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Using beneficial nematodes for crop insect pest control

Carol Miles, Caitlin Blethen, Randy Gaugler, and Todd Murray
Nematodes are non-segmented, elongated roundworms that are colorless,
without appendages, and
usually microscopic. There are non-beneficial
and beneficial nematodes.
Non-beneficial nematodes are
also called plant parasitic nematodes and cause damage to crops and other types of plants.
nematodes attack soilborne insect pests, yet are not
harmful to humans, animals, plants, or earthworms,
and can therefore be
used as biological control organisms (Denno et al. 2008). Beneficial nematodes that
cause disease within an insect are referred
to as “entomopathogenic” and have the ability to kill insects
(Figure 1).
ntomopathogenic nematodes from the families Steinernematidae and Heterorhabditidae have
proven to be
the most effective as biological control organisms (Kaya and
Gaugler 1993). They are soil-inhabiting
organisms and can be used effectively to control soilborne insect pests, but are generally not effective when
applied to control insects in the leaf canopy. When considered as a group
of nearly 30 species, each with
its own suite of preferred hosts,
nematodes can be used to control a wide range of
insect pests, including a variety of caterpillars, cutworms, crown borers, grubs, corn root worm,
thrips, fungus gnat, and beetles.
ntomopathogenic nematodes have been
released extensively in crop fields
with negligible effects on nontarget insects
and are regarded as exceptionally safe to the environment.
The keys to success with
ntomopathogenic nematodes are (1) understanding their
life cycles and
functions; (2) matching the correct nematode species with
the pest species; (3) applying them during
appropriate environmental conditions (soil temperature,
soil moisture, sunlight); and (5) applying them
only with compatible pesticides.
Because e
ntomopathogenic nematodes are living organisms, they require
careful handling to survive shipment and
storage as well as appropriate environmental conditions to
survive in the soil after application.
It is unknown whether introduced species of
ntomopathogenic nematodes will overwinter in the Pacific
Northwest and successfully colonize insect hosts in subsequent years. While much of the research regarding
ntomopathogenic nematodes has been conducted outside of the region, the information in this publication
can be successfully applied throughout the Pacific Northwest. Many
species of
nematodes occur naturally in
the region, including Heterorhabditismarelatus. Heterorhabditis marelatus
has been shown
to control some insect crop pests (Berry et al. 1997).
The life cycle of most nematodes includes an egg stage,
four juvenile stages, and an adult stage. The third
juvenile stage of
ntomopathogenic nematodes is referred to
as the “infective juvenile” or “dauer” stage
and is the
only free-living stage (Figure 2). The infective juvenile is capable of surviving in the soil, where it
locates, attacks,
and infects a pest insect (Poinar 1990). Under optimal
conditions, it takes 37 days for
steinernematids and
heterorhabditids to complete one life cycle inside a
host from egg to egg.
Emergence of infective juveniles
from the host requires about 611 days for steinernematids and 1214 days
for heterorhabditids (Kaya and
Koppenhöfer 1999). Figure 3 is a diagram of the life
cycle of
ntomopathogenic nematodes from host infection to
emergence from the host.
Figure 1.
Tens of thousands of infective juveniles of the e
nemato de Steinernema carpocapsae
spilling out of a Galleria wax moth larva in search of new hosts (photo provided by Randy Gaugler).
Figure 2. An infective juvenile entomopathogenic nematode exhibiting the “J”-shaped resting position
(photo provided by David Shapiro-Ilan).
Figure 3.
Diagram of the generalized life cycle of entomopathogenic nematodes (image provided by David Shapiro-
An understanding of host-finding strategies
will help you properly match
ntomopathogenic nematode
to pest insects to ensure infection and control (Gaugler
1999). Only
ntomopathogenic nematodes
in the infective juvenile stage will
survive in the soil and find and penetrate insect
pests. Infective juvenile
ntomopathogenic nematodes locate
their hosts in soil by means of two strategiesambushing and cruising
(Gaugler et al. 1989). Ambusher
species include Steinernema carpocapsae and S. scapterisici; cruisers
include Heterorhabditis bacteriophora and S. glaseri. S. riobravis and S. feltiae
do a bit of both
ambushing and cruising (Campbell
and Gaugler 1997).
ntomopathogenic nematodes that use the ambushing
strategy tend to remain
stationary at or near the
soil surface and locate host insects by direct contact (Campbell et al. 1996).
An ambusher searches
by standing on its tail so that most of its body is in the air, referred to as
“nictation.” The nictating
nematode attaches to and attacks passing insect
hosts. Ambusher
ntomopathogenic nematodes most effectively
control insect pests that are highly mobile at the
surface, such as cutworms, armyworms, and
mole crickets.
Entomopathogenic nematodes that use the cruising strategy
are highly mobile and able to
move throughout the soil profile. Cruisers locate their host by
sensing carbon dioxide or other
volatiles released
by the host. Cruiser
ntomopathogenic nematodes are most effective
sedentary and slow-moving insect pests at
various soil depths, such as white grubs and root
Generally, several
ntomopathogenic nematodes will infect a single insect
host. The infective juvenile
nematodes penetrate the
insect’s body cavity, usually through natural body
openings such as the mouth,
anus, genital pore, or breathing
pore (spiracle) (Figure 4). Heterorhabditids can break the outer
cuticle of
the insect using a dorsal “tooth” or hook.
