**Insect Pests of Chickpea and Lentil**

**Pod Borers: *Helicoverpa armigera* and *Helicoverpa punctigera***

Nearly 60 insect species are known to feed on chickpea, of which the pod
borers *Helicoverpa armigera* and *Helicoverpa punctigera* (Lepidoptera:
Noctuidae) are the major pests. The former is a major pest of chickpea
in Asia, Africa, and Australia, while the latter is confined to
Australia. *Helicoverpa-*inflicted losses to chickpea crops in the
semi-arid tropics are estimated at over US \$328 million annually. Pod
borers rarely become a serious pest on lentil. Worldwide, losses due to
*Heliothis*/*Helicoverpa* in cotton, legumes, vegetables, cereals, etc.,
exceed \$2 billion, and the cost of insecticides used to control these
pests is over \$1 billion annually. There are several common names for
pod-borers, namely cotton bollworm, corn earworm, African cotton
bollworm, native budworm, old world bollworm, legume pod borers, gram
pod borer, and tomato fruit worm.

**Geographic distribution**

*Helicoverpa armigera* is widely distributed in Asia, Africa, Australia,
and the Mediterranean Europe, while *H. punctigera* is restricted to
southern regions of Australia. Additionally, there are reports of *H.
armigera* outbreaks in Hungary, Italy, Romania, Slovakia, Spain, Sweden,
Switzerland, and the United Kingdom.

**Host range**

*Helicoverpa armigera* and *H. punctigera* are major pests of cotton,
pigeonpea, chickpea, sunflower, tomato, maize, sorghum, pearl millet,
okra, *Phaseolus* spp., vegetables, tobacco, linseed, a number of fruits
(*Prunus, Citrus,* etc.), and forest trees. In recent years, *H.
armigera* damage has been reported in carnation, grapevine, apple,
strawberries, finger millet, etc. *Helicoverpa punctigera* is a major
pest of cotton, corn, sorghum, tomato, chickpea and other grain legumes.

**Nature of damage**

*Helicoverpa* females lay eggs singly on leaves, flowers, and young
pods. The larvae initially feed on the foliage (young leaves) in
chickpea and a few other legumes (Fig. 1), but mostly on flowers and
flower buds in cotton, pigeonpea, etc. The young seedlings of chickpea
may be destroyed completely, particularly under tropical climates in
southern India. Larger larvae bore into pods/bolls and consume the
developing seeds inside the pod (Fig. 2). In Australia where the climate
is cooler, the *Helicoverpa* populations build up in spring, attacking
chickpea in late spring before moving on to summer crops growing in the
sub-tropical regions.

**Life cycle**

The oviposition period lasts for 5 to 24 days, and a female may lay up
to 3,000 eggs, mainly at night on leaves, flowers, and pods (Fig. 3).
The egg incubation period depends on temperature, and varies between 2
to 5 days (usually 3 days). Duration of larval development depends not
only on the temperature, but also on the nature and quality of the host
plant, and varies between 15.2 days on maize to 23.8 days on tomato
(Fig. 4). The number of larval instars varies from 5 to 7, with six
being most common. The larvae pupate in the soil (Fig. 5). The pre-pupal
period lasts for 1 to 4 days. The larvae spin a loose web of silk before
pupation. In non-diapausing pupae, the pupal period ranges from about 6
days at 35°C to over 30 days at 15°C. The diapausing period for pupae
may last several months. Pale colored adults are produced from pupae
exposed to temperatures exceeding 30°C. In captivity, longevity varies
from 1 to 23 days for males and 5 to 28 days for females (Fig. 6).

*Helicoverpa armigera* exhibits a facultative diapause, which enables it
to survive adverse weather conditions in both winter and summer. The
winter diapause is induced by exposure of the larvae to short
photoperiods and low temperatures. In China and India, *H. armigera*
populations are comprised of tropical, sub-tropical, and temperate
ecotypes. In subtropical Australia, *H. armigera* undergoes diapause
during winters when the temperatures are low. High temperatures can also
induce diapause. It enters a true summer diapause when the larvae are
exposed to very high temperatures (43°C for 8 h daily), although the
proportion of females entering diapause is nearly half compared to that
of males. At these temperatures, non-diapausing males are sterile. In
Australia, *H. punctigera* has been observed to enter a diapause in
spring when temperatures are quite high and plant hosts are scarce.

**Management**

**Economic thresholds.** Monitoring of *Helicoverpa* populations is
necessary to determine if threshold has been exceeded and control
measures are required. Action thresholds based on egg numbers have been
used to make control decisions. One larva per meter row in chickpea
causes economic loss. A simple rule of thumb based on monsoon rains and
November rainfall has been developed to forecast *H. armigera*
populations in India. Models for long-range forecasts of *H. armigera*
and *H. punctigera* populations in Australia have also been developed.
These population-forecasting models may be incorporated into crop
production models for pest management. In Australia, three crops,
cotton, tomato and maize, have high levels of *Helicoverpa* attack and
require multiple sprays of pesticides. Of the legume crops, field peas
and chickpeas are spring flowering crops grown in the southern regions
of Australia, and usually suffer sporadic damage from *H. punctigera*,
requiring a single pesticide application only.

**Host plant resistance.** The development of crop cultivars resistant
or tolerant to *H. armigera* and *H. punctigera* has considerable
potential for use in integrated pest management, particularly under
subsistence farming conditions in developing countries. Several chickpea
germplasm accessions (ICC 506EB, ICC 10667, ICC 10619, ICC 4935, ICC
10243, ICCV 95992, and ICC 10817) with resistance to *H. armigera* have
been identified, and varieties such as ICCV 7, ICCV 10, and ICCL 86103
with moderate levels of resistance have been released for cultivation
(Fig. 1). However, most of these lines are highly susceptible to
Fusarium wilt. Therefore, concerted efforts have been made to break the
linkage by raising a large population of crosses between *Helicoverpa-*
and wilt-resistant parents. Several wild relatives of chickpea have
shown high levels of resistance to *H. armigera,* and efforts are
underway to transfer resistance from the wild relatives into high
yielding varieties of chickpea

**Genetically modified crops.** In recent years, genetic engineering has
enabled the introgression of genes from distantly related organisms
(i.e., *Bacillus thuringiensis*) into crops such as cotton, corn,
pigeonpea, and chickpea. Chickpea cultivars ICCV 1 and ICCV 6 have been
transformed with *cry IAc* gene. Insect feeding assays indicated that
the expression level of the *cry IAc* gene was inhibitory to the
development and feeding by *H. armigera*. Efforts are underway at
ICRISAT to develop transgenic chickpeas for resistance to pod borer. A
resistance management strategy has been developed for transgenic cotton
growing in Australia to prevent undesirable side effects, including the
development of resistance to *Bt*, which will also be applicable to
chickpea in case transgenic chickpeas are released for cultivation.

######## Cultural manipulation of the crop and its environment. A number of cultural practices such as time of sowing, spacing, fertilizer application, deep ploughing, interculture, and flooding have been reported to reduce the survival and damage by *Helicoverpa* species. Inter-cropping or strip-cropping with marigold, sunflower, linseed, mustard, or coriander can minimize the extent of damage to the main crop. Strip-cropping also increases the efficiency of chemical control. Hand-picking of large larvae can reduce *Helicoverpa* damage. However, the adoption of cultural practices depends on the crop husbandry practices in a particular agro-ecosystem. An area-wide management strategy has been implemented in regions of Queensland and New South Wales, Australia, to suppress local population densities of *H. armigera*, with chickpea trap crops playing an important role in this strategy. The chickpea trap crop is planted after the commercial crops to attract *H. armigera* emerging from winter diapause. The trap crops are destroyed before larvae commence pupation. As a result, the overall *H. armigera* pressure on summer crops is reduced, resulting in greater opportunity for adoption of soft control options, reduced insecticide use, and greater activity of the natural enemies. 

**Natural enemies.** The importance of biotic and abiotic factors on the
seasonal abundance of *H. armigera* and *H. punctigera* is poorly
understood. Some parasitic wasps avoid chickpea due to dense layers of
trichomes and their acidic exudates. *Trichogramma* egg parasitoids are
seldom present in high numbers in chickpea crops in India. The
ichneumonid wasp, *Campoletis chlorideae* is an important larval
parasitoid of *H. armigera* on chickpea in India. The dipteran
parasitoids *Carcelia illota*, *Goniophthalmus halli*, and *Palexorista
laxa* have been reported to parasitize up to 54% of the larvae on
chickpea. Predators such as *Chrysopa* spp*., Chrysoperla* spp.*, Nabis*
spp*., Geocoris* spp*., Orius* spp*.,* and *Polistes* spp. are common in
India. Provision of bird perches or planting of tall crops that serve as
resting sites for insectivorous birds such as Myna (*Acridotheris
tritis)* and Drongo (*Dicrurus macrocercus)* helps to reduce the numbers
of *H. armigera* larvae.

**Biopesticides and natural plant products.** The use of microbial
pathogens such as *H. armigera* nuclear polyhedrosis virus (HaNPV),
entomopathogenic fungi, *Bacillus thuringiensis* (*Bt*), nematodes, and
natural plant products such as neem, custard apple, and karanj
(*Pongamia pinnata)* kernel extracts have shown some potential to
control *H. armigera*. HaNPV has been reported to be a viable option to
control *H. armigera* in chickpea in India. Jaggery (locally made brown
sugar from sugarcane juice) (0.5%), sucrose (0.5%), egg white (3%), and
chickpea flour (1%) increase the activity of HaNPV. In Australia, the
efficacy of HaNPV in chickpea has been increased by the addition of milk
powder, and more recently the additive Aminofeed® (Farma-Chem,
Australia). The entomopathogenic fungus *Nomuraea rileyi* (10^6^ spores
per ml) resulted in 90 to 100% mortality of the larvae. Another
entomopathogenic fungus, *Beauveria bassiana* (2.68 x 10^7^ spores per
ml) resulted in 10% reduction in damage by *H. armigera* over the
control plants. *Bt* formulations are also used as sprays to control
*Helicoverpa*. Spraying *Bt* formulations in the evening results in
better control than spraying at other times of the day.

**Chemical control.** Management of *Helicoverpa* in India and Australia
in chickpea and other high-value crops relies heavily on insecticides.
There is substantial literature on the comparative efficacy of different
insecticides against *Helicoverpa*. Endosulfan, cypermethrin,
fenvalerate, methomyl, thiodicarb, profenophos, spinosad, and indoxacarb
have been found to be effective for controlling *H. armigera*. Spray
initiation at 50% flowering has been found to be most effective.
Development of resistance to insecticides is a major problem in *H.
armigera,* but not in *H. punctigera* because of its high mobility.
*Helicoverpa armigera* populations in several regions have developed
resistance to pyrethroids, carbamates, and organophosphates.
Introduction of new compounds such as thiodicarb, indoxacarb, and
spinosad has helped in overcoming development of resistance in *H.
armigera* to conventional insecticides.

**Integrated pest management (IPM).** Several management tactics have
been investigated, which provide a framework for improved management of
pod borers in chickpea and lentil cropping systems worldwide. For
example, crop cultivars with resistance to *Helicoverpa* (derived
through conventional plant breeding or biotechnological approaches) can
play an important role. Cultural practices such as deep ploughing,
interculture, flooding, and intercropping could potentially reduce the
intensity of *Helicoverpa*. Although the role of natural enemies as
biological control agents is unclear, their impact could be improved by
reducing pesticide applications, and adopting cropping practices that
encourage their activity. Most studies have shown that insecticide
applications are more effective than neem kernel extracts, *Bt*, HaNPV,
or augmentative releases of natural enemies. However, biopesticides and
synthetic insecticides, applied alone, together, or in rotation, are
effective for *Helicoverpa* control in chickpea. Moreover, scouting for
eggs and young larvae is critical for initiating timely control
measures. Insecticides with ovicidal action, and/or systemic action are
effective against *Helicoverpa* during the flowering stage. Finally, the
development of transgenic plants with different insecticidal genes,
molecular marker assisted selection, and exploitation of the wild
relatives of *Cicer* and *Lens* species should be pursued to develop
comprehensive programs for *Helicoverpa* management on chickpeas and
lentils.

