
Parasites and Pathogens of Insects. Edited by Beckage, N.E., Thompson, S.N., Federici, B. Academic Press, New York (1993).
Pest management in the subtropics. Biological control - A Florida perspective. Edited by Rosen, D., Bennett, F.D. and Capinera, J.L., Intercept, Andover, UK (1994).
Proceedings of Workshop on Optimal Use of Insecticidal Nematodes in Pest Management. Edited by Polavarapu, S., Rutgers University, New Brunswick, New Jersey (1999).
Recent Advances in Biological Control of Insect Pests by Entomogenous Nematodes in Japan. Edited by Ishibashi, N., Saga University, Japan (1987).
Recent Advances in Nematology. Edited by Dwivedi, B.K., Bioved Research Society, Allahabad, India (1992).
Slug and Snail Pests in Agriculture. Edited by Henderson, I., Symposium Proceedings No. 66, British Crop Protection Council Farnham, UK (1996).
Slugs & Snails: Agricultural, Veterinary & Environmental Perspectives. Monograph No. 80, British Crop Protection Council, Thornton Health, UK (2003).
Survival Strategies of Entomopathogenic Nematodes. Edited by Glazer, I., Richardson, P., Boemare, N., Coudert, F., EUR 18855 EN Report (1999).
The Biology of Nematodes. Edited by Lee, D.L., Taylor & Francis, London (2002).
The Natural History of Nematodes by Pionar, G.O.Jr., Prentice-Hall, Englewood Cliffs, New Jersey (1983).
Tick-Born Diseases and their Vectors. Edited by Dusbabek, R., Bukva, V., SPB Academic Publishers, The Hague, The Netherlands (1991).
Tick Born Pathogens at the Host Vector Interface: A Global Perspective. Edited by Coons, L., Rothschild, M., Krugel, National Park, South Africa (1995).
Tortricid Pests, Their Biology, Natural Enemies and Control. Edited by Van der Geest,L.P.S., Evenhius, H.H., elsevier Science, Amsterdam, The Netherlands (1991).
Tylenchida Parasites of Plants and Insects. By Siddiqi, M.R., CAB Farnham Royal, Slough, UK (1986).
Tylenchida Parasites of Plants and Insects. By Siddiqi, M.R., CAB International, Wallingford, UK (2000).
Use of Pathogens in Scarab Pest Management. Edited by Jackson, T.A., Glare, T.R., Ag Research, Lincoln, New Zealand (1992).

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List of Research and Extension Journals Accepting Papers on Beneficial Nematodes / May 2, 2008 by Ganpati Jagdale