Once inside the body cavity of the host, the infective
release bacteria that live symbiotically
within the
ntomopathogenic nematode’s gut but do not harm
the nematode. The nematode-bacterium
relationship is highly specific: only Xenorhabdus spp.
bacteria co-
exist with steinernematids, and only Photorhabdus bacteria co-exist with heterorhabditids. Once
into the host, the bacteria multiply quickly
and under optimal conditions cause the host to die
within 24
to 48 hours.
ntomopathogenic nematodes feed on both bacteria they release and
host insect tissue. After a few days
inside the host,
ntomopathogenic nematodes mature to the adult stage. These
nematodes produce hundreds of thousands of
new juveniles that may undergo several
life cycles within
a single host (Figure 5). When the host has
been consumed, the infective juveniles, armed with a fresh
supply of bacteria, emerge from the empty
shell of the host, move into the soil, and begin the
search for a
new host. A protective exterior cuticle
surrounds the infective juvenile, protecting it from
the environment
and predators. Under ideal
conditions, steinernematids emerge 611 days after
initial infection and
heterorhabditids emerge 1214
days after initial infection (Kaya and Koppenhöfer
1999). The duration of
infective juvenile survival in soil
is unknown because they can become prey
to invertebrates and
Figure 4. Infective juvenile entomopathogenic nematodes entering a host through the insects genital pore
(photo provided by Randy Gaugler).
Figure 5.
Entomopathogenic nematodes produce Hhundreds of thousands of
new juveniles
entomopathogenic nematodes that may undergo several
life cycles within a single host insect
provided by Randy Gaugler)
Over 30 species of
ntomopathogenic nematodes have been
identified. Eight species have been
worldwide and seven are currently available in the
United States: Steinernema
carpocapsae, S. feltiae,
S. glaseri, S. riobravis, Heterorhabditis bacteriophora,
H. megidis, and H. marelatus.
The eighth species, S.scapterisci, although once commercialized, is no longer
available (Kaya and
Koppenhöfer 1999).
The different species of
ntomopathogenic nematodes vary in
the range of insects they attack,
environmental needs,
and stability in commercial products (Gaugler 1999).
A given species of
ntomopathogenic nematode may also control a particular pest more effectively than another
Therefore, the insect pest must be identified
before choosing the
ntomopathogenic nematode species most
appropriate for biological control. Table 1 is a general guide
of available
ntomopathogenic nematodes
and the pests they
have been shown to control. Table 2 provides a description
of each species of
commercially available
BIOLOGICAL CONTROL ORGANISMS1 (From Shapiro-Ilan and Gaugler 2010)
Crop(s) Targeted
Pest Common Name
Pest Scientific Name
Efficacious Nematodes2
Artichoke plume moth
Platyptilia carduidactyla
Lepidoptera: Noctuidae
Sc, Sf, Sr
Banana moth
Opogona sachari
Hb, Sc
Banana root borer
Cosmopolites sordidus
Sc, Sf, Sg
Sphenophorus spp. (Coleoptera:
Turf, vegetables
Black cutworm
Agrotis ipsilon
Berries, ornamentals
Black vine weevil
Otiorhynchus sulcatus
Hb, Hd, Hm, Hmeg, Sc, Sg
Fruit trees, ornamentals
Synanthedon spp. and other sesiids
Hb, Sc, Sf
Home yard, turf
Cat flea
Ctenocephalides felis
Citrus, ornamentals
Citrus root weevil
Pachnaeus spp. (Coleoptera:
Sr, Hb
Pome fruit
Codling moth
Cydia pomonella
Sc, Sf
Corn earworm
Helicoverpa zea
Sc, Sf, Sr
Corn rootworm
Diabrotica spp.
Hb, Sc
Cranberry girdler
Chrysoteuchia topiaria
Crane fly
Diptera: Tipulidae
Citrus, ornamentals
Diaprepes root weevil
Diaprepes abbreviatus
Hb, Sr
Fungus gnat
Diptera: Sciaridae
Sf, Hb
Grape root borer
Vitacea polistiformis
Hz, Hb
Iris borer
Macronoctua onusta
Hb, Sc
Forest plantings
Large pine weevil
Hylobius albietis
Hd, Sc
Vegetables, ornamentals
Liriomyza spp. (Diptera: Agromyzidae)
Sc, Sf
Mole cricket
Scapteriscus spp.
Sc, Sr, Sscap
Nut and fruit trees
Navel orangeworm
Amyelois transitella
Fruit trees
Plum curculio
Conotrachelus nenuphar
Turf, ornamentals
Scarab grub3
Coleoptera: Scarabaeidae
Hb, Sc, Sg, Ss, Hz
Shore fly
Scatella spp.
Sc, Sf
Strawberry root weevil
Otiorhynchus ovatus
Bee hives
Small hive beetle
Aethina tumida
Hi, Sr
Sweet potato
Sweetpotato weevil
Cylas formicarius
Hb, Sc, Sf
1 ANematodes listed provided at leastt least one scientific study reported 75% suppression of
these pests using the nematodes indicated in field or greenhouse experiments.
2 Nematode species are abbreviated as follows: Hb = Heterorhabditis bacteriophora, Hd = H.
downesi, Hi = H. indica, Hm = H. marelata, Hmeg = H. megidis, Hz = H. zealandica, Sc =
Steinernema carpocapsae, Sf = S. feltiae, Sg = S. glaseri, Sk = S. kushidai, Sr = S. riobrave, Sscap
= S. scapterisci, Ss = S. scarabaei.