###### **Selected References**

Commonwealth Agricultural Bureau International (CABI). 1993.
Distribution Maps of Plant Pests, No. 15. Commonwealth Agricultural
Bureau International, Wallingford, UK.

Fitt, G. P. 1989. The ecology of *Heliothis* species in relation to
agro-ecosystems. Annu. Rev. Entomol. 34:17-52.

Fitt, G. P., and Cotter, S. C. 2005. The *Helicoverpa* problem in
Australia: Biology and Management. In: *Heliothis/Helicoverpa*
Management: Emerging Trends and Strategies for Future Research (Sharma,
H.C., ed.). Oxford and IBH Publishing, New Delhi, India. pp. 45-61.

King, A. B. S. 1994. *Heliothis/Helicoverpa* (Lepidoptera: Noctuidae).
In: Insect Pests of Cotton (Matthews, G.A., and Tunstall, J.P., eds.).
CAB International, Wallingford, UK. pp. 39-106.

Maelzer, D.A., and Zalucki, M.P. 2000. Long range forecasts of the
numbers of *Helicoverpa punctigera* and *H. armigera* (Lepidoptera:
Noctuidae) in Australia using the Southern Oscillation Index and the sea
surface temperature. Bull. Entomol. Res. 90:133-146.

Matthews, M. 1999. Heliothine Moths of Australia. A Guide to Pest
Bollworms and Related Noctuid Groups. Monograph on Australian
Lepidoptera, Volume 7. CSIRO Publishing, P O Box 1139, 150 Oxford
Street, Callingford, Victoria, 3066, Australia, 320 pp.

Romeis, J., and Shanower, T.G. 1996. Arthropod natural enemies of
*Helicoverpa armigera* (Hübner) (Lepidoptera: Noctuidae) in India.
Biocontr. Sci. Tech. 6:481-508.

Sharma, H. C. (ed.). 2005. *Heliothis/Helicoverpa* Management: Emerging
Trends and Strategies for Future Research. Oxford and IBH Publishers,
New Delhi, India, 469 pp.

(Prepared by H. C. Sharma, T. J .Ridsdill-Smith and S. L. Clement)

![](media/image1.jpeg){width="2.4in" height="1.8in"}

Fig. 1. Leaf damage by *Helicoverpa armigera* in chickpea (Left --
Resistant line ICC 506EB, and Right - Susceptible line ICC 3137).
(Courtesy ICRISAT)

![](media/image2.jpeg){width="3.1256944444444446in"
height="1.0555555555555556in"}

Fig. 2. Pod damage by *Helicoverpa armigera.* (Courtesy ICRISAT)

![](media/image3.jpeg){width="3.0in" height="2.046527777777778in"}

Fig. 3. Eggs of *Helicoverpa armigera* on chickpea. (Courtesy ICRISAT)

![](media/image4.jpeg){width="2.0in" height="0.7201388888888889in"}

Fig. 4. A) Larva of *Helicoverpa armigera* (Photo: ICRISAT), and B) *H.
punctigera.* (Courtesy Richard Lloyd)

![](media/image5.jpeg){width="2.3756944444444446in"
height="1.461111111111111in"}

Fig. 5. Pupa of *Helicoverpa armigera.* (Courtesy ICRISAT)

![](media/image6.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 6. Adult of *Helicoverpa armigera.* (Courtesy ICRISAT)

## 

## Pea moth: *Cydia nigricana*

Pea moth *Cydia* (*Laspeyresia*) *nigricana* (Lepidoptera: Tortricidae)
is mainly found on wild and cultivated peas, but it also feeds on
lentils. It is commonly known as Erbsenwickler (DE), tordeuse du pois
(FR), polilla del guisante (ES), tortrice dei piselli (IT), traça da
ervilha (PT), ärtvecklare (SE) or pea moth. The lack of efficient
control methods, together with a low damage threshold in green peas,
makes pea moth an economically important insect in Europe.

**Geographic distribution**

Pea moth, *C. nigricina* has a Paleoarctic distribution, ranging from
Europe to China and Japan. It is widespread in colder climates, and is
found up to 64° latitude in Scandinavia. It has been introduced to North
Africa, and also to Canada and USA, where it is most abundant in the
northeastern part of the country.

**Host plants**

Various wild and cultivated Leguminosae, especially species of *Vicia*
and *Lathyrus*, garden pea, clover, and lentil. It also feeds on
chickpea and lupine.

**Nature of damage**

The larvae attack the seeds inside the pods (Fig. 1). Damage is detected
only when the pods are opened. Larval feeding spoils the pods with
excrement and silk (Fig. 2). One larva damages up to 6 seeds, but
usually only 1 to 2 are severely damaged. Seed damage, presence of
larvae, and stains lead to rejection of consignments by the processors.

**Life cycle**

The adult moth is small (15 mm wingspan) and delicate. It is
distinguished by grey-brown forewings with traces of ocherous, and
conspicuous black and white costal strigulae and interspaces. The
ocellus, which is otherwise typical for *Cydia*, is poorly developed
(Fig. 3).

Adults appear during the flowering stage and lay eggs on plants (Fig.
4). A female may lay up to 200 eggs. Embryonic development is completed
in 6 to 10 days. Egg-laying occurs 5 to 11 days after eclosion. The
female deposits 1 to 3 eggs on the stipules or the leaflets (Fig. 5).
The larva is yellowish white with dark spots and short hairs, measures
13 to 18 mm, and has a light brown head (Fig. 2). Larval development is
completed in 18 to 30 days, and there are five instars. The larva leaves
the pod and migrates to the ground where it spins a cocoon containing
particles of soil. Larvae hibernate in the soil and pupate in spring.
There is one generation per year.

Moth eclosion is influenced by soil temperatures and photoperiod.
Temperatures between 21 to 25° C, and 70 to 90% R.H. are favorable. The
sum of effective temperatures for insect development is 442°C. Hot and
dry weather is unfavorable to the insect.

**Mating behavior and chemical ecology**

Male moths become active during the late afternoon in broad sunlight and
are seen to actively fly over host plants. They readily orient to
pheromone-releasing females or synthetic pheromone lures over a
distance. After landing, the male starts wingfanning and rhythmically
extrudes its genital claspers at the tip of the erect abdomen. The
female then walks towards the male, touching him with her antennae, and
the male then attempts copulation. The female response to the male
courtship is largely mediated by volatile chemicals released from
androconial scent organs on the male hindwing. Male-produced pheromones
are, however, active only at close distance, while female-produced sex
pheromone attracts the males over 50 m or more.

Pea moth females are attracted to peas for oviposition. The eclosion of
adults is tightly correlated with flowering of pea. The females are
attracted to pea fields from a long distance. However, the chemicals
encoding host plant attraction have not been elucidated. One open
question is whether the pea moth females mate on the host plant or at
the site of emergence, before reaching a suitable host plant. Better
knowledge of the role of host plant cues in pea moth reproductive
behavior is the key to integrated management strategies.

**Management**

**Cultural control.** Low cropping intensity and area wide separation of
grain peas from vegetable peas are key factors in reducing pea moth
infestations. Pea fields of the previous year are the main source of
infestation. The distance to newer fields should exceed the flight range
of pea moths, which has been estimated to be 2 km or more. High pea moth
populations have also been found in grasslands and natural habitats
where wild Leguminosae grow.

Early-sowing in combination with short-duration genotypes reduces the
availability of peas at the susceptible phenological stages, i.e.
flowers and young pods, during the main season. Intercropping with
barley, deep plowing of fields with overwintering larvae, and
eradication of weeds are other methods to reduce pea moth damage.

**Natural enemies.** The egg parasitoid, *Trichogramma evanescens,* can
be used for biological control of this pest, although under practical
conditions, efficacy needs to be improved. Naturally- occurring
hymenopterous parasitoids, though abundant, are apparently not capable
of reducing pea moth populations to below economic thresholds.

**Microbial control**. The granulosis virus of the codling moth, *Cydia
pomonella,* is reported to be effective under laboratory and small scale
field experiments. However, efficacy was unsatisfactory in several
large-scale field tests, even when high dosages were applied.

**Chemical control.** Pyrethroids or carbamates are commonly used, but
the control efficacy of insecticide sprays is often limited.
Pheromone-baited traps are efficient tools to time such insecticide
sprays.

Chemical control of *C. nigricana* is difficult, since the pea moth is
protected from insecticide sprays in soil or in pods during most of its
life cycle. Only the neonate larvae are susceptible to insecticides,
before they penetrate the pods. Timing of insecticide sprays is achieved
by pheromone-baited monitoring traps. In spite of optimized timing of
pesticide application, it is often not possible to reduce pea moth
infestations below the low damage threshold of 0.5 to 1% in green peas,
and likely in lentils.

**Pheromones**. The pea moth pheromone is codlemone acetate,
(*E*,*E*)-8,10-dodecadienyl acetate. The geometric isomers of this
compound, which are quickly formed on pheromone lures, are strong
attraction antagonists. Isomerization of peromone lures within few days
makes it impossible to use synthetic pheromone for monitoring in fields
traps. A less attractive pheromone mimic is used instead.

Isomerization of the main pheromone compound is, on the other hand, not
an obstacle for pheromone-mediated mating disruption. A repellent blend
of pheromone and antagonistic isomers was efficient for population
control by mating disruption in isolated pea fields. Main obstacles to a
more widespread use of mating disruption in pea moth control are the
availability of a suitable dispenser material and the cost of dispenser
application in pea fields. In comparison, mating disruption has been
successfully used against codling moth, a closely related species.

**Integrated pest management**. Early-sowing, short-duration genotypes,
and intercropping can be combined with insecticide treatments.
Pheromone-baited monitoring traps are an inexpensive and efficient tool
to time sprays. Further development of pheromone-mediated mating
disruption, resistant cultivars, and identification of plant volatile
cues that attract gravid females for oviposition would be a significant
step towards sustainable and more efficient control of pea moth.

**Selected References**

Bengtsson, M., Karg, G., Kirsch, P. A, Löfqvist, J., Sauer, A., and
Witzgall, P. 1994. Mating disruption of pea moth *Cydia nigricana* F.
(Lepidoptera: Tortricidae) by a repellent blend of sex pheromone and
attraction inhibitors. J. Chem. Ecol. 20:871-887.

Beniwal, S. P. S., Bayaa, B., Weigand, S., Makkouk, K. H., and Saxena,
M. C. 1993. Field guide to lentil diseases and insect pests.
http://www.icarda.org/Publications/Field_Guides/Lentil/Lentil.htm.

Darty, J. M., and Wimmer, F. 1983. Lentil: Control of the pea midge and
the peamoth (*Contarinia lentis*). Phytoma 347:29-31.

Gould, H. J., and Legowski, T. J. 1964. Spray warnings for pea moth
(*Laspeyresia nigricana*) based on its biology in the field. Entomol.
Exp. Appl. 7:131-138.

Payne, C. C. 1981. The susceptibility of the pea moth, *Cydia
nigricana*, to infection by the granulosevirus of the codling moth,
*Cydia pomonella*. J. Invert. Pathol. 38:71-77.

Witzgall, P., Bengtsson, M., Unelius, C. R., and Löfqvist, J. 1993.
Attraction of pea moth *Cydia nigricana* F. (Lepidoptera: Tortricidae)
to female sex pheromone (*E*,*E*)-8,10-dodecadien‑1‑yl acetate, is
inhibited by geometric isomers (*E*,*Z*), (*Z*,*E*) and (*Z*,*Z*). J.
Chem. Ecol. 19:1917-1928.