Acta Entomologia Bohemoslovaca
Acta Horticulturae
Acta Parasitologica
Acta Phytopathologica et Entomologica Hungarica
Acta Phytophylacica Sinica
Actes des Colloques Insectes Sociaux
Advances in Parasitology
Agricultural Ecosystems and Environment
Agricultural Systems
Agro Food Industry Hi-Technology
American Bee Journal
American Naturalist
Annals of Applied Biology
Annals of the Entomological Society of America
Annals of the New York Academy of Science
Annual Review of Entomology
Annual Review of Microbiology
Annual Review of Phytopathology
Applied Entomology and Zoology
Applied and Environmental Microbiology
Applied Microbiology and Biotechnology
Applied Soil Ecology
Arthropod Management Tests
Australian Journal of Experimental Agriculture
Behaviour
Biocontrol Science and Technology
Biocontrol
Biodiversity and Conservation
Biological Control
Biology and Fertility of Soils
Biotechnology and Bioengineering
Biotechnology Progress
Brighton Crop Protection Conference- Pest and Disease
Bulletin of Entomological Research
Bulletin of Entomological Society of America
Bulletin of the Faculty of Agriculture, Saga University
Bulletin of the Georgian Academy of Sciences
Bulletin of the Institute of Maritime Tropical Medicine, Gdynia
Bulletin of the Institute of Zoology, Academia Sinica
Bulletin of the International Organization for Biological and Integrated Control of Noxious Animals and Plants
Bulletin OILB/SROP
California Agriculture
Canadian Entomologist
Canadian Journal of Zoology
Cellular and Molecular Life Sciences
Chinese Journal of Tropical Crops
Comparative Biochemistry and Physiology B
Crop Protection
Current Genetics
Current Opinion in Microbiology
Ecology
Egyptian Journal of Biological Pest Control
Egyptian Journal of Agronematology
Entomologia Expermentalis et Applicata
Entomophaga
Environmental Entomology
Experientia
Experimental and Applied Acarology
Experimental Parasitology
FEMS Microbiology Letters
Florida Entomologist
Folia Parasitologica
Forest Ecology and Management
Forest Research
Gene
Indian Journal of Agricultural Sciences
Indian Journal of Entomology
Indian Journal of Nematology
Insect Science and Its Application
Israel Journal of Medical Science
Integrated Pest Management Reviews
International Journal for Parasitology
International Journal of Nematology
International Journal of Parasitology
International Journal of Systematic Bacteriology
International Journal of Systematic and evolutionary Microbiology
International Organization for Biological and Integrated Control Bulletin
International Research Communications System Medical Science: Microbiology, Parasitology and Infectious Diseases
IOBC/WPRS Bulletin
Irish Journal of Agricultural and Food Research
Japanese Journal of Applied Entomology and Zoology
Japanese Journal of Nematology
Journal for Hawaiian and Pacific Agriculture
Journal of Agricultural Research
Journal of American Mosquito Control Association
Journal of Animal Ecology
Journal of Applied Ecology
Journal of Applied Entomology- Zeitschrift fur Angewandte Entomologie
Journal of Arboriculture
Journal of Australian Entomological Society
Journal of Bacteriology
Journal of Economic Entomology
Journal of Egyptian Society of Parasitology
Journal of Entomological Science
Journal of Environmental Horticulture
Journal of chemical Ecology
Journal of Clinical Microbiology
Journal of General Microbiolgy
Journal of Helminthology
Journal of Industrial Microbiology and Biotechnology
Journal of Insect Pathology
Journal of Invertebrate Pathology
Journal of Kansas Entomological Society
Journal of Medical Entomology
Journal of Molluscan Studies
Journal of Natural Products
Journal of Nematology
Journal of Parasitology
Journal of Genetic Microbiology
Journal of the Australian Entomological Society
Journal of the Entomological Society of British Columbia
Journal of the Georgia Entomological Society
Journal of Thermal Biology
Journal of West China University of Medical Sciences
Korean Journal of Applied Entomology
Korean Journal of Turfgrass Science
Medical and Veterinary Entomology
Memoirs of the Entomological Society of Canada
Microbial Ecology
Molecular Phylogenetics and Evolution
Mosquito News
Mushroom News
Natural Enemies of Insects
Nature
Nature Biotechnology
Nematology
Nematropica
Netherlands Journal of Plant Pathology
New Zealand Entomologist
New Zealand Journal of Experimental Agriculture
New Zealand Journal of Zoology
Oecologia
Pakistan Journal of Nematology
Parasitology
Parasitology Research
Pedobiologica
Pest Management Science
Phytoparasitica
Phytoprotection
Plant Protection Quarterly
Polskie Pismo Entomologiczne
Proceedings of the American Chemical Society, Division of Environmental Chemistry
Proceedings of the Florida State Horticultural Society
Proceedings of the Entomological Society of Washington
Proceedings of the Helminthological Society of Washington
Proceedings of the North Central Branch of Entomological Society of America
Research and Reviews in Parasitology
Revista de la Sociedad Entomologica Argentina
Revista de Proteccion Vegetal
Revue de Nematologie
Russian Journal of Nematology
Science
Sri Lanka Journal of Tea Science
SPOR/WPRS Bulletin
Systematic and Applied Microbiolgy
Systematic Parasitology
The Canadian Entomologist
Trends in Parasitology
Veterinary Dermatology
Zhonghua Kunchong

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Symbiotic bacteria of Heterorhabdits nematodes- Photorhabdus species / April 6, 2008 by Ganpati Jagdale