3 Efficacy against various pest species within this group varies among nematode species.
1999 and Berry 2000).
Steinernema carpocapsae. An ambusher type, it is most effective against highly mobile surface
insects such as webworms, cutworms, armyworms, girdlers, and wood borers. Most
effective at soil surface temperatures between 70°F and 85°F. Can be formulated in a
partially desiccated state in clay granules to provide several months of room-temperature
shelf life.
S. feltiae. Combines ambusher and cruiser strategies and attacks immature fly larvae (dipterous
insects), including mushroom flies, fungus gnats, and crane flies. Maintains infectivity at low
soil temperatures, even below 50°F. Has relatively low stability in formulation and a short
shelf life.
S. glaseri. A large cruiser type that attacks white grubs and other beetle larvae, particularly scarabs.
Expensive and difficult to produce and manage due to its tendency to lose its bacterial
symbiont. Highly active and robust infective juveniles are difficult to contain within
S. riobravis. Combines ambusher and cruiser strategies, and attacks corn earworms, citrus root
weevils, pink bollworms, and mole crickets. Isolated from the Rio Grande Valley of Texas.
Maintains infectivity at soil temperatures above 95°F and in semi-arid conditions.
Heterorhabditis bacteriophora. A cruiser-type nematode that attacks caterpillar and beetle
larvae, including root weevils, particularly black vine weevil. Most effective in warm
temperatures (above 68°F). Infective juveniles persist only a few days in the soil. Has poor
stability in formulation and a short shelf life.
H. marelatus. A cruiser-type nematode that attacks beetle larvae , including white grubs and
root weevils. Isolated from the Oregon coastal region. Active at cool soil temperatures (50
H. megidis. A large cruiser-type nematode that has been effective in controlling black vine weevil
larvae. Has not been widely researched or tested for insect control. Isolated in Ohio, researched
and developed in Europe, and now available in the United States. Tends to have poor formulation
stability and a short shelf life.
Obtaining E
Perhaps the biggest challenge to the use of
nematodes as effective biological control
is the variable quantity and quality of nematodes in
commercial products (Gaugler et al. 2000).
ntomopathogenic nematodes
are cultured on a large scale in laboratories and are
available from many
commercial suppliers in North
America and Europe. In past assessments of cottage
industry commercial
products, most contained lower
numbers of
ntomopathogenic nematodes than the suppliers claimed. In
addition, in some cases the species of
ntomopathogenic nematodes in
the product were mixed and
therefore inconsistent with
the product label. The industry has made progress,
however, in increasing the
quality of its products. See Table 3 for lists of commercial suppliers compiled by university and state
government professionals.
ntomopathogenic nematodes can also be purchased
through gardening mail-
order catalogs and at some
local agricultural and nursery supply stores.
Commercial Sources of Insect Parasitic Nematodes. 2000. Compiled by P. Grewal and K. Power.
Ohio State University.
Commercial Producers and Suppliers of Nematodes. 2010. Compiled by D.I. Shapiro-Ilan and
R. Gaugler. Cornell University.
Vendors of Beneficial Organisms in North America. 2010. Compiled by J. White and
D. Johnson. University of Kentucky College of Agriculture.
Suppliers of Beneficial Organisms in North America. 1997. Compiled by C.D. Hunter.
California Environmental Protection Agency, Department of Pesticide Regulation,
Environmental Monitoring and Pest Management Branch.
Shelf Life
In general,
ntomopathogenic nematodes do not have a long
shelf life. Many microbial insecticides,
Bacillus thuringiensis, have a resting stage facilitating
long-term storage. The infective juvenile
ntomopathogenic nematode
stage is not a resting stage; juveniles are metabolically
active and use energy
reserves while in formulation
(Lewis 1999). For this reason, it is advisable to order
nematodes only 34 days prior to application.
ntomopathogenic nematodes should be shipped by
overnight delivery in their
infective juvenile stage and used
within 12 days after arrival.
Examine the
ntomopathogenic nematodes upon receipt to make sure
they arrived alive. The shipment
container should not
feel warm or hot. Open the container and check the
color and odor of the
nematodes. To the naked eye,
the nematodes on a sponge formulation will appear as
a light tan or gray
paste, while nematodes in vermiculite or liquid suspension will not be discernible
from the carrier
material. The container should have
a mild odor; if there is a strong smell, like ammonia,
then it is likely
the nematodes are dead. If the formulation is a sponge or vermiculite, remove
a tiny portion of the
product with tweezers and place
in a teaspoon of cool water (approximately 60ºF) for
six hours. If the
formulation is a liquid suspension, swirl the liquid to ensure distribution of the nematodes and remove a
small droplet (about 0.05 ml).
Place the soaked nematode sample (from the sponge
or vermiculite) or the droplet from the liquid suspen-
sion on a slide or in a small, clear glass bowl. View
the samples with a hand lens (15X) or microscope.
ntomopathogenic nematodes will be mobile and have
a bend to their shape. S. carpocapsae has a
“J” shape and will move only when prodded with a
pin or needle (Figure 2). All other
nematodes will move in an
“S” pattern (Lewis 1999). If the nematodes are straight
and not moving, it is
likely they are dead. A mortality rate of 10% should be expected. If more than
20% of the nematodes are
dead, inform your supplier
ntomopathogenic nematodes should be stored in their shipment containers under refrigeration until
ready for use. The
storage life of
ntomopathogenic nematodes is species- and formulation-dependent.