Witzgall, P., Bengtsson, M., Karg, G., Bäckman, A. C., Streinz, L.,
Kirsch, P. A., Blum, Z., and Löfqvist, J. 1996. Behavioral observations
and measurements of aerial pheromone concentrations in a mating
disruption trial against pea moth *Cydia nigricana* F. (Lepidoptera,
Tortricidae). J. Chem. Ecol. 22:191-206.

Wright, D. W., and Geering, Q. A. 1948. The biology and control of the
pea moth *Laspeyresia nigricana* Stephh. Bull. Entomol. Res. 39:57-48.

(Prepared by G. Thöming, H. Saucke and P. Witzgall)

![](media/image7.jpeg){width="3.0in" height="1.2402777777777778in"}

Fig. 1. *Cydia nigricana* larvae damage the lentil seeds inside a pod.
(Courtesy P. Witzgall)

![](media/image8.png){width="1.9305555555555556in" height="2.0in"}

Fig. 2. *Cydia nigricana* larval feeding spoils pods with excrement and
silk. (Courtesy P. Witzgall)

![](media/image9.jpeg){width="3.0in" height="1.8902777777777777in"}

Fig. 3. Adult of *Cydia nigricana.* (Courtesy P. Witzgall)

![](media/image10.jpeg){width="3.0in" height="2.640277777777778in"}

Fig. 4. Adults of *Cydia nigricana* appear during the flowering stage
and lay eggs on plants. (Courtesy P. Witzgall).

![](media/image11.png){width="1.3611111111111112in" height="2.0in"}

Fig. 5. Eggs of *Cydia nigricana.* (Courtesy P. Witzgall)

Lima Bean Pod Borer, *Etiella zinckenella*

Lima bean pod borer, *Etiella zinckenella* (Lepidoptera: Pyralidae) is
an important insect pest of several pulse crops, including lentil. It is
commonly known as lima bean pod borer, spiny pod borer, or pea pod
borer. It is an occasional pest of lentil, and is not a pest of
chickpea.

**Geographical distribution**

The lima bean pod borer, *E. zinckenella* is widely distributed in Asia,
Africa, and Europe. It is also a serious pest in Australia, New
Caledonia, Papua New Guinea, and Solomon Islands. In the American
continent, it is present in Canada, USA, West Indies, and Central
America.

**Host range**

It feeds on several leguminous species, especially lima bean, cowpea,
pigeonpea, lentil, horse gram, green gram, field pea, and sunhemp.

**Nature of damage**

The presence of a hole on the pod surface, dry light-colored frass, and
webbing in the pod are indications of infestation by the lima bean pod
borer, *E. zinckenella*. As a result of insect damage, the pods are
poorly developed. Individual seeds have holes and internal portions are
gutted out (Fig. 1). The pods are partially or completely consumed
inside. Externally, the pods give shrunken appearance, with small
surface punctures. Larvae generally feed on maturing pods. Lima bean pod
borer population builds up by the end of the season, when the
temperature is high. The caterpillar is greenish or pinkish red, with a
yellow head. It moves violently when disturbed. The adults are small,
brown, and active at night (Fig. 2). Its infestation can be detected by
the presence of small punctures on the surface of the pods, and the
larvae can be observed by splitting the pods.

**Life cycle**

The adult moths are 10 to 12 mm long with a wingspan of 22 to 28 mm. The
forewings are brown-gray with a white anterior margin (Fig. 3). Mating
takes place at night or in dark places, and the females lay eggs on
young pods in clusters of 2 to 12. A female lays 47 to 178 eggs in 5 to
6 days. Eggs are laid near the calyx of the flowers or on pods. The
larva immediately bores into the pods and feeds internally. Larvae
attain a maximum length of 15 mm, and are greenish with a brown line
(Fig. 4). The larvae often move from one pod to another. The average
egg, larval, pre-pupal, and pupal periods on lentil have been reported
to be 5.4, 17.2, 2.3, and 13.8 days, respectively. The larvae enter
diapause in winter. Pupation normally takes place in the soil, but
sometimes on pods (Fig. 5). One generation is completed in about 4 weeks
under favorable conditions, and there are 3 to 5 generations per year.
The adults survive for one week, and the females live longer than the
males.

**Management**

**Host plant resistance.** Host plant resistance can play a major role
in reducing the losses due to *E. zinckenella* in lentil. Short-duration
genotypes are more susceptible than the medium- and long-duration
genotypes. The line LH 90-39 is resistant, while LL 147 is tolerant to
*E. zinckenella* damage. Lines P 927 and P 202 have been reported to be
resistant, and yield 52.9 and 43.5% more than L 9-12, respectively.

**Natural enemies.** Several natural enemies have been reported to
control *E. zinckenella.* These include *Bracon etiellae*, *B.
pectoralis*, *Phanerotoma planifrons*, *Pigeria piger*, *P.
hendecasisella*, *Exorista roborator*, and *Tetrastichus* spp. (Fig. 6).
However, there are no reports on use of natural enemies for classical
biological control of the lima bean pod borer.

**Chemical control**. This is an occasional pest of lentil, but
insecticide application may be necessary under heavy infestation. Sprays
of methidathion (0.5 kg ai ha^-1^), deltamethrin (38 g ai ha^-1^), or
endosulfan (6 ml L^-1^) at flowering and early pod setting have been
reported to provide effective control of this pest. Abamectin 1.8% EC (1
ml 20 L^-1^), α-cypermethrin 10% (1.5 ml 2 L^-1^), and cyhalothrin 2.5%
(2 ml 2 L^-1^) have also been found to provide effective control.

**Integrated pest management.** The Lima bean pod borer infestations
quite often are low, and do not warrant control. There is a positive
relationship between moths caught in sweep nets during flowering and
pod-formation and seed damage, and therefore, rough predictions of
damage can be made based on moth catches to undertake control measures.
Under heavy infestations, application of insecticides with a strong
contact and systemic action may be effective. Varieties that are less
susceptible to the pod borer may be recommended for cultivation in areas
endemic to this pest.

**Selected References**

Brar, J. S., Verma, M. M., Sandhu, T. S., Singh, B., and Gill, A. S.
1989. LL 147 variety of lentil (*Lens culinaris* L.). J. Res. Punjab
Agric. Univ. 26:170.

# CABI (Commonwealth Agricultural Bureau International). 2002. Crop Protection Compendium*.* Commonwealth Agricultural Bureau International, Wallingford, UK.

Jaglan, M. S., Sucheta, Khokhar, K. S., and Kumar, S. 1995. Biology of
lentil pod borer, *Etiella zinckenella* Treitschke on lentil and pea.
Ann. Biol. 11:224-228.

# Nair, M. R. G. K. 1975. Insects and Mites of Crops in India. Indian Council of Agricultural Research, New Delhi, India. 405 pp.

# Sandhu, G. S., and Verma, G. C. 1968. *Etiella zinckenella* Treitschke (Lepidoptera: Phyticidae) as a pod borer of lentil in Punjab. J. Bombay Nat. Hist. Soc. 65:799.

(Prepared by C.P. Srivastava and H.C. Sharma)

# ![](media/image12.jpeg){width="2.3756944444444446in" height="1.5763888888888888in"}

# Fig. 1. Lentil seed damage by pod borer, *Etiella zinckenella.* (Courtesy ICRISAT)

# 

# ![](media/image13.jpeg){width="3.0in" height="1.9930555555555556in"}

# Fig. 2. Adult of pod borer, *Etiella zinckenella.* (Courtesy ICRISAT)

# ![](media/image14.jpeg){width="3.0in" height="1.6in"}

# Fig. 3. Wing span and coloration of pod borer, *Etiella zinckenella* adult. (Courtesy ICRISAT)

# ![](media/image15.jpeg){width="3.0in" height="1.9930555555555556in"}

# Fig. 4. Larva of pod borer, *Etiella zinckenella.* (Courtesy ICRISAT)

# 

# ![](media/image16.jpeg){width="3.0in" height="1.9930555555555556in"}

# Fig. 5. Pod borer, *Etiella zinckenella* pupation in the pod. (Courtesy ICRISAT)

# 

# ![](media/image17.jpeg){width="3.0in" height="1.9930555555555556in"}

# Fig. 6. Pod borer, *Etiella zinckenella* larval parasitoid, *Tetrastichus* spp. (Courtesy ICRISAT) 

### Leaf Weevils: *Sitona crinitus*

Leaf weevil, *Sitona crinitus* (*Sitona macularius*) (Coleoptera:
Curculionidae) is one of the main insect species attacking lentil. The
adults feed on the foliage, but larvae cause the main damage. The larvae
are a serious pest on N~2~ fixing nodules of lentils in West Asia. They
are commonly known as lead leaf weevil or *Sitona* weevil.

**Geographical distribution**

*Sitona crinitus* is one of the main insect species attacking lentil in
West Asia (Turkey, Syria, Lebanon, and Jordan), southern Europe, North
Africa, and the former USSR.

**Host range**

*Sitona crinitus* shows a distinct feeding preference among the grain
legumes. *Vicia sativa* is more preferred than *Medicago polymorpha*,
*Lathyrus sativus*, *L. ochrus*, and *L. cicera*. *Lens culinaris* is
the next most severely damaged species, followed by *Trifolium
angustifolium*. *Lathyrus* *ochrus* is the least damaged species. Both
chickpea and faba bean are non-host plants of *S. crinitus*.

**Nature of damage**

Both adults and larvae of *S. crinitus* damage (Fig. 1) the lentil crop,
but larvae are the main damaging stage. The adult weevils feed on
foliage in a characteristic manner, making semicircular notches from the
leaf edges early in the season. The adult feeding usually does not
affect yields, unless populations are very high and/or unfavorable
environmental conditions limit the growth of the lentil seedlings, and
the plants cannot compensate the damage to foliage quickly. The larvae
are a serious pest on N~2~ fixing nodules of lentils in West Asia and
North Africa. Nodule damage is higher in early-sown than in late-sown
lentils. There is a positive correlation between visual damage score and
nodule damage by *S. crinitus*. Visual damage can be used for evaluating
a large number of genotypes for *Sitona* resistance under field
conditions. The white *Sitona* larvae can be seen inside the nodules in
uprooted lentil plants. Damage by *Sitona* reduces the nitrogen-fixing
ability of the crop. Mineral nitrogen does not compensate for the
damaged nodules, and fails to supplement fixed nitrogen for yield
increase. The foliage of damaged plants assumes a yellow appearance
similar to nitrogen deficiency characteristics. At times, the
infestation of leaflets may be \>90% and the larvae destroy most of the
nodules. At high infestation levels (\>90% nodule damage), the insect
caused 17.7 and 14.1% loss in straw and grain yields, respectively.
*Sitona crinitus* is also an efficient vector of broad bean stain
cosmovirus (BBSV) in lentil.

**Life cycle**

The adult weevils have a grey-brown and elongated body of 3 - 4 mm
length. The pronotum has three straight longitudinal light lines, and
the elytra have three rows of dark and white spots. The females lay
spherical yellow eggs (Fig. 2), which later turn black. The larvae are
cream-white, with a brown head capsule, legless, while the pupae are
white.

Overwintered adults of *S. crinitus* appear in the second half of March
and feed on young shoots and leaves, while the larvae (Fig. 3) appear
when the climatic conditions are suitable, and have root nodules to feed
upon. The spring migration of adults and the number of months spent in
hibernation have a significant effect on adult lifespan. In the
Mediterranean region, where hot and dry summers prevail, the adults
(Fig. 5) aestivate in the soil, and start emerging in December/January.
Sometimes, the adults emerge in May, when the lentil matures. The adults
of the previous generation die in April/May. There is only one
generation per year, and the adults live for almost one year. After
copulation, the females start laying eggs on the soil around the lentil
plants or loosely on the leaves, which later fall to the ground. The
oviposition period lasts for several months, and each female lays
several hundred eggs. Temperatures \<10°C induce quiescence in eggs of
*S. crinitus*. The neonate larvae move into the soil and infest the
nodules. Each larva may consume many nodules during development until
pupation. It pupates in the soil. Depending upon temperatures, the
larval and pupal period may last for 5 - 6 and 3 - 4 weeks,
respectively.