Heterorhabditis amazonensis- undescribed
H. argentinensis- P. temperata
H. bacteriophora- Photorhabdus luminescens subsp. laumondii TT01, P. luminescens kayaii subsp. nov., P. luminescens thracensis subsp. nov., P. temperate
H. baujardi- undescribed
H. brevicaudis- P. luminescens
H. downesi- Photorhabdus sp
H. floridensis- undescribed
H. georgiana- undescribed
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 XlNach
H. mexicana- undescribed
H. poinari- Photorhabdus sp
H. safricana- undescribed
H. taysearae- undescribed
H. zealandica- P. temperata

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Symbiotic bacteria of Steinernematid nematodes- Xenorhabdus species / April 6, 2008 by Ganpati Jagdale

Steinernema abbasi- undescribed
S. aciari- undescribed
S. affine-Xenorhabdus bovienii
S. akhursti- undescribed
S. anatoliense- undescribed
S. apuliae- undescribed
S. arenarium- X. kozodoii
S. ashiuense- undescribed
S. asiaticum- undescribed
S. australe- X. magdalenensis
S. backanense- undescribed
S. beddingi- undescribed
S. bicornutum- X. budapestensis
S. carpocapsae- X. nematophila
S. caudatum- undescribed
S. ceratophorum- undescribed
S. cholashanense- undescribed
S. cubanum- X. poinarii
S. cumgarense- undescribed
S. diaprepesi- undescribed
S. eapokense- undescribed
S. feltiae- X. bovienii
S. glaseri- X. poinarii
S. guangdongense- undescribed
S. hebeiense- undescribed
S. hermaphroditum- undescribed
S. intermedium - X. bovienii
S. jollieti-undescribed
S. karii- undescribed
S. khoisanae- undescribed
S. kraussei- X. bovienii
S. kushidai- X. japonica
S. leizhouense- undescribed
S. litorale- undescribed
S. loci- undescribed
S. longicaudum- undescribed
S. monticolum- undescribed
S. neocurtillae- undescribed
S. oregonense- undescribed
S. pakistanense- undescribed
S. puertoricense- X. romanii
S. rarum- X. szentirmaii
S. riobrave- Xenorhabdus sp
S. ritteri- Xenorhabdus sp
S. robustispiculum- undescribed
S. sangi- undescribed
S. sasonense- undescribed
S. scapterisci- X. innexi
S. scarabaei- X. koppenhoeferi
S. serratum- X. ehlersii
S. siamkayai- X. stockiae
S. sichuanense- X. bovienii
S. silvaticum- undescribed
S. tami- Xenorhabdus sp
S. texanum- undescribed
S. thanhi- undescribed
S. thermophilum- X. indica
S. websteri- undescribed
S. weiseri- undescribed
S. yirgalemense- undescribed

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Beneficial Nematodes: Steinernema and Heterorhabditis species / March 30, 2008 by Ganpati Jagdale

Entomopathogenic nematodes in the genera Steinernema and Heterorhabditis are recognized as insect-parasitic nematodes, beneficial nematodes, biocontrol agents, biological control agents, biological insecticides or biopesticides. These nematodes are also recognized as pathogens or microbial control agents because of their symbiotic association with bacteria (Xenorhabdus and Photorhabdus spp.) that are mainly pathogenic to insects. Because of mutualistic relationship with pathogenic bacteria these nematodes are named as entomopathogenic nematodes.

These beneficial nematodes contribute to the regulation of natural populations of insects. However, the population of naturally occurring entomopathogenic nematodes is normally not high enough to manages soil dwelling plant pests. Therefore, during last 3-4 decades, these live nematodes have been commercially mass produced and inundatively applied to control many garden insects, turfgrass insects, nursery insects, greenhouse insects and insects that feed on different field crops.

Use of this natural control of insects is beneficial for both the environment and humans because it reduces use of chemical insecticides/pesticides.

These biopesticides (entomopathogenic nematodes and their symbiotic bacteria) are safe to produce and not harmful to users, application personnel, mammals, most beneficial insects or plants.