Specific storage instructions will be included with the
ntomopathogenic nematode shipment and should
followed. Table 4 is a summary of storage times for
ntomopathogenic nematodes in different
formulations. Storing
nematodes under refrigeration will increase their shelf
life, but their infectivity will
still decrease the longer
they are in storage. When the storage life has expired,
expect 70100% mortality of
the nematodes (Grewal
NEMATODES (adapted from Grewal 1999b).
Storage Duration
Nematode Species
Room Temperature
Liq uid concentrate
S. carpocapsae
S. riobravis
56 days
34 days
1215 days
79 days
S. carpocapsae
23 months
H. bacteriophora
12 months
Ver miculite
S. feltiae
H. megidis
45 months
23 months
Alg inate gels
S. carpocapsae
S. feltiae
34 months
1 month
69 months
45 months
Flowable gels
S. carpocapsae
S. glaseri
35 months
Water dispersible
S. carpocapsae
S. feltiae
S. riobravis
2 months
23 months
912 months
57 months
45 months
Wettable p ow der
S. carpocapsae
S. feltiae
H. megidis
23 months
23 months
68 months
56 months
45 months
Nematode w oo l
H. bacteriophora
21 days
Soil Conditions
ntomopathogenic nematodes can die if they are applied to
soils that are too dry, too hot, or too cold,
or if they
are exposed to ultraviolet (UV) light from the sun.
Nematodes live in the water-filled spaces, or
between soil particles. They need water to move and
successfully locate a host, and oxygen to
survive. Heavy clay soils hold water
well, but may contain too little oxygen, and the small
pore space may
restrict nematode movement. Sandy
soils must be irrigated to maintain the water-filled pores. If applying
ntomopathogenic nematodes after an extended dry period, break the crust of the dry soil with a rake
or harrow and irrigate the soil before the application, preferably to a depth of 46 inches. Remove
plant debris before the application, as it will prevent the nematodes from reaching the soil surface.
Soil temperatures between 77
F and 82
F are ideal
for applying all
ntomopathogenic nematode species.
In general, soil
temperatures greater than 85
F can decrease the
efficacy of some nematode species, while
soil temperatures less than 50
F can immobilize others at
the soil surface, causing them to be exposed to
light that can kill them. The range of soil temperatures that nematode species can survive and infect
host insects does vary, however. For example, S.
feltiae can be effective at 57
F, while S. riobravis
be effective at 95
ntomopathogenic nematodes should be applied late in the day or on
a cool, overcast day when light
and temperatures
are low. Apply them to moist soil following either
a rainfall or irrigation, and lightly
irrigate afterward. This washes them
into the soil and decreases soil surface temperatures.
Do not over-
irrigate, as saturated soil will impede
nematode activity due to lack of oxygen.
Preparing for Application
ntomopathogenic nematodes should be prepared for field application no earlier
than one hour ahead of
time. If nematodes
are in a liquid suspension, shake the shipment container well and pour the liquid into
the application
container (e.g., tank, backpack sprayer, or watering
can). Rinse the shipment container
twice with cool
water (approximately 60
F), and pour the rinse water
into the application container. If
nematodes are on a
sponge, soak the sponge in one gallon of cool water
for 10 minutes, then pour the
water into the application container. Rinse the sponge several times, pouring
the rinse water into the
application container after
each rinse. If nematodes are in vermiculite, add the
mixture directly to water in the
application container and stir until dispersed. Once
the nematodes have
been mixed with water, agitate
the mixture every five minutes to keep the nematodes
in suspension and
supplied with oxygen.
Application Equipment
Read the product label for specific application instructions.
ntomopathogenic nematodes that are
formulated with vermiculite
may be best applied as a granular product. Other formulations can be applied
using standard liquid pesticide,
fertilizer, and irrigation equipment with pressures of
up to 300 PSI.
Electrostatic, fan, pressurized, and mist
sprayers can be used. If tanks are agitated through
sparging (recirculation of the spray mix), or if the temperature in the tank rises above 86
F, the
will be damaged. Irrigation systems may
also be used for applying most species; however, high-pressure
recycling pumping systems are not good
delivery systems (Shetlar 1999). An excellent overview of
sprayer equipment is provided in the Private
Applicator Pesticide Education Manual, EM020 (Ramsay et al.
Remove all screens smaller than 50-mesh from your
spray or irrigation equipment to allow nematodes
pass through the system. Check spray nozzle orifices
for clogging during application. Direct spray
at the soil to maximize the number of nematodes being
applied directly to the soil. A large spray
volume is
ideal. Volumes of 26 gallons of water per 1,000 square
feet (87260 gallons per acre) are
recommended on
most nematode labels (Shetlar 1999). The water in
the spray will wash the nematodes
from plant surfaces into the soil. Lower volume spray applications of 0.51.0 gallon per 1,000 square
feet (2045 gallons per
acre) can be used if the area or field is irrigated prior
to and immediately
ntomopathogenic nematode application.
Overhead irrigation following nematode
will wash the nematodes from plant surfaces into the
soil. If the spray droplets are allowed
to dry prior to
this irrigation, the nematodes will be exposed to UV
light and die while still on the plant
Applying nematodes during a rainfall will also ensure
that nematodes reach the soil surface.