**Management**

**Cultural control.** Crop rotation can reduce the infestation to some
extent, as the weevils emerge from the soil after aestivation. They are,
however, strong fliers and can migrate long distances. Weedy fields are
more prone to *Sitona* damage. Delaying planting by about two weeks
reduces *Sitona* infestation. However, early sowing combined with the
control of *Sitona* and weeds, and P application gives higher net
returns with virtually no risk of economic loss to the farmers.

**Host plant resistance.** Previous research on host plant resistance
against *S. crinitus* did not produce conclusive results, but recent
studies indicated that eight accessions of wild lentils (ILWL 110, ILWL
136, ILWL 166, ILWL 203, ILWL 207, ILWL 245, ILWL 254, and ILWL 258)
have high level of resistance to *Sitona* (≤10% nodule damage). The
resistant accession ILWL 245 belongs to *L. culinaris* subsp.
*orientalis*, the progenitor of the cultivated lentil, and can be
crossed with the cultivated lentils. This line is being used to
understand the inheritance of *Sitona* weevil resistance, and to
introgress resistance genes to cultivated lentils (Fig. 5). Isoflavone
reductase levels can be used to select lentils for resistance to
*Sitona*. The *cryIII* toxin expression in nodules has been found to
result in significant reduction in nodule damage by *S. lineatus* on
*Pisum sativum,* and *S. hispidulus* on *Medicago sativa*, and could be
used as strategy to produce *Sitona*-resistant lentils.

**Chemical control.** Carbofuran and aldicarb reduce nodule damage
significantly (Solh et al., 1986), and yield increases due to carbofuran
application are generally greater in early than in late-sown crop.
Lentil seeds can be treated with furathiocarb (12 ml kg^-1^ seed) or
apply granular insecticides such as carbofuran 5% G 10 kg^-1^ ha) at
planting. Use of carbofuran improves both nodulation and grain
production. Rhizobial inoculation and phosphorus application increase
lentil productivity in arid locations under Mediterranean environments.
If no preventive control is taken, and a high infestation is observed in
the field, spray with phosmet (1 kg a.i.^-1^ ha). This is less effective
than granular application and seed treatment. Alternatively,
chlorpyriphos, malathion, or oxy-demeton methyl can also be applied in
case of severe infestation. The water extract of *Melia azedarach*
kernels (50 g^-1^liter) significantly reduced *S. crinitus* adult damage
on lentil leaves for one week.

**Integrated Pest Management.** Crop rotation, delaying planting date by
about two weeks than the normal sowing, planting resistant varieties,
and development of *Bt*-transgenic lentils having *cryIII* toxin
expression in nodules could be used to manage *Sitona* damage in
lentils. Seed treatment with furathiocarb (12 ml kg^-1^ seed) or
application of granular insecticide (carbofuran 5% G @ 10 kg^-1^ ha)
reduces nodule damage significantly. The water extract of neem kernels
(@ 50 g^-1^liter) significantly reduces *S. crinitus* damage.
Chlorpyriphos, malathion, or oxy-demeton methyl can be sprayed in case
of severe infestation.

# Selected References

Bezdicek, D. F., Quinn, M. A., Forse, L., Heron, D., and Kahn, M. L.
1994. Insecticidal activity and competitiveness of *Rhizobium* spp.
containing the *Bacillus thuringiensis* subsp. *tenebrionis* --endotoxin
gene (cryIII) in legume nodules. Soil Biol. Biochem. 26:1637-1646.

El Damir, El Bouhssini, M., and Al Salty, M. N. 2000. Effect of *Melia
azedarach* (L.) (Meliaceae) fruit extracts on *Sitona crinitus* (H.)
(Coleoptera: Curculionidae) adults feeding. Arab J. Plant Prot.
18:64-67.

El Damir, M., Al-Salti, M. N., and El Bouhssini, M. 2001. Ecology and
biology of *Sitona crinitus H.* in northern Syria. Bassel Al-Assad J.
Engin. Sci. 14:109-119.

Islam, R., and Afandi, F. 1982. Effect of some insecticides on
nodulation and yield of two lentil cultivars. Lentil Exp. News Serv.
9:24-25.

Perez Andueza, G., Mozos Pascual de los, M., and Portillo Rubio, M.
2004. Main pests of lentil (*Lens culinaris* Medikus) in Castilla La
Mancha (central Spain): crop losses and influence on yield parameters.
Boletin de Sanidad Vegetal, Plagas 30:763-772.

Tahhan, O., and Hariri, G. 1983. The feeding and chemical control of
*Sitona maculatus* (Marsham) (Coleoptera: Curculionidae) on lentils in
northern Syria. Lens Newslett. 10:32-34.

Weigand, S., Lateef, S. S., El Din, N. E. S., Mahmoud, S. F., Ahmed, K.,
and Ali, K. 1994. Integrated control of insect pests of cool season food
legumes. Pages 679-694 in: Expanding the Production and use of Cool
Season Food Legumes, Proceedings of the Second International Food Legume
Research Conference on Pea, Lentil, Faba Bean, Chickpea, and Grasspea,
12-16 April 1992, Cairo, Egypt. Kluwer Academic Publishers Group,
Dordrecht, Netherlands.

(Prepared by M. El Bouhssini)

![](media/image18.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 1. Chickpea leaf damage by *Sitona crinitis.* (Courtesy ICARDA)

![](media/image19.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 2. Egg of *Sitona crinitis.* (Courtesy ICARDA)

![](media/image20.jpeg){width="2.0in" height="1.3666666666666667in"}

Fig. 3. Larva of *Sitona crinitis.* (Courtesy ICARDA)

![](media/image21.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 4. Adult of *Sitona critinis.* (Courtesy ICARDA)

![](media/image22.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 5. Reaction of chickpea genotypes to damage by *Sitona crinitis.*
(Courtesy ICARDA)

**Armyworms: *Spodoptera* spp.**

The beet armyworm, *Spodoptera exigua* (Noctuidae: Lepidoptera), is a
pest of chickpea in Mexico and north India, and became a serious pest of
chickpea (vegetative stage) in some regions of southern India in the
latter part of the 20^th^ century. Although chickpea is susceptible to
attack by *S. exigua*, it is not a pest of chickpea in the U.S. Pacific
Northwest. The western yellow-striped armyworm, *S. praefica* (Grote)
(Noctuidae: Lepidoptera), is a sporadic pest of chickpea and other grain
legumes in the U.S. Pacific Northwest.

**Geographic distribution**

Native to the Orient, the beet armyworm has become established in much
of the world, including Europe, Asia, Russia, Africa, Australia, India,
Pacific Islands, North America, and the West Indies. It is common in the
United States, but rarely overwinters where frost kills its host plants.
*Spodoptera praefica* is known only from the western United States,
principally California, Oregon and Washington.

**Host range**

The beet armyworm has a wide host range, occurring as a serious pest of
vegetable crops (asparagus, bean, beet, broccoli, cabbage, cauliflower,
celery, chickpea, corn, cowpea, eggplant, lettuce, onion, pea, pepper,
potato, radish, spinach, sweet potato, tomato, and turnip) in many parts
of the world. On field crops, it is of economic concern on alfalfa,
corn, cotton, peanut, safflower, sorghum, soybean, sugarbeet, and
tobacco. The western yellow-striped armyworm is a pest of several crops,
including lentil and chickpea in the U.S. Pacific Northwest.

**Nature of damage**

The young larvae of *S. exigua* initially feed gregariously on chickpea
foliage. As the larvae mature, they become solitary and eat large
irregular holes in foliage (Fig. 2). As a leaf feeder, beet armyworm
consumes much more chickpea tissue than the chickpea pod borer
*Helicoverpa armigera*, but has not been reported as a serious pest on
pods. Infestations of *S. praefica* usually appear late in the chickpea
growing season in the U.S. Pacific Northwest, with potential to inflict
economic damage by consuming foliage and pods.

**Life cycle**

The *S. exigua* females lay eggs in clusters of 50 to 150 eggs (Fig. 1).
A female lays 300 to 600 eggs on the lower surface of the leaves, near
the blossoms, and the terminal branches. The eggs are greenish white in
color, and covered with a layer of whitish scales, which gives the egg
mass a fuzzy or cottony appearance. Eggs hatch in 2 to 3 days during
warm weather. There are normally five instars, although additional
instars have also been reported. Duration of the instars I to V has been
observed to be 2.5, 1.5, 1.2, 1.5, and 3.0 days, respectively. The
larvae are pale green or yellow in color during the first and second
instars, but acquire pale stripes during the third-instar (Fig. 2).
During the fourth-instar, the larvae become darker dorsally, and possess
a dark lateral stripe (Fig. 3). Fifth-instar larvae are quite variable
in appearance, and tend to be green dorsally with pink or yellow color
ventrally, and a white stripe laterally. Sometimes, the larvae assume a
dark color. The body is practically devoid of hairs and spines. Pupation
occurs in the soil (Fig. 4). The pupal chamber is constructed from sand
and soil particles held together with an oral secretion, which hardens
upon drying. The pupa is light brown in color and measures 15 to 20 mm
in length. Duration of the pupal stage is 6 to 7 days during warm
weather. The moths are of moderate size, with wingspan of 25 to 30 mm.
The forewings are mottled gray and brown, and have irregular banding
pattern, and a light bean-shaped spot (Fig. 5). The hindwings are of
uniform gray or white color, and trimmed with a dark line at the
margins. Mating occurs soon after emergence, and oviposition begins
within 2 to 3 days. Oviposition extends over a 3 to 7 day period, and
the adults live for 9 to 10 days. Seasonal activity varies considerably
according to climate. The life cycle can be completed within 24 days.
There are several generations in a year, which vary across regions,
depending upon the climate.

The life cycle of *S. praefica* on chickpea in the U.S. Pacific
Northwest has not been studied, although its movement from weed hosts to
cultivated lentil and its economic impact on this crop in this region
has been described.

**Management**

**Economic thresholds and population monitoring.** Pheromone traps can
be used to detect the presence of beet armyworm adults. Visual sampling
for damage and larvae, combined with an action threshold of 0.3 larvae
per plant has been used in cabbage. However, there is no information on
action thresholds in chickpea. Regular monitoring of crops is
recommended because adult moths frequently invade the crops from
adjacent crops and weeds. Pheromones can also be used to disrupt mating
and inhibit or eliminate reproduction. Saturation of the atmosphere
around beet armyworm-susceptible crops might reduce mating by 97%.
Economic thresholds for *S. praefica* on chickpea have not been
established. However, a population of one larva per 1,000 cm^2^ on
lentils with green pods will cause 10% loss. Sweep-netting of larvae
will establish the presence of larvae in agricultural crops and a sex
attractant for trapping male *S. praefica* could potentially be used to
monitor male flights in chickpea.

**Host-plant resistance.** There is no published information on
resistance of chickpea varieties to *S. exigua* and *S. praefica*.
However, several progenies from interspecific hybridization involving
cultivated chickpea (*Cicer arietinum*) and a wild relative (*C.
reticulatum*) have exhibited high levels of resistance to *S. exigua*
(Fig. 6).

**Natural enemies.** Common natural enemies of *S. exigua* in U.S.
agricultural crops are braconids (*Chelonus insularis*, *Cotesia
marginiventris*, and *Meteorus autographae*) and a tachinid (*Lespsia
archippivora*). Also, predators like minute pirate bugs (*Orius* spp.),
big-eye bugs (*Geocoris* spp.), damsel bugs (*Nabis* spp.), and the
predatory shield bug (*Podisus maculiventris*) attack eggs and small
larvae. Red imported fire ant (*Solenopsis invicta)* attack pupae. In
India, *S*. *exigua* is a host of *Campoletis chlorideae* (Hymenoptera:
Ichneumonidae), an important parasitoid of the chickpea pod borer
*Helicoverpa armigera*. Natural enemies of *S. praefica* include many
parasitic insects and predators, but their impact on this insect in U.S.
chickpea crops is unknown.