Since entomopathogenic nematodes do not cause any health risk to the consumers of nematode treated agricultural produce and damage to the environment, they are exempted from registration requirements in most countries.

These biological control agents have also no detrimental effect on other benefical nematodes including bacterial feeders, some fungal feeders (Aphelenchus sp.), predatory nematodes and other soil microbial communities.

But entomopathogenic nematodes can be detrimental to plant-parasitic nematodes that are responsible for causing a tremendous economic loss to our agriculture industry throughout world. It has been demonstrated that entomopathogenic nematodes can suppress the populations of many economically important plant-parasitic nematodes including foliar nematodes, potato cyst nematodes, ring nematodes, root-knot nematodes, root lesion nematodes, sting nematodes, stubby root nematodes and stunt nematodes.

Scientific publications on Entomopathogenic Nematodes / March 22, 2008 by Ganpati Jagdale

Scientific Publications by Dr. Ganpati B. Jagdale on insect-parasitic nematodes (EPNs) I. Book Chapters

Tomalak, M., Piggott, S. and Jagdale, G. B. 2005. Glasshouse applications. In: Nematodes As Biocontrol Agents. Grewal, P.S. Ehlers, R.-U., Shapiro-Ilan, D. (eds.). CAB publishing, CAB International, Oxon. Pp 147-166.

II. Research Publications

Jagdale, G.B., Kamoun, S., Grewal, P.S. 2009. Entomopathogenic nematodes induce components of systemic resistance in plants: Biochemical and molecular evidence. Biol. Control.51: 102-109
Hoy, C. W., Grewal, P. S., Lawrence, J. L., Jagdale, G., Acosta, N. 2008. Canonical correspondence analysis demonstrates unique soil conditions for entomopathogenic nematode species compared with other free-living nematode species. Biol. Control. 46: 371-379.
Jagdale, G. B. and Grewal, P. S. 2008. Influence of the entomopathogenic nematode Steinernema carpocapsae infected host cadavers or their extracts on the foliar nematode Aphelenchoides fragariae on Hosta in the greenhouse and laboratory. Biological Control 44: 13-23.
Shabeg, S .B., Jagdale, G. B., Cheng, Z, Hoy, C. W., Miller, S. A. and. Grewal, P. S. 2007. Indicative value of soil nematode food web indices and trophic group abundance in differentiating habitats with a gradient of anthropogenic impact. Environmental Bioindicators 2: 146-160. Jagdale, G. B., Casey, M. L., Grewal, P. S. and Luis Cañas. 2007. Effect of entomopathogenic nematode species, split application and potting medium on the control of the fungus gnat, Bradysia difformis (Diptera : Sciaridae), in the greenhouse at alternating cold and warm temperatures. Biological Control 43: 23-30. 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., Saeb, A. T., Nethi Somasekhar and Grewal, P. S. 2006. Genetic variation and relationships between isolates and species of the entomopathogenic nematode genus Heterorhabditis deciphered through isozyme profiles. Journal of Parasitology 92: 509- 516. Sandhu, S. K., Jagdale, G. B., Hogenhout, S. A. and Grewal, P. S. 2006. Comparative analysis of the expressed genome of the entomopathogenic nematode, Heterorhabditis bacteriophora. Molecular and Biochemical Parasitology 145: 239-244. Grewal, P. S., Susan Bornstein-Forst, S., Burnell, A. M., Glazer, I. and Jagdale, G. B. 2006. Physiological, genetic, and molecular mechanisms of chemoreception, thermobiosis and anhydrobiosis in entomopathogenic nematodes. Biological Control 38: 54- 65. 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 Parasitol 91: 988-994. Jagdale, G. B., Casey, M. L., Grewal, P. S. and Lindquist, R. K. 2004. Application rate and timing, potting medium and host plant on the efficacy of Steinernema feltiae against the fungus gnat, Bradysia coprophila, in floriculture. Biological Control 29: 296-305. 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. Grewal, P. S. and Jagdale, G. B. 2002. Enhanced trehalose accumulation and desiccation survival of entomopathogenic nematodes through cold preacclimation. Biocontrol Science and Technology 12: 533- 545. Jagdale, G. B. and Gordon, R. 1998. Effect of propagation temperatures on temperature tolerances of entomopathogenic nematodes. Fundamental and Applied Nematology 21: 177-183. Jagdale, G. B. and Gordon, R. 1998. Variable expression of isozymes in entomopathogenic nematodes follows laboratory recycling. Fundamental and Applied Nematology 21: 147-155. Jagdale, G. B. and Gordon, R.1997. Effect of temperature on the activities of glucose-6-phosphate dehydrogenase and hexokinase in entomopathogenic nematodes (Nematoda: Steinernematidae). Comparative Biochemistry and Physiology 118A: 1151-1156. Jagdale, G. B. 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 Gordon, R. 1997. Effect of recycling temperature on the infectivity of entomopathogenic nematodes. Canadian Journal of Zoology 75: 2137-2141. Jagdale, G. B., Gordon, R. and Vrain, T. C. 1996. Use of cellulose acetate electrophoresis in the taxonomy of steinernematids (Rhabditida, Nematoda). Journal of Nematology 28: 301-309. Jagdale, G. B. and Gordon, R. 1994. Distribution of catecholamines in the nervous system of a mermithid nematode, Romanomermis culicivorax. Parasitology Research 80: 459-466. Jagdale, G. B. and Gordon, R. 1994. Distribution of FMRF-amide-like peptide in the nervous system of a mermithid nematode, Romanomermis culicivorax. Parasitology Research 80: 467-473. Jagdale, G.B. and Gordon, R. 1994. Role of catecholamines in the reproduction of Romanomermis culicivorax. Journal of Nematology 26: 40-45. Jagdale, G.B. and Gordon, R. 1994. Caudal papillae in Romanomermis culicivorax. Journal of Nematology 26: 235-237.