Use equipment that is clean and free of pesticide
residues. Also, do not mix
ntomopathogenic nematodes
with nitrogen
fertilizers, particularly urea (Grewal 2000). Although
there is evidence that nematodes are
tolerant to many
herbicides and fungicides, they are sensitive to certain
insecticides and nematicides. Refer
to the nematode
product label for specific listings of chemicals that are lethal to
nematodes. To check that live
nematodes are being applied to the soil, set pans or
containers on the soil
surface prior to application.
Immediately after application, use a hand lens (15X)
or a microscope to check
the liquid in the pans or
containers for live, moving nematodes.
Application Rates
Before applying any biological control, including
ntomopathogenic nematodes, read the product label
for specific application instructions. A broadcast application rate of 1
billion nematodes per acre is
generally recommended
to control most soil insects. For smaller areas, the
recommended application
rate is 250,000 nematodes
per square meter. If nematodes are banded (applied in a band beside the crop
row), a lower rate may be
applied. Research at the University of Florida has
demonstrated that a rate of
up to 200 million nematodes per acre applied in a band provided effective
control of root weevil in citrus
orchards (Duncan et
al. 1999). More research is needed to determine
specific rate responses for each
species of
nematode in various cropping systems to control
specific pests. Calibrate
equipment to ensure appropriate application
rates. An excellent overview of sprayer calibration is provided
in the Private Applicator Pesticide Education Manual, EM020 (Ramsay et al. 2009).
Evaluating nematode applications
It can be difficult to be sure if the e
nematodes reached the soil and the target pests, as it
is very laborious to recover the cadavers of the insects they have killed. There are two simple tests that
can be used to assess the efficacy of all e
nematode species. Both tests employ Gallaria
mellonella waxworms (Berry 2007), which are the caterpillar stage of a waxmoth species that are
extremely susceptible to e
nematode infection. Galleria waxworms are readily available
at fishing bait and pet supply stores.
For the first test, place 23 Gallaria waxworms in a tea strainer and bury the strainer 4 inches deep in the
soil. You can bury the Gallaria waxworms either just before you apply the e
or anytime afterwards. It is best to place several baited strainers in the area where nematodes are being
appliedeither 34 strainers for a garden area or approximately 10 strainers per acre. Remove the
strainers from the soil after 2 days, rinse the waxworms with distilled water, and store them on moistened
filter paper or thick paper towel in a dark location at room temperature. Check the waxworms regularly
over the next 710 days to look for nematode infection. Infected waxworms will usually change color;
Steinernematid-infected waxworms will turn yellow, tan, or brown, while Heterorhabditid-infected
waxworms will turn pink or purple. If the waxworms turn black, they were likely killed by other means.
For the second test, collect e
nematode-treated soil from the treated area at least one day
after nematodes have been applied. Collect at least 10 soil samples from a garden area and 20 soil
samples per acre. Each soil sample should be approximately ¼ cup from a depth of 4 inches. Mix the soil
together, place ¼ cup into a wax cup, and place a Galleria waxworm on top of the soil. Use 23 cups for
a garden area or approximately 10 cups per acre. Place the cups in a dark area at room temperature for 2
days. Rinse, store, and evaluate the waxworms as described above.
Pesticide Safety
ntomopathogenic nematodes, like all biological control
organisms, are considered pesticides. However,
nematodes are exempted from federal pesticide registration requirements under
40CFR152.20(a) because they belong to a group of specific biocontrol agents. Questions regarding
pesticide registration should
be directed to your state’s Department of Agriculture
Pesticide Division.
Washington State Department of Agriculture
Pesticide Management Division
PO Box 42589, Olympia, WA 98504-2589
Phone: 1-877-301-4555, Fax: (360) 902-2093,
Oregon State Department of Agriculture
Pesticides Division
635 Capitol Street NE, Salem, OR 97301-2532
Phone: (503) 986-4635, Fax: (503) 986-4735,
Idaho State Department of Agriculture Division
of Agricultural Resources Pesticide Division
PO Box 790, Boise, ID 83701-0790
Phone: (208) 332-8500, Fax: (208) 334-2170, Email:
The environmental benefits of using
ntomopathogenic nematodes include
no concern with re-entry
times, residues, groundwater
contamination, or pollinators. Any pesticide, including biological controls,
applied to certified organic
farms must be approved by the organic certification
program. A list of
approved organic materials is
available from your state’s Department of Agriculture
Organic Program.
Washington State Department of Agriculture
Organic Food Program
PO Box 42560, Olympia, WA 98504-2560
Phone: (360) 902-1805, Fax: (360) 902-2087,
Oregon State Department of Agriculture Food
Safety Division
635 Capitol Street NE, Salem, OR 97031-2532
Phone: (503) 986-4550, Fax: (503) 986-4747,
Idaho State Department of Agriculture Organic
PO Box 790, Boise, ID 83701
Phone: (208) 332-8675, Fax: (208) 334-2170,
More Information about E
Insect Parasitic NematodesOhio State University
Provides a comprehensive bibliography of research
literature that permits quick access to all published
particularly field trials, for any target insect.
Includes an electronic expert panel of
authorities who will respond to questions.