**Biopesticides and natural plant products.** Nuclear polyhedrosis virus
isolated from beet armyworm is fairly effective as a bioinsecticide
under greenhouse conditions, where inactivation by ultraviolet light of
sunlight is not a severe problem. The nuclear polyhedrosis virus is
highly specific to beet armyworm, and is considered to be an important
mortality factor under natural conditions. The fungi *Beauveria
bassiana* and *Nomuraea rileyi* are pathogenic to *S. exigua* and larvae
are susceptible to neem products. A nuclear polyhedrosis virsus has been
found in *S. praefica*.

**Chemical control.** Information on insecticidal control of the beet
armyworm comes from publications on U.S. crops other than chickpea. For
example, eggs and young larvae can be controlled with foliar
applications of 5% cottonseed oil, but this concentration is damaging to
some plant species. Moreover, development of resistance to insecticides
is a major problem in the management of *S. exigua* because it attacks
several crops, which results in frequent insecticide applications and
status as a secondary or induced pest in some crops. Insecticides have
not been evaluated for control of *S. praefica* on chickpea, although
they are used against damaging populations in the U.S. Pacific
Northwest.

**Integrated pest management.** Improving the prospects for protecting
chickpea crops from *Spodoptera* infestations requires the development
of integrated pest management control and management tactics that would
include, in large part, adult monitoring with pheromone traps, field
sampling to estimate damaging populations, and the use of biopesticides
and natural plant products that are compatible with natural enemies. In
India**,** insecticides (endosulfan, fenvalerate, cypermethrin) can be
used to control outbreaks of *S. exigua.* In the absence of integrated
pest management programs for most of the grain legume insect pests in
North America, including *S. praefica*, the present control strategy
involves insecticides. The development and deployment of *Bt*-transgenic
chickpea has received attention, and could be a promising tool for
protecting chickpea crops from damage by *Spodoptera* infestations.

**Selected References**

Capinera, J. L. 2001. Handbook of Vegetable Pests. Academic Press, San
Diego, USA.

Commonwealth Agricultural Bureau (CABI). 1972. Distribution Maps of
Plant Pests. Map 302. CAB International, Wallingford, Oxfordshire, UK.

Clement, S. L., Wightman, J. A., Hardie, D. C., Bailey, P., Baker, G.,
and McDonald, G. 2000. Opportunities for integrated management of insect
pests of grain legumes. Pages 467-480 in: Linking Research and Marketing
Opportunities for Pulses in the 21^st^ Century, Knight, R., ed. Kluwer
Academic Publishers, Dordrecht, The Netherlands.

Eveleens, K. G., van den Bosch, R., and Ehler, L. E. 1973. Secondary
outbreak induction of beet armyworm by experimental insecticide
applications in cotton in California. Environ. Entomol. 2:497-503.

Gutierrez, P. E., Cortez, M. E., and Ayala, O. J. L. 1986. Population
dynamics of arthropod pests and beneficial animals in chickpea planted
on six dates and the effects on yield and grain quality. Revista
Chapingo (Mexico) 11-12:63-68.

Ruberson, J. R., Herzog, G. A., Lambert, W. R., and Lewis, W. J. 1994.
Management of the beet armyworm (Lepidoptera: Noctuidae) in cotton: role
of natural enemies. Florida Entomol. 77:440-453.

Sharma, H. C., Gowda, C. L. L., Stevenson, P. C., Ridsdill-Smith, T. J.,
Clement, S. L., Ranga Rao, G. V., Romies, J., Miles, M., and Bouhssini,
M. 2007. Host plant resistance and insect pest management in chickpea.
Pages 520-537 in: Chickpea Breeding and Management, S. S.Yadav, R. J.
Redden, W. Chen, and B. Sharma, eds. CAB International, Oxfordshire, UK.

Todd, E. L., and Poole, R. W. 1980. Keys and illustrations for the
armyworm moths of the noctuid genus *Spodoptera* Guenée from the Western
Hemisphere. Annals. Entomol. Soc. Amer. 73:722-738.

Wakamura, S., and Takai, M. 1992. Control of the beet armyworm in open
fields with sex pheromone. Pages 115-125 in: Diamondback Moth and other
Crucifer Pests, Talekar, N. S., ed. Asian Vegetable Research and
Development Center (AVRDC), Taipei, Taiwan.

(Prepared by R. Ahmad, P. Durai Murugan, P. Arjuna Rao and S.L. Clement)

![](media/image23.jpeg){width="3.0in" height="1.9930555555555556in"}

Fig. 1. Eggs of *Spodoptera exigua.* (Courtesy ICRISAT)

![](media/image24.jpeg){width="3.0in" height="1.9930555555555556in"}

Fig. 2. Larva of *Spodoptera exigua* feeding on chickpea. (Courtesy
ICRISAT)

![](media/image25.jpeg){width="3.0in" height="2.05in"}

Fig. 3. Fourth instar larvae of *Spodoptera praefica* feeding on
chickpea. (Courtesy W. Kaiser)

![](media/image26.jpeg){width="3.0in" height="1.9930555555555556in"}

Fig. 4. Eggs of *Spodoptera exigua.* (Courtesy ICRISAT)

![](media/image27.jpeg){width="3.0in" height="1.9930555555555556in"}

Fig. 5. Adult of *Spodoptera exigua.* (Courtesy ICRISAT)

![](media/image28.jpeg){width="3.0in" height="2.2736111111111112in"}

Fig. 6. Dead first instar larvae of *Spodoptera exigua* on resistant
hybrid chickpea. (Courtesy L. Elberson).

**The Black Cutworm: *Agrotis ipsilon***

The black cutworm, *Agrotis ipsilon* (Lepidoptera: Noctuidae) is a
polyphagous pest on several field crops. The other two species, *Agrotis
flammatra* and *A. spinifera* are of minor importance.

**Geographic distribution**

*Agrotis* species are widely distributed world over, and are pests of
several crops in different agro-ecosystems. *Agrotis ipsilon* is widely
distributed in Europe, North Africa, Syria, Japan, China, Indonesia,
Australia, New Zealand, Hawaii, Sri Lanka, Myanmar, and India. In
Europe, it is found in almost all countries extending from Ireland and
Portugal in the west to Bulgaria and Romania in the east, and from
Norway and Finland in the north to Sicily in the south. It is
widespread, and more damaging in the northern hemisphere than in the
southern hemisphere. It annually reinvades temperate areas,
overwintering in warmer or subtropical regions. Long distance dispersal
of adults has long been suspected in Europe, China, and North America.
The cutworms move north in the spring and south in autumn. Studies in
the United States demonstrated northward displacement of moths during
the spring in the range of 1,000 km in two to four days when assisted by
northward wind. Similar displacement to south and southwest has been
documented in the autumn.

**Host range**

It feeds on chickpea, pea, lentil, potato, and other crops in North
India. It also damages cauliflower, cotton, maize, strawberry, tomato,
grapevine, alfalfa, rice, sorghum, sweet potato, tobacco, wheat,
mustard, linseed, sugarcane, cucurbits, poppy, beet, soybean, cabbage,
clover, sunflower, artichoke, pepper, vegetables, and several weeds.
Among weeds suitable for larval development are *Poa pratensis, Rumex
crispus, Chenopodium album, Barbarea vulgaris,* and *Amaranthus
retroflexus*.

**Nature of damage**

The larvae feed on leaves, stems, and roots of many field crops,
including chickpea and lentil. Up to 10% plant damage has been recorded
at 40 days after crop emergence in chickpea. The older larvae cut the
plant above the root crown (Fig. 1). Most of the plant is not consumed
after cutting, and larvae move to another plant leaving the earlier one
to wither and dry. Some species feed on the upper leaves before moving
to the soil surface (Fig. 2). Heavy damage of cutworms occurs in areas
that remain flooded during the rainy season.

**Life cycle**

The moths are grey-brown with a wingspan of 40 to 50 mm (Fig. 3).
Forewings are light brown, patterned with an ipsilon shape, and the
hindwings are creamy white with brown edges. The moths appear in the
plains in October and come out at dusk and fly until darkness sets in.
After mating for 4 to 6 days, the female begins to oviposit at night and
lay creamy-white, dome shaped eggs in clusters. These are laid singly or
in small groups on moist soil, weeds, or leaves of host plants.
Oviposition continues for 5 to 11 days. The pre-oviposition and
oviposition periods vary from 3.9 to 5.5 and 5.8 to 8.3 days,
respectively. A female may lay 639 to 2252 eggs. Eggs are sub-spherical
usually dome shaped with 30 to 40 longitudinal ribs and about 0.5 mm in
diameter. The freshly laid eggs are cream colored and turn reddish
yellow and finally blackish before hatching. The egg, larval, and pupal
periods vary from 2.7 to 5.1, 18.2 to 39.5, and 31.4 to 69.8 days,
respectively. Larval mortality is as high as 70% during the early
instars. In summer, it survives on the weeds in wastelands. The young
larvae are nocturnal, pale yellowish green with a blackish head, and
pro-notal shield. The larva is covered with small dusky tubercles, each
bearing a seta. When full grown, the larva is 40 to 50 mm long, slate
grey or dark green in color with shiny and greasy skin. Each body
segment has four or more distinct spots on each side. On the dorsal
surface, along the median lines, two bright lines run longitudinally.
The ventral surface is light grey. Larvae remain below the surface of
the ground, under clods of soil, or other shelters during the day. The
first two larval stages feed on the foliage. The third and later stages
often become cannibalistic and adopt solitary habits. They grow as much
as 50 mm in length and can be found coiled round a damaged plant. Mature
larvae burrow deeply into the ground and pupate within cells (Fig. 4),
from which the adults emerge. Depending on the climate, there may be one
or several generations in a year.

**Management**

**Population monitoring**. Control measures can be undertaken based on
economic thresholds. The economic threshold is 1.0 larva per 10 plants
in northern plains of India. Light traps can be very effective for
monitoring as well as managing the cutworms. However, light traps are
not consistently efficient. In USA, light traps are most effective in
the summer and autumn. Pheromone traps are more effective during the
spring flight, when the larvae present the greatest threat to young
plants. The yellow white traps captured significantly more male moths.
Large larvae burrow in the soil, and are difficult to observe. However,
larvae can be sampled with bait traps and this is the most effective
means for monitoring cutworms prior to crop emergence or planting of
seedlings. The trap has a container, which is baited with fresh plant
material and/or bran with vermiculite so that the larvae can take
shelter. Larvae are captured in baited containers more effectively if
the vermiculite is not too near the surface. Trapping efficiency is
enhanced if a screen cylinder, which provides a visual stimulus to the
cutworms, is suspended above the baited container. If plants are present
in the field, they compete with the bait in the traps and trap
efficiency declines markedly.

**Cultural control.** Deep plowing of the fields before planting and
after crop harvest reduces cutworm damage, which turn up the larvae and
pupae to the soil surface, making them susceptible to predators and
sunshine. The plants at times are able to recover from cutworm damage.
The abundance of *A. ipsilon* in some areas is partly affected by
rainfall. In the drier areas (e.g., Syria) infestations are higher in
years with high rainfall. In fact, flooding fields is recommended as a
control measure in some cases. Weed hosts in outlying areas are often
preferred sites of oviposition and serve as food for the cutworms during
the off-season.

**Natural enemies.** In India, the braconids; *Microgaster* sp., *Bracon
kitcheneri*, and *Fileanta ruficanda*. parasitize the cutworm larvae,
while *Broscus punctatus* and *Liogryllus bimaculatus* are common
predators. *Apanteles ruficrus* is another important parasitoid. In USA,
among the wasps known to attack this cutworm are braconids: *Apanteles
marginiventris*, *Microplitis feltiae*, *Microplitis kewleyi*, *Meteorus
autographae*, and *Meteorus leviventri*; ichneuminids: *Campoletis
argentifrons*, *Campoletis flavicincta*, *Hyposoter annulipes*, and
*Ophion flavidus*. Larvae parasitized by *M. leviventris* consume about
24% less foliage and cut about 36% fewer seedlings. Overall parasitism
could be as high as 69%. *Amara impuncticollis* and *Philonthus* sp*.*
predate on all instars of *A. ipsilon*.