Symbiotic bacterial genus, Photorhabdus / March 22, 2008 by Ganpati Jagdale

known species of symbiotic bacterial genus Photorhabdus associated with a nematode genus Heterorhabditis. Identification based on colony morphology and molecular techniques

Photorhabdus luminescens (Thomas and Poinar 1979) Boemare et al. 1993
P. temperata
P. luminescens subsp. luminescens subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
P. luminescens subsp. akhurstii subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
P. luminescens subsp. kayaii subsp. nov., Hazir, Stackebrandt, Lang, Schumann, Ehlers & Keskin, 2004
P. luminescens subsp. laumondii subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
P.luminescens subsp. sonorensis, Orozco, Hill & Stock, 2013
P. temperata sp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
P. temperata subsp. temperata subsp. nov., Fischer-Le Saux, Viallard, Brunel, Normand & Boemare, 1999
P. luminescens subsp. thracensis subsp. nov., Hazir, Stackebrandt, Lang, Schumann, Ehlers & Keskin, 2004

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Symbiotic bacterial genus, Xenorhabdus Thomas and Poinar 1979 / March 20, 2008 by Ganpati Jagdale

known species of symbiotic bacterial genus Xenorhabdus Thomas and Poinar 1979 associated with a nematode genus Steinernema. Identification based on colony morphology and molecular techniques

Xenorhabdus beddingii (Akhurst 1986) Akhurst and Boemare 1993
X. bovienii (Akhurst 1983) Akhurst and Boemare 1993
X. budapestensis Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
X. cabanillasii Tailliez, Pagès, Ginibre & Boemare, 2006
X. doucetiae Tailliez, Pagès, Ginibre & Boemare, 2006
X. ehlersii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
X. griffiniae Tailliez, Pagès, Ginibre & Boemare, 2006
X. hominickii Tailliez, Pagès, Ginibre & Boemare, 2006
X. indica Somvanshi, Lang, Ganguly, Swiderski, Saxena, & Stackebrandt 2006
X. innexi Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005
X. japonica Nishimura et al. 1995
X. koppenhoeferi Tailliez, Pagès, Ginibre & Boemare, 2006
X. kozodoii Tailliez, Pagès, Ginibre & Boemare, 2006
X. magdalenensis, Tailliez, Pages, Edgington, Tymo, & Buddie, 2012
X. mauleonii Tailliez, Pagès, Ginibre & Boemare, 2006
X. miraniensis Tailliez, Pagès, Ginibre & Boemare, 2006
X. nematophila (Poinar and Thomas 1965) Thomas and Poinar 1979
X. poinarii (Akhurst 1983) Akhurst and Boemare 1993
X. romanii Tailliez, Pagès, Ginibre & Boemare, 2006
X. stockiae Tailliez, Pagès, Ginibre & Boemare, 2006
X. szentirmaii Lengyel, Lang, Fodor, Szállás, Schumann, Stackebrandt, 2005