Center for Vector BiologyRutgers University
Emphasizes a balance of basic and applied research
to achieve fundamental understandings of the insect-
nematode host-parasite relationship to increase
efficacy and integration of entomopathogenic nematodes in
pest management systems. Includes a slide
collection, list of publications, and summaries of
projects on
production, behavioral
and developmental ecology, and genetic engineering of entomopathogenic
Plant and Insect Parasitic NematodesUniversity of Nebraska
Includes discussion of the
species Steinernema and Heterorhabditis, including
culture, storage, transport,
and images. Includes an electronic copy of the paper.
Biological Control News
University of Wisconsin
Provides back issues of the newsletter Biological Control News. Also includes an
alphabetical index of
biological control-related
articles. For example, clicking on the N button
will provide a list of topics
on nabid bug,
National Academy of Science Report, Nealiotus
curculionis, and nematodes. Click on any
of the
topics for a complete article listing.
Berry, R. 2007. Application and Evaluation of Entomopathogens for Control of Pest Insects in Mint. In
Field Manual
of Techniques in Invertebrate Pathology: Application and Evaluation of Pathogens for Control of Insects and Other
Invertebrate Pests
. 2nd ed. Edited by L.A. Lacey and H.K. Kaya, pp. 599-608.Dordrecht, The Netherlands: Springer.
Berry, R. 2000. Control of Root Weevils in Nursery Crops with Entomopathogenic Nematodes. In
Proceedings: Beneficial Nematode, Application in
Greenhouse, Nursery and Small-Fruit Operations
, pp. 2229.
Edited by P. Gothro, Oregon State University.
Berry, R.E., J. Liu, and E. Groth. 1997. Efficacy and Persistence of Heterorhabditis marelatus (Rhabditida:
Heterorhabditidae) Against Root Weevils (Coleoptera: Curculionidea) in Strawberry.
Environmental Entomology
26(2): 465470.
Campbell J., and R. Gaugler. 1997. Inter-specific Variation in Entomopathogenic Nematode Foraging Strategy:
Dichotomy or Variation Along a Continuum?
Fundamental & Applied Nematology
Campbell, J., E. Lewis, F. Yoder, and R. Gaugler. 1996. Entomopathogenic Nematode Spatial Distribution in
113: 473482.
Denno, R.F., D.S. Gruner, and I. Kaplan. 2008. Potential for Entomopathogenic Nematodes in Biological Control:
A Meta-Analytical Synthesis and Insights from Trophic Cascade Theory.
Journal of Nematology
40(2): 61-72.
Duncan, L. W., D. I. Shapiro, C. W. McCoy, and J. H. Graham. 1999. Entomopathogenic Nematodes as a Component
of Citrus Root Weevil IPM. In
Proceedings: Optimal Use of Insecticidal Nematodes in Pest
pp. 7990. Edited by S. Polavarapu, Rutgers University.
Gaugler, R., J. Campbell, and T. McGuire. 1989. Selection for Host Finding in Steinernema feltiae.
Journal of
Invertebrate Pathology
54: 363372.
Gaugler, R. 1999. Matching Nematode and Insect to Achieve Optimal Field Performance. In
Proceedings: Optimal Use of Insecticidal Nematodes
in Pest Management,
pp. 914. Edited by S. Polavarapu,
Rutgers University.
Gaugler, R., P. Grewal, H.K. Kaya, and D. Smith-Fiola. 2000. Quality Assessment of Commercially Produced
Entomopathogenic Nematodes.
Biological Control
17: 100109.
Kaya, H.K., and R. Gaugler. 1993. Entomopathogenic Nematodes.
Annual Review of Entomology
38: 181206.
Kaya, H.K., and A.M. Koppenhöfer. 1999. Biology and Ecology of Insecticidal Nematodes. In
Optimal Use of Insecticidal Nematodes
in Pest Management
, pp. 18. Edited by S. Polavarapu, Rutgers University.
Lewis, E.E. 1999. Handling, Transport, and Storage of Insecticidal Nematodes. In
Workshop Proceedings:
Use of Insecticidal Nematodes in Pest Management
, pp. 2530. Edited by S. Polavarapu, Rutgers University.
Poinar, G.O. 1990. Biology and Taxonomy of Steinernematidae and Heterorhabditidae. In:
Nematodes in Biological Control
, pp. 2361. Edited by R. Gaugler and H.K. Kaya. Boca Raton, FL: CRC Press.
Polavarapu, S. 1999. Insecticidal Nematodes for Cranberry Pest Management. In
Workshop Proceedings: Optimal
Use of Insecticidal Nematodes in Pest
, pp. 7990. Edited by S. Polavarapu, Rutgers University.
Ramsay, C., R. Hines, and C. Foss. 2009. Private Applicator Pesticide Education Manual: A Guide to Safe Use and
Washington State University Extension
Shapiro-Ilan, D.I., and R. Gaugler. 2010. Nematodes: Rhabditida: Steinernematidae & Heterorhabditidae. In:
Biological Control:
A Guide to Natural Enemies in North America
. Edited by A. Shelton, Cornell University.
Shetlar, D.J. 1999. Application Methods in Different Cropping Systems. In
Workshop Proceedings: Optimal
Use of
Insecticidal Nematodes in Pest Management
, pp. 3136. Edited by S. Polavarapu, Rutgers University.
About the Authors
Carol Miles, Vegetable Specialist, Department of Horticulture and Landscape Architecture, Washington
State University, Mount Vernon Northwest Washington Research and Extension Center.
Caitlin Blethen, former assistant to Carol Miles, Vegetable Horticulture, Washington State University.
Randy Gaugler, Entomologist, Director for the Center for Vector Biology, Rutgers University.