**Biopesticides.** *Metarhizium anisopliae* and nuclear polyhedrosis
virus have been found to be effective in reducing the population of
cutworm. An entomopathogenic nematode, *Steinernema carpocapsae* strain
has also been found to be effective. In central USA, another
entomopathogenic nematode, *Hexamermis arvalis* is known to parasitize
60% larvae.

**Chemical control.** Cutworms are often difficult to control,
especially when populations are epidemic in proportion. Large
populations may cause severe crop damage with indications that the pest
is the black cutworm. Unfortunately, by the time the pest is identified,
the cutworms would have already developed into a life stage, which is
not as susceptible to insecticides as the early larval stages. The
sporadic nature of cutworm populations can make preventive treatments
futile. Their soil-dwelling habits beneath foliage make the control
difficult with insecticides since they do not reach the target. One way
to control cutworms is to broadcast a poison bait prepared with wheat
bran, cotton or groundnut cake, moistened with water, and trichlofon,
carbaryl, or parathion at 10 kg ha^-1^ in the evening. Endosulfan dusts
or sprays, and endosulfan bait have also been found to be effective for
cutworm control. Carbaryl, phosalone, chlorpyriphos, quinalphos, phorate
and carbofuran are the other effective insecticides.

# **Selected References**

Ahmad, R. 2003. Insect pests of chickpea and their management. Pages
229-260 in: Chickpea Research In India. Indian Institute of Pulses
Research, M. Ali, S. Kumar, and N. B. Singh, eds. Kanpur, Uttar Pradesh,
India.

Amitava, K., Paul, S., Basu, A., and Chettri, M. 2003. Field evaluation
of bio-pesticides and insecticides against cutworm in plains of West
Bengal. *Indian J. Potato* 30:155-156.

Chaudhary, J. P., and Malik, V. S. 1981. Extent of damage and screening
of dust formulations of insecticides against gram cutworm *Agrotis
ipsilon* Hufn. on gram, *Cicer arietinum* L. Indian J. Entomol.
43:153-157.

Hill, D. S. 1983. *Agrotis ipsilon* (Hfn.). Pages 357-358 in:
*Agricultural Insect Pests of the Tropics and Their Control*. 2nd Ed.
Cambridge University Press, Cambridge, UK.

Kumar, K., Sinha, R. B. P., and Sinha, P. K. 1983. Testing of different
insecticidal formulations against greasy surface caterpillars, *Agroris
ipsilon* Rutt. J. Soil Biol. Ecol. 3:35-38.

# Nair, M. R. G. K. 1975. *Insects and Mites of Crops in India*. Indian Council of Agricultural Research, New Delhi, India. 405 pp.

(Prepared by R. Singh and S.K. Singh)

![](media/image29.jpeg){width="2.3756944444444446in"
height="1.5763888888888888in"}

Fig 1. Nature of damage and the larvae of black cutworm, *Agrotis
ipsilon.* (Courtesy ICRISAT)

![](media/image30.jpeg){width="1.8833333333333333in"
height="1.9944444444444445in"}

Fig. 2. Black cutworm, *Agrotis ipsilon* larva feeding chickpea foliage.
(Courtesy ICRISAT)

![](media/image31.jpeg){width="2.498611111111111in"
height="1.4652777777777777in"}

Fig. 3. Adult of black cutworm, *Agrotis ipsilon.* (Courtesy ICRISAT)

![](media/image32.jpeg){width="2.3756944444444446in"
height="1.5763888888888888in"}

Fig. 4. Pupa of black cutworm, *Agrotis ipsilon.* (Courtesy ICRISAT)

**Leafminers: *Liriomyza cicerina* and *Chromatomyia horticola***

Chickpea leafminer, *Liriomyza cicerina* (Diptera: Agromyzidae) is an
important pest of chickpea in West Asia, North Africa, and southern
Europe. *Chromatomyia horticola* is a polyphagous pest and feeds on
greenhouse plants in Italy, and United Kingdom. It also attacks chickpea
in Spain and India.

**Geographical distribution**

*Liriomyza cicerina* causes heavy losses to chickpea in West Asia, North
Africa, and southern Europe. In the Mediterranean region, the chickpea
leafminer occurs in high densities every year. *Chromatomyia horticola*
is widely distributed in Europe, Asia, Africa, North America, Australia,
Pacific Islands, and New Zealand.

**Host range**

*Liriomyza cicerina* is a major pest of chickpea in West Asia and North
Africa. It has also been reported to feed on lentil, pea, faba bean,
groundnut, beet, capsicum, *Brassica* spp., *Allium* spp., and several
other crop and weed hosts. *Chromatomyia horticola* has been reported to
feed on chickpea, field beans, alfalfa, faba bean, field beans, lablab,
lentil, and mung bean.

**Nature of damage**

The females puncture the upper surface of chickpea leaflets with the
ovipositor and feed on the exudates, which results in a stippled pattern
on the leaflets. In some feeding punctures, the eggs are inserted just
under the epidermis. The leafminer larvae feed in the leaf mesophyll
tissue forming a serpentine mine, which later becomes a blotch (Fig. 1).
The mining activity of the larvae reduces photosynthetic capacity of the
plant, and under heavy infestation, may cause desiccation and premature
leaf fall. Leafminer damage at times may destroy young seedlings,
resulting in leaf drop and reduces the crop yield. Up to 30% yield
losses have been reported in chickpea due to leafminer damage in Syria.
The larvae of *C. horticola* make tunnels in the leaves. The second
generation of *C. horticola* has been reported to damage sunflower in
Hungary. Although the damage is very conspicuous, most of the time it is
not economically significant.

**Life cycle**

The wingspan of *L. cicerina* varies from 1.3 to 1.6 mm. Legs are
normally brown and striated, but several specimens may have yellowish
legs (Figs. 2 and 3). It completes the life cycle in 20 to 30 days. For
egg laying, the female makes a tubular leaf puncture to deposit the eggs
singly. Some punctures are without eggs, which are considered as feeding
punctures. The larva begins feeding immediately after eclosion and feeds
on the mesophyll tissue until it is ready to emerge from the leaf after
four larval moults (Fig 2). When the larva is ready to pupate, it cuts a
semicircular slit in the leaf surface at or near the end of the mine and
falls down from the leaf to the ground to pupate in the soil (Fig. 2).
It has an obligatory diapause. There are 3 to 4 generations in a year,
and the larvae are present throughout the vegetative period of host
plants. Maximum damage has been recorded during pod formation causing 31
to 86% loss in leaf mass, and 33% yield loss in grain yield. In Syria,
the first generation emerging from diapause appears in late March, and
the second-generation reaches peak in the mid May. With the maturity of
the chickpea plants, the leafminers disappear. Leaf mining at times does
not influence yield, but reduces seed weight.

The *C. horticola* is active from December to April-May in India, and is
believed to pass the rest of the period in the pupal stage in the soil.
The adults emerge in the beginning of December. A female may lay 290 to
353 eggs singly, in the leaf tissues. The eggs hatch in 2 to 4 days and
the larvae feed between the lower and upper epidermis by making zigzag
tunnels. The larvae complete development in about 3 to 6 days, and
pupate within the galleries. The larvae are minute, slender, and can be
seen feeding within the tunnels. The adults emerge from the pupae in 6
days and life cycle is completed in 13 to 16 days. There are 4 to 5
generations from December to May, and maximum incidence has been
observed in March in India. Longevity of the males is longer when fed on
sugar or honey. On the other hand, sugar or honey-fed females lived
shorter lives than those fed on the host plant. The fecundity increases
with an increased availability of sugar or honey. Females of *C.
horticola* prefer to lay eggs on leaves already infested with the same
species than the un-infested leaves.

**Management**

Cultural manipulation, scouting, monitoring of insect pest populations
and their natural enemies, use of natural plant products,
bio-pesticides, and need-based insecticide application can be taken up
for management of agromyzid flies. Integrated pest management approaches
have been evaluated for controlling leafminers, especially *L. cicerina*
with minimum effects on natural enemies of the pest. Yellow moistened
traps are more effective in capturing adults of *L.* *cicerina* than the
Tullgren funnels or net sweeps.

**Cultural control.** Early-plantings of chickpea in winter are less
damaged by the leafminer compared to the spring-sown crop. The second
generation of the leafminer coincides with young seedlings of the
spring-sown chickpea plants (in contrast to the winter-sown crop); hence
the higher level of infestation in spring-sown chickpea. Different
combinations of fertilizers have no effect on the leafminer incidence.
Spreading of harvested plants on plastic sheets to facilitate collection
of pupae and deep plowing are safe control methods against the pupae.

**Host plant resistance.** Varieties with larger leaflets are more
preferred by the leaf miner than those with small leaflets. Oxalic acid
content in chickpea leaves has been reported to be correlated with the
level of resistance to leafminer (Fig. 4). Over 200 accessions of eight
wild *Cicer* species have been evaluated for resistance to *L. cicerina*
under natural infestation. Two accessions of *C. cuneatum* (ILWC 40 and
ILWC 187) and 10 accessions of *C. judaicum* were found to be highly
resistant to leafminer damage. Another 18 lines of *C. judaicum*, four
of *C*. *pinnatifidum,* and *C. reticulatum* showed moderate levels of
resistance. ILWC 39, ILC 3800, ILC 5901, and ILC 7738 are resistant to
leafminer damage. These sources of resistance have been used
successfully at the International Center for Agricultural Research in
Dry Areas (ICARDA) in breeding chickpea cultivars with resistance to
leaf miner. Seven of these lines (FLIP 2005-1C, FLIP 2005-2C, FLIP
2005-3C, FLIP 2005-4C, FLIP 2005-5C, FLIP 2005-6C, and FLIP 2005-7C)
have good agronomic background, seed size, and plant type, and have been
distributed to national programs for evaluation under local conditions.

**Biological control.** Six braconids and 12 eulophids parasitize *L.
cicerina* of which, *Diglyphus isaea* is the most common parasitoid. It
has been commercialized in China for the management of leafminers on
other crops. *Opius monilicornis* parasitized up to 29.8% larvae of *L.
cicerina* in chickpea fields (Fig. 5). In Spain, *O. monilicornis* and
*D. isaea* are major parasitoids of chickpea leafminer. In Syria, *O.
monilicornis* has been found to be the most effective parasitoid of
leafminer in chickpea, parasitizing up to 70% larvae. The majority of
the larval parasitoids (*D. isaea*, *Hemiptarsenus* *zilahisebessi*,
*Chrysonotomyia* sp., *Pnigalio* sp., and *Cirrospilus* sp.) emerge
during the active season of leafminers, including *L.* *cicerina*, and a
good synchronization has been observed between the time of host
emergence and abundance of the larval parasitoid, *D. isaea*.

*Pseudopezomachus masii*, *Aphidius* sp., *Chrysocharis* *pubicornis*,
*Neochrysocharis formosa*, *C. lyonitae*, *D. isaea,* *Pediobius
acanthi,* and *Sphegigaster* spp. have been reported parasitizing *C.
horticola*. *Dacnusa rodriguezi* has been found to be an important
parasitoid of *C. horticola*. *Dacnusa sibirica* and *D. isaea* have
also been reported parasitizing larvae of *C. horticola* in chickpea.
The parasitism of *C. horticola* is greater in the sub-humid than in the
sub-arid regions. The hymenopteran parasitoids, *D. horticola*,
*Hemiptarsenus indicus* and *Pediobius* *indicus* are associated with
*C.* *horticola* in India.