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Life cycle of entomopathogenic nematodes (EPNs) / March 16, 2008 by Ganpati Jagdale

Entomopathogenic nematode life cycle

EPNs complete most of their life cycle in insects with an exception of infective juveniles, the only free-living stage found in soil.
Infective juveniles of both Steinernema and Heterorhabditis locate a host and enter through its natural body openings such as mouth, anus or spiracles.
Infective juveniles of Heterorhabditis also enter through the intersegmental members of the host cuticle.
Infective juveniles then actively penetrate through the midgut wall or tracheae into the insect body cavity (hemocoel) containing insect blood (haemolymph).
Once in the body cavity, infective juvenile releases symbiotic bacteria from its intestine 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 insect cadaver.
Heterorhabditid and Steinernematid nematodes differ in their mode of reproduction. For example, in heterorhabditid nematodes, the first generation individuals are produced by self-fertile hermaphrodites (hermaphroditic) but subsequent generation individuals are produced by cross fertilization involving males and females (amphimictic). In Steinernematid nematodes with an exception of one species, all generations are produced by cross fertilization involving males and females (amphimictic).
Depending on availability of food resource, both 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 (from infective juvenile penetration to infective juvenile emergence) is completed within 12- 15 days at room temperature. The optimum temperature for growth and reproduction of nematodes is between 25 and 30⁰C.

Species of the genus Heterorhabditis Poinar, 1976 / March 15, 2008 by Ganpati Jagdale

Known species of Heterorhabditis Poinar, 1976 with a biocontrol potential- Identification based on morphological and molecular techniques

Heterorhabditis amazonensis Andalo, Nguyen, & Moino, 2006
H. argentinensis Stock, 1993
H. atacamensis, Edgington, Buddie, Moore, France, Merino, & Hunt, 2011
H. bacteriophora Poinar, 1976
H. baujardi Phan, Subbotin, Nguyen & Moens, 2003
H. brevicaudis Liu, 1994
H. downesi Stock, Griffin & Burnell, 2002
H. floridensis Nguyen, Gozel, Koppenhofer, & Adams, 2006
H. georgiana Nguyen, Shapiro-Ilan, & Mbata, 2008
Heterorhabditis gerrardi, Plichta, Joyce, Clarke, Waterfield, & Stock, 2009
H. hambletoni (Pereira, 1937) Poinar, 1976
H. hawaiiensis Gardner, Stock & Kaya, 1994
H. heliothidis (Khan, Brooks & Hirschman, 1976) Poinar, Thomas & Hess, 1977
H. hepialius Stock, Strong & Gardner, 1996
H. hoptha (Turco, 1970), Poinar, 1979
H. indica Poinar, Karunakar & David, 1992
H. marelata Liu & Berry, 1996
H. megidis Poinar, Jackson & Klein, 1988
H. mexicana Nguyen, Shapiro-Ilan, Stuart, MCCoy, James & Adams, 2004
H. poinari Kakulia & Mikaia, 1997
H. safricana Malan, Nguyen, deWaal, & Tiedt, 2008
Heterorhabditis sonorensis, Stock, Rivera-Orduno, & Flores-Lara, 2009
H. taysearae Shamseldean, El-Sooud, Abd-Elgawad & Saleh, 1996
H. zealandica Poinar, 1990

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Nematode Information