Todd Murray, Extension Educator, Washington State University, Skamania County.
... This procedure was repeated thrice and the average number of live and dead IJs was determined for each sponge type. Live IJs moved rapidly or had a Jshaped position (Miles et al., 2000). Immobile and straight were probed to confirm if they were alive or dead. ...
Sponges are one of the cheapest and most suitable substrates used to formulate and/or store the infective juveniles (IJs) of entomopathogenic nematodes (EPNs). Our study investigated the survival and infectivity of the IJs on five different sponges compared to that in an aqueous suspension (control). The sponges were Oasis® floral, Nanosponge, ScotchbriteTM, or Lysol® and natural sea sponge. EPN species tested were Heterorhabditis bacteriophora, Steinernema carpocapsae and S. feltiae. The recovery efficiency of the IJs from sponges was initially assessed. Subsequently, IJs were stored in the sponges and placed in plastic bags or Falcon tubes and incubated at 10o or 27℃ for 8 months or 11 weeks, respectively. IJ survival and infectivity were monitored monthly for the 10°C and weekly for 27℃ in these sponge types. The IJs were recovered from the sponges, and their survival was based on observing their movement under a dissecting microscope, and infectivity was based on larval mortality in Galleria mellonella. Recovery efficiency of IJs was best for the Oasis floral sponge for all nematode species ranging between 83 and 91%. The survival and infectivity of stored IJs in all sponge types and control for both 10o and 27℃ gradually decreased over time. IJs stored in Scotchbrite, Lysol, and Nanosponge had the best survival and infectivity, whereas Oasis floral and natural sea sponges showed the poorest results. After 8 months at 10℃ in plastic bags, the survival ratio of all IJs in these three sponges (Scotchbrite, Lysol, and Nanosponge) was approximately 55%. IJs in Scotchbrite and Nanosponge were also able to survive and retain their infectivity at 27℃ for 3 months. IJs stored in Falcon tubes had survival that ranged from 26 to 53% at 27°C and 55 to 77% at 10°C. H. bacteriophora IJs lost their infectivity when stored at 27°C after 10 weeks. However, S. carpocapsae and S. feltiae exhibited 85% infectivity when stored in Scotchbrite and 50% in Nanosponge, respectively. Overall, sponges made from polyurethane (Scotchbrite) followed by melamine (Nanosponge) and cellulose (Lysol) are recommended for long-term nematode storage and transportation of nematode samples. However, Oasis floral sponge may be preferred for short-term IJ formulation for field applications because of easier recovery of IJs.
Myiasis caused by Lucilia sericata (Diptera: Calliphoridae) is widely distributed throughout the world and affects both humans and animals. In addition, L. sericata larvae and adults may play a role in spreading causal agents of mycobacterial infections. Therefore, it is important to establish new and safe alternative methods of controlling this blowfly. The insecticidal effectiveness of four commercially available essential oils [lettuce (Lactuca sativa), chamomile (Matricaria chamomilla), anise (Pimpinella anisum), rosemary (Rosmarinus officinalis)] against third larval instars of L. sericata was evaluated. The effects of sublethal concentrations of these oils on pupation rates, adult emergences, sex ratios, and morphological anomalies were also determined. The oils were highly toxic to L. sericata larvae, with median lethal concentrations (LC(50) ) of 0.57%, 0.85%, 2.74%, and 6.77% for lettuce, chamomile, anise, and rosemary oils, respectively. Pupation rates were markedly decreased after treatment with 8% lettuce oil, and adult emergence was suppressed by 2% lettuce and chamomile oils. Morphological abnormalities were recorded after treatment with all tested oils, and lettuce was the major cause of deformation. There was a predominance of males over females (4 : 1) after treatment with lower concentrations of chamomile and rosemary; such a skew toward males would lead to a population decline. The four tested oils are inexpensive and may represent new botanical insecticides for controlling blowflies.
Full-text available
The monogeneric nematode families Steinernematidae (Steinernema) and Heterorhabditidae (Heterorhabditis), mutually associated with the pathogenic bacteria Xenorhabdus, are similar in their actions. The free-living, non-feeding infective juveniles possess attributes of both insect parasitoids or predators (they have chemoreceptors and are motile) and microbial pathogens (virulence, high reproductive capacity, numerical but no functional response to host population changes). Their potential as agents of biological control are accentuated by their dispersability using spray equipment, their compatibility with many pesticides, and their amenability to genetic selection. Major sections of this review are on: taxonomy; biology of the nematode-bacterium complex; host range (insects are killed so rapidly that highly adapted host-parasite relationships do not form); behaviour; ecology (including dispersal and host finding, survival, interspecific competition, and recycling and epizootiology); genetics; commercialisation; and efficacy. -P.J.Jarvis
Full-text available
The quality of the most widely commercialized entomopathogenic nematodes, Steinernema carpocapsae (Weiser) and Heterorhabditis bacteriophora Poinar, was assessed from 30 shipments from six United States suppliers mailed to three locations. These suppliers comprise a cottage industry aimed in large part at a mail-order market. Most companies were accessible and reliably shipped pure populations of the correct species on time, in sturdy containers, and often with superb accompanying instructions. Nematodes were received in satisfactory condition with acceptable levels of viability. Consistency, however, was a problem, with each supplier having one or more weak spots to bolster. Most shipments did not contain the expected nematode quantity, and one shipment had no nematodes. Pathogenicity of several products against Galleria mellonella (L.) larvae was not equivalent to our controls. H. bacteriophora was not always available when ordered. A few products contained mixed populations of both nematode species. Application rate recommendations provided by some suppliers appeared unsound. We conclude that (1) the entomopathogenic nematode cottage industry lacks rigorous quality control, (2) self-regulation is problematic without feedback on quality, and (3) consumers are rarely able to provide this feedback. Improved reliability by the nematode industry will most likely be achieved via industry-generated agreement on standards for quality.