**Chemical control**. To control *L. cicerina* in chickpea, application
of monocrotophos or endosulfan at the flowering stage has been found to
be most effective. Neem seed kernel extract (3 and 4 kg per 10 L water)
has been found to be effective against the larvae of *L. cicerina* for
15 days. Neem oil (azadirachtin 0.03% w/w) (40 ml per 20 L water),
sprayed 3 to 5 times at weekly intervals, starting at about flowering
stage, effectively reduced the mining caused by the chickpea leafminer.
Neem sprays had little effect on the level of parasitism. The plots
sprayed with deltamethrin (50% EC) (25 ml per 100 L water) reduced the
level of parasitism by about 50%. Methamidaphos is quite effective
against *L. cicerina* on chickpea.

*Chromatomyia horticola* could be effectively controlled by dimethoate,
monocrotophos, methyl-parathion, phosphamidon, and promecarb.
Phosphamidon and dimethoate also reduced the numbers of pupae.
Oxy-methyl demeton (0.025%) and dimethoate (0.03%) are very effective in
checking the leaf infestation and maggot population. Decamethrin 0.008%,
fenvalerate 0.02%, and cypermethrin 0.01% are more effective than
oxy-methyl demeton (0.025%) and dimethoate (0.03%) in managing *C.
horticola*. Adults of *C. horticola* are highly susceptible to
isoxathion, chlorpyriphos, ethofenprox, emamectin benzoate, and
pyridalyl.

**Integrated pest management**. Early plantings are less damaged by *L.
cicerina* in West Asia. Monitoring of pest populations and their natural
enemies should be undertaken for effective use of natural plant products
and bio-pesticides under lower population densities. This will be useful
for the encouragement of natural enemies. Leafminer-resistant varieties
should be planted in areas endemic to leafminer damage. Need-based
insecticide application can be taken up for the control of leafminers
under heavy infestations.

# **Selected References**

Anonymous. 1965. *Phytomyza atricornis* (Distribution map). Distribution
Maps of Plant Pests. (December) Map 205. Commonwealth Agricultural
Bureau International, Wallingford, Oxfordshire, UK.

Bene, G. del. 1989. Natural enemies of *Liriomyza trifolii* (Burgress),
*Chromatomyia horticola* (Gourreau) and *Chromatomyia syngenesiae* Hardy
(Diptera: Agromyzidae) in Tuscany. Redia 72:529-544.

El Bouhssini, K. Mardini, K., Malhotra, R. S., Joubi, A., and Kagka, N.
2008. Effects of planting date, varieties and insecticides on chickpea
leaf miner (*Liriomyza cicerina* R.) infestation and the parasitoid
*Opius monilicornis* F. Crop Prot. 27:915-919.

El-Serwy, S. A. 2003. Studies on the leafminers: *Liriomyza cicerina*
(Rondani) and *Liriomyza bryoniae* (Kaltenbach) (Diptera: Agromyzidae)
and their parasitoids in faba bean in Egypt. Egyptian J. Agric. Res.
81:1581-1593.

Pardo, J., Tormos, J., and Jimenez, R. 2000. Morphological details of
the final larval instar of *Dacnusa* *rodriguezi* Docavo &Tormos
(Hymenoptera: Braconidae) parasitoid species of *Chromatomyia horticola*
(Goureau) (Diptera: Agromyzidae). Fragmenta-Entomologica 32:299-303.

Parrella, M. P. 1987. Biology of *Liriomyza.* Annu. Rev. Entomol.
32:201-224.

Singh, K. B., and Weigand, S. 1996. Registration of three
leafminer-resistant chickpea lines:ILC 3800, ILC 5901 and ILC 7738. Crop
Sci. 36:472.

Uygun, N., Polatoz, Z., and Baspinar, H. 1995. Faunistic studies on
Agromyzidae (Diptera) in the South-East Mediterranean region of Turkey.
Turk. Ent. Derg. 19:123-136.

Weigand, S. 1990. Insect pests of chickpea and in the Mediterranean area
and possibilities for resistance. Options Mediterraneennes- Series A-
Seminaires- Mediterraneennes-No. 9:73-76.

(Prepared by R. Lal, H. R. Rohilla, and M. El Bouhssini)

![](media/image33.jpeg){width="2.3756944444444446in"
height="0.9451388888888889in"}

Fig. 1. Leaf mining (Right -- leaf mining. Left -- larva coming out of
the mine) by chickpea leaf miner, *Liriomyza cicerina.* (Courtesy
ICARDA).

![](media/image34.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 2. Adult, pupal, and larval stages of the chickpea leaf miner,
*Liriomyza cicerina.* (Courtesy ICARDA).

![](media/image35.jpeg){width="2.5in" height="1.1944444444444444in"}

Fig. 3. Adult of *Liriomyza cicerina.* (Courtesy ICARDA)

![](media/image36.jpeg){width="2.0in" height="1.5131944444444445in"}

Fig. 4. Varietal reaction to damage by chickpea leaf miner, *Liriomyza
cicerina.* (Courtesy ICARDA)

![](media/image37.jpeg){width="2.0in" height="1.3666666666666667in"}

Fig. 5. Parasitoid, *Opius monilicornis* laying egg inside the chickpea
leaf miner, *Liriomyza cicerina.* (Courtesy ICARDA)

**Black Aphid: *Aphis craccivora***

The black aphid, *Aphis craccivora* (Hemiptera: Aphidiidae) is an
important pest of chickpea in the Indian Subcontinent, and is an
occasional pest in the Pacific Northwest USA. *Aphis craccivora* is also
important as a vector of the chickpea stunt disease. It is commonly
known as cowpea aphid, black aphid, black legume aphid, groundnut aphid,
bean aphid, lucerne aphid, black lucerne aphid, and African bean aphid.

**Geographic distribution**

##  The black aphid, *A. craccivora* is distributed in Europe, Asia, Africa, Australasia and Pacific Islands, North and Central America, and South America. It is widely distributed in the tropics, where it is one of the most common aphid species.

## 

## Host range

##  *Aphis craccivora* has an extensive host range, and damages several grain legumes including chickpea, lentil, cowpea, groundnut, alfalfa, and other legumes. It also infests *Glaricidia* sp., *Capsella bursa-pastoris*, *Chenopodium album*, *Polygonum lapathifolium*, and *Rumex crispus*. *Aphis craccivora* is a key pest of lentil in Castilla La Mancha (central Spain).

#### Nature of damage

Both nymphs and adults suck the plant sap from the tender leaves, stems
and pods, and mostly colonize the young leaves and growing points, which
become characteristically deformed (Fig. 1)*.* Yield can be drastically
reduced, and if infestations are early and severe, plants can be killed
completely. Infestations during the bloom and early pod stages reduce
yield and crop quality by removing plant sap, impairing pod appearance,
reducing seed fill, and by the presence of aphid honeydew. Plants with
high populations also have reduced ability to fix nitrogen. In many
areas, *A. craccivora* is more important as a vector of viruses than as
a direct plant feeder. While feeding, the aphid produces considerable
amount of honeydew upon which sooty mold grows. The black sooty mold
reduces photosynthesis and may make leaves unpalatable to livestock. The
honeydew also makes the plants sticky, which causes problems during
harvesting and threshing.

**Life cycle**

The black aphid, *A. craccivora* is readily distinguishable from other
aphids. It is a relatively small insect, and the adults are usually
shiny black, while the nymphs are slate gray. The appendages are usually
whitish with blackish tips. The life cycle of *A. craccivora* passes
through nymph to adult stage, which is completed in 8 to 10 days. The
aphids are active throughout the year, and a female can produce over 100
nymphs in 15 days. There are several generations in the tropics. Winged,
sexual forms of *A. craccivora* also occur in some regions, e.g., in
India, but not in others.

**Management**

**Monitoring.** Aphid infestation should be monitored regularly,
particularly when the crops begin to flower. Action thresholds are 1 to
2 aphids per leaf, 2 to 3 aphids per stem tip, or 9 to 13 aphids per
sweep, if a sweep net is used for sampling.

**Cultural Control.** Early sowing leads to early canopy closure, which
also helps in reducing the spread of viruses in chickpea. Aphid
infestation is greater under wider spacing.

**Biological control.** *Aphis craccivora* is attacked by several
parasites and predators. Ladybirds, *Coccinella septempunctata, C.
transversalis, C. nigritis, Menochilus sexmaculatus* (Fig. 2), and
*Brumus suturalis*; and chrysopids, *Chrysoperla* spp. are common aphid
predators. The parasitoids such as *Trixys indicus* and *Lipolexix
scutellaris* are important natural enemies of *A. craccivora* in India.
Although parasitism as high as 95% has been documented, aphid population
levels can be so high that enough non-parasitized individuals remain to
cause significant injury.

**Biopesticides.** Plant products such as azadirachtin, neem oil, and
pyrethrins have shown some suppression of *A. craccivora* populations,
but do not provide effective control of the aphids.

**Host plant resistance.** A number of genotypes have been reported to
be less susceptible to aphid damage. Chickpea varieties with low
trichome density or those devoid of trichomes, and low acid exudates are
highly susceptible to aphid damage.

**Chemical control**. Generally, there is no need for aphid control on
chickpea in India, but chemical control may become necessary to prevent
secondary spread of the chickpea viruses. A number of insecticides such
as methomyl, oxy-demeton methyl, and monocrotophos are effective for
aphid control. *Aphis craccivora* has also developed resistance to some
commonly used insecticides in India.

**Management.** Biological control with predatory coccinellids has a
considerable potential for management of aphids. Aphids infest lentil
fields only at later stages of crop development (podding), where no
spray is necessary. If infestation occurs before pod maturity,
insecticide sprays may be given. Aphids are attacked by a number of
natural enemies, especially coccinellids. Although the larvae and adults
feed on aphids, the extremely fast reproduction rate of the aphids
prevents the coccinellids from reducing infestation levels sufficiently.
Under heavy infestation, application of selective insecticides for aphid
control may be necessary.

###### Selected References

CABI (Commonwealth Agricultural Bureau International). 1983. *Aphis
craccivora*. Distribution Maps of Plant Pests. Map 99. Commonwealth
Agricultural Bureau International, Wallingford, UK.

Dhingra, S. 1994. Development of resistance in the bean aphid, *Aphis
craccivora* Koch. to various insecticides used for nearly a quarter
century. J. Entomol. Res. 18:105-108.

Kaiser, W. J., Ghanekar, A. M., Nene, Y. L., Rao, B. S., and Anjaiah, V.
1990. Viral diseases of chickpea. Pages 139-142 in: Chickpea in the
Nineties: Proceedings of the Second International Workshop on Chickpea
Improvement, 4-8 December 1989, International Crops Research Institute
for the Semi-Arid Tropics (ICRISAT), Patancheru, Andhra Pradesh, India.

Lal, S. S., Yadava, C. P., Dias, C. A. R., and Nigam, R. 1989. Effect of
planting density and chickpea cultivars on the infestation of black
aphid, *Aphis craccivora* Koch. Madras Agric. J. 76:461-462.

Loss, S., Brandon, N., and Siddique, K. H. M. (Eds.). 1998. The Chickpea
Bulletin 1326. Western Australian Department of Agriculture, Perth,
Australia. 76 pp.

Perez Andueza, G., Mozos Pascual de los, M., and Portillo Rubio, M.
2004. Main pests of lentil (*Lens culinaris* Medikus) in Castilla La
Mancha (central Spain): crop losses and influence on yield parameters.
Boletin de Sanidad Vegetal, Plagas 30:763-772.

Reed, W., Cardona, C., Sithanantham, S., and Lateef, S. S. 1987. The
chickpea insect pests and their control. Pages 283-318 in: The Chickpea,
M. C. Saxena and K. B. Singh, eds. CAB International, Wallingford, UK.
pp. 283-318.

Singh, R., and Tripathi, N. 1987. Record of parasitoids from Tarai belt
of eastern Uttar Pradesh. J. Aphidol. 1:89-92.

Singh, S. R., and Van Emden, H. F. 1979. Insect pests of grain legumes.
Annu. Rev. Entomol. 24:255-278.

Waterhouse, D. F. 1998. Biological Control of Insect Pests: Southeast
Asian Prospects. Australian Council of International Agricultural
Research (ACIAR) Monograph No. 51.

(Prepared by H. C. Sharma, M. K. Dhillon and J. Romeis**)**

![](media/image38.jpeg){width="2.0in" height="2.546527777777778in"}

Fig. 1. Black aphid, *Aphis craccivora* infesting chickpea. (Courtesy
ICRISAT)

![](media/image39.jpeg){width="2.0in" height="1.3666666666666667in"}

Fig. 2. Coccinellid beetle feeding on *Aphis craccivora* in chickpea.
(Courtesy ICRISAT)

Pea Aphid: *Acyrthosiphon pisum*

Pea aphid, *Acyrthosiphon pisum* (Hemiptera: Aphididae), commonly known
as pea aphid, green pea aphid, and green alfalfa aphid, is an important
pest of lentils. It is a key pest of lentil in Castilla La Mancha
(central Spain), and is also an important pest of lentil in India.

**Distribution**

## The pea aphid, *A. pisum* has a worldwide distribution, and is a major pest of lentil in America, Europe, Africa, Australia, and Asia. 

**Host range**

Many species of grain legumes including pea, lentil, alfalfa, faba bean,
*Melilotus officinalis*, *Ononis repens*, *Sarothamnus scoparius*,
*Trifolium* spp., *Vicia cracca*, etc. are important hosts of pea aphid,
*A. pisum*.

**Nature of damage**

Both nymphs and adults suck the plant sap from the tender leaves, stems
and pods, and mostly colonize the young leaves and growing points, which
become characteristically deformed*.* Yield may be drastically reduced
if infestations are early and severe. Aphid infestations during the
bloom and early pod stages reduce yield and crop quality by removing
plant sap, impairing pod development, reducing seed fill, and by the
presence of aphid honeydew. Plants with high aphid populations also have
a reduced ability to fix nitrogen. Pea aphids can transmit many viruses
including pea enation mosaic virus, even to crops that they rarely feed
on, such as cucurbits.

**Life cycle**

Pea aphid, *A. pisum* is 3 to 4 mm long, greenish, has long legs,
and reproduces parthenogenetically (females produce offspring without
mating). The aphid has viviparous colonies (Fig. 1) of wingless females
(apterae), which later convert to winged forms for further dispersion.
The winged females (alatae) of this aphid are carried by the wind to the
host plants. Live female nymphs (no egg stage necessary) are produced
throughout the year. Females take 9 to 12 days to complete development.
There are three moults, and each female produces up to 150 nymphs. There
are 13 to 20 generations per year. Adults may or may not have wings, but
produce winged individuals for dispersion if conditions become crowded
or plants become desiccated. Sexual forms occur in the fall, and eggs
are laid after mating. Populations tend to be lower after cold, snowless
winters, or springs with persistent wet weather. The net reproductive
rate and post-reproductive period, adult longevity, survival, fecundity,
and the lifespan have been found to be significantly longer on lentils
than on other crops. Daily nymph production is significantly correlated
with minimum temperature.

**Management**

**Monitoring.** Start aphid monitoring when plants begin to flower.
Action thresholds vary from 1 to 2 aphids per leaf, 2 to 3 aphids per
stem tip, or 9 to 13 aphids per sweep, if a sweep net is used for
sampling.

**Cultural control.** No specific cultural practices have been
recommended for aphid control in lentil. However, harvesting or spraying
nearby alfalfa, vetch, or clover before winged adults are formed in the
spring minimizes aphid damage.

**Host plant resistance.** Varieties differ in their susceptibility to
pea aphid damage. Varieties with aphid resistance should be planted in
areas endemic to infestations by this aphid. The genotype, French Indigo
has high levels of antibiosis, while Spanish Brown has high levels of
antixenosis and tolerance to *A. pisum*.

**Biological control.** There are many natural enemies such as predatory
coccinellids, lacewings, flower fly larvae, predatory midges, and
braconid wasps that help reduce aphid numbers. However, under heavy
infestation, natural enemies may not provide adequate control. Fungi
such as *Pandora neoaphidis*, can control high aphid populations during
warm, humid, and wet weather.

**Chemical control**. In case of severe infestations, foliar application
of dimethoate, pirimicarb or other systemic insecticides provides
adequate control.

**Integrated pest management.** Host plant resistance and biological
control with predatory coccinellids has a potential for the management
of aphids in lentil. Aphids often infest lentil fields only at later
stages of crop growth (podding), where no spray is necessary. If
re-infestation occurs before pod maturity, insecticide sprays may be
given. Aphids attract a number of natural enemies, especially
coccinellids. Although the larvae and adults feed on aphids, the
extremely fast reproduction rate of the aphids prevents the coccinellids
from reducing infestation levels sufficiently.

###### **Selected References**

Andarge, A., and Van Der Westhuizen, M. C. 2004. Mechanisms of
resistance of lentil *Lens culineris* Medicus, genotypes to the pea
aphid *Acyrthosiphon pisum* Harris (Heniptera: Aphididae). Int. J. Trop.
Insect Sci. 24:249-254.

Aydin, H., Muehlbauer, F. J, and Kaiser, W. J. 1987. Pea enation mosaic
virus in lentil (*Lens culinaris*). Plant Dis. 71:635-638.

CABI (Commonwealth Agricultural Bureau International). 1982.
*Acyrthosiphon pisum*. In: Distribution Maps of Plant Pests, December
1982, Map 23. Commonwealth Agricultural Bureau International,
Wallingford, UK.

Lal, S. S. 1992. Insect pests of lentil and their management: review.
Agric. Rev. 13:225-232.

Perez Andueza, G., Mozos Pascual de los, M., and Portillo Rubio, M.
2004. Main pests of lentil (*Lens culinaris* Medikus) in Castilla La
Mancha (central Spain): crop losses and influence on yield parameters.
Boletin de Sanidad Vegetal, Plagas 30:763-772.

Wale, M., Jembere, B., and Seyoum, E. 2000. Biology of the pea aphid,
*Acyrthosiphon pisum* (Harris) (Homoptera: Aphididae) on cool-season
legumes. Insect Sci. Appl. 20: 171-180.

Weigand, S., Lateef, S. S., El Din Sharaf, N., Mahmoud, S. F., Ahmed,
K., and Ali, K. 1994. Integrated control of insect pests of cool season
food legumes. Pages 679-674 in: Expanding the Production and Use of Cool
Season Food Legumes, F. J. Muehlbauer and W. J. Kaiser, eds. Kluwer
Academic Publishers, Dordrecht, The Netherlands.

Yadava, C. P., and Ahmad, R. 2000. Insect pests of lentil and their
management. Appl. Entomol. 2:82-95.

(Prepared by J. Romeis, M.K. Dhillon, and R. Ahmad)

![](media/image40.jpeg){width="2.0in" height="1.3666666666666667in"}

Fig. 1. Pea aphid, *Acyrthosiphon pisum* infesting lentil. (Courtesy
Jorg Romeis)

**Lygus bugs: *Lygus* spp.**

Tarnished plant bug, *Lygus lineolaris* and the western tarnished plant
bug, *L. hesperus* (Hemiptera: Miridae) are pests of lentil in western
USA. Feeding by *Lygus* bugs produces depressed, chalk-colored lesions
(chalky spot) on the lentil seeds where the bug has fed. These bugs are
pests of lentil throughout the Palouse area (eastern Washington and
northern Idaho) in the USA. In years where pea aphid populations are
very low, early sprays are not applied and this results in greater
damage by the *Lygus* bugs. Scouting for *Lygus* bug is critical to
prevent damage to the crop. Close examination of the plants is the best
way to see *Lygus* bugs, which are present under the curly leaves of the
lentil plants during the daytime, and rarely seen on the visible
portions of the crop.

**Distribution**

Tarnished plant bug, *L. lineolaris* (Fig. 1) is distributed in North
America, Central America, and the Caribbean; while the western tarnished
plant bug, *L. hesperus* (Fig. 2), is distributed in North America, The
European tarnished plant bug, *L. pratensis*, is distributed throughout
Europe and Asia (Afghanistan, China, Persia, Turkey).

## 

**Host range**

*Lygus hesperus* is a polyphagous pest on several crops and weeds.
Alfalfa is the primary host of western tarnished plant bug, *L.
hesperus*, which is also an important pest of sorghum in USA. *Lygus
lineolaris* is polyphagous and attacks a wide range of fruits,
vegetables, and ornamentals. Economic losses have been recorded in
cotton, alfalfa, snap bean, lima bean, soybean, lentil, tomato, and
nursery stock. *Lygus rugulipennis* damages *Vicia sativa*, *Trifolium
pratense*, *Medicago sativa*, red clover, etc. *Lygus pratensis* damages
a wide range of field and garden crops.

## Nature of damage

The *Lygus* bugs puncture the tissue and feed on immature reproductive
structures causing \"chalky spot syndrome\" (physical deformations) on
lentil seeds, which increases the prevalence of shriveled, unfilled
pods, and seed abortion. Incidence of growing-point injuries by *Lygus*
spp. is a serious problem on cauliflower in Sweden. The bugs are most
active during the early-morning, and cause most damage to newly emerging
seedlings.

## Life cycle

##  The adult bugs are about 6 mm long and 2.5 mm wide, flattened, oval, and of various colors from pale green to yellowish brown, and have a distinctive triangle or \"V\" mark about one-third of the distance down the back, just in front of the wings. The legs and antennae are relatively long. The young nymphs are pale-green in color and look like aphids, except that they are more active. Older nymphs are quite variable in color, and are similar to adults. The older nymphs develop prominent black dots on the top of the thorax and abdomen. The adults overwinter in protective hedgerows, fallen leaves, and other plant debris. Emerging in mid-April to late-May, they feed on weed hosts. The eggs are laid on the stems and petioles and the egg incubation period last for about 10 days. The 1^st^ generation adults appear in mid-June to mid-July. They leave the host weeds, and disperse into the adjacent crops. The second generation nymphs feed on the host plants during August, causing economic damage. A smaller third generation may emerge in September. 

##  Overwintering adults of *L. lineolaris* can be found in dead weeds, leaf litter, under tree bark, in rock piles in fields, timber margins, stream banks, and roadside. Adults become active in early spring and feed on newly developing buds and shoots. Eggs are often deposited in leaf petioles or at the base of the leaf blade, but the preferred location varies with the crop attacked (Fig. 3). Eggs are usually deposited singly, but occasionally more than one egg may be found at an oviposition site. The eggs hatch in about 7 to 10 days. The newly hatched nymph (Fig. 4) is about one mm long, yellowish-green nymphs emerge and begin feeding (Fig. 5). There are five nymphal stages. The life cycle is completed in 3 to 4 weeks. There are 2 to 3 generations per year. Population peaks for adults generally occur in early July, early August, and early September. 

**Management**

Although *Lygus* bugs are not serious pests of lentil, and do not
require any control measures, an understanding of management practices
well in advance would be helpful in managing them on lentil as and when
they reach economic threshold levels.

**Cultural control.** Strip cutting is used to limit *Lygus* migration
from forage lucerne to susceptible neighboring crops when the lucerne is
harvested. The localized treatment of alfalfa strips significantly
reduces the extent of damage on lettuce plots adjacent to the trap
crops. Removal of plant materials following seed harvest during late
summer does not provide significant control of bugs for the following
year. No-tillage encourages population buildup of *L. lineolaris* in
maize and soybean. Alfalafa can be used as a strip cop in cotton to keep
*L. lineolaris* population under check. Some of these practices may be
suitably modified to reduce the damage by *Lygus* bugs in lentil.

**Biological control.** Several parasitoids (*Anaphes iole*, *Peristenus
stygicus*