Full-text available
A hybridized Foundation population of the entomopathogenic nematode Steinernema feltiae was bidirectionally selected for enhanced and diminished host-finding ability. While there was no response to selection for diminished host-finding, 13 rounds of selection for enhancement produced a 20- to 27-fold increase. Moreover, the proportion of infectives initiating positive chemotaxis was increased from less than one-third to more than 80%. Nematodes failing to migrate out of the inoculation zone declined from 33 to 8% after 6 rounds of selection. Relaxation of selection pressure produced a gradual decrease in host-finding. This regression, coupled with the high realized heritability for enhanced host-finding (0.64), suggests that wild-type populations take a passive approach to host-finding. Because for inundative biological control it is desirable that infective stages quickly initiate host-seeking movements, improved host-finding may result in improved field efficacy.
Full-text available
L'aptitude de six espèces de nématodes entomopathogènes ayant des comportements différents dans la recherche d'hôtes ayant eux-mêmes des taux de mobilité variables (#Galleria mellonella$ avec ou sans restriction de mouvement) a été évaluée. #Steinernema carpocapsae$ et #S. scapterisci$ ont tendance à se tenir droit sur leur queue sans se déplacer pendant des périodes de temps assez prolongées mais avec des mouvements de pendulation. Ces espèces utilisent une stratégie d'embuscade et sont les plus efficaces pour trouver les larves se déplaçant sans restriction de mouvement. #Heterorhabditis bacteriophora$ et #S. glaseri$ ne montrent pas de mouvement de pendulation et ont été les plus efficaces pour trouver les larves à déplacement restreint, ce qui est typique d'espèces cherchant leur hôte en se déplaçant. Une autre espèce sans mouvement de pendulation, #S. feltiae$, et une espèce à pendulation peu fréquente, #S. riobravis$, peuvent atteindre les deux types d'hôtes, ce qui suggère qu'elles ont un comportement intermédiaire de recherche d'hôtes. La stratégie intermédiaire de #S. feltiae$ pourrait être due à sa capacité à élever de plus de 30% son corps au-dessus du support (mouvement de pendulation), et ce, plus fréquemment que les autres espèces sans mouvement pendulaire. (Résumé d'auteur)
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Understanding the temporal and spatial distribution of entomopathogenic nematodes is essential for determining the role of these insect parasites in soil communities and ultimately for their use in suppression of pest insect populations. We measured the vertical and horizontal distribution of endemic populations of entomopathogenic nematodes (Steinernema carpocapsae and Heterorhabditis bacteriophora) in turfgrass. Vertical distribution was determined by taking soil cores every 3 h from 05.00 to 23.00 h, over 4 days, and dividing the cores into 8, 1 cm deep sections. Steinernema carpocapsae was recovered primarily near the soil surface: 50% of positive sections were recovered in the thatch or first 1 cm of soil. S. carpocapsae recovery was lower during the middle of the day and none were recovered in the upper section. H. bacteriophora was recovered uniformly throughout the top 8 cm of soil and its vertical distribution did not change over the course of the day. Horizontal distribution was measured as the number of nematodes recovered from cores taken from 12 randomly selected 0.3 x 0.8 m sections from within four 15.3 x 15.3 m plots. Samples were collected biweekly over a 9-month period. H. bacteriophora had a patchier distribution than S. carpocapsae and both nematode species had more patchy distributions then their potential hosts. Our results support the hypothesis that these two species of nematode utilize different foraging strategies; S. carpocapsae primarily a surface adapted ambusher and H. bacteriophora as a cruise forager.
Peppermint (Mentha piperita), native spearmint (M. spicata), and Scotch spearmint (M. cardiaca) are grown commercially for their essential oils. Essential oils, which are produced in leaf glands primarily on the lower leaf surface, contain volatile terpenes and sesquiterpenes that produce the unique flavoring for candy, chewing gum, toothpaste, medicines and other products. The oil, initially recovered by on-farm steam distillation, is further processed and blended to produce oils to meet specifications requested by product manufacturers.
We compared the efficacy of Heterorhabditis marelatus Liu & Berry, a newly described species of entomopathogenic nematode, with that of H. bacteriophora Poinar against the root weevils, Otiorhynchus ovatus (L.) and O. sulcatus in strawberry. In the laboratory, H. marelatus was significantly more virulent than H. bacteriophora on O. ovatus and O sulcatus 7 d after nematode application at 14°C. In field experiments, H. marelatus applied at concentrations of 52 and 136 infective juveniles (IJs) per square centimeter, reduced root weevil larvae and pupae 75.3 and 77.4%, respectively, 20 d after nematode application. H. bacteriophora applied at concentrations of 128 and 379 IJs per square centimeter reduced root weevils 50.0 and 74.0%, respectively. Both nematode species were detected up to 30 dafter application by baiting with Galleria mellonella larvae in soil samples collected from the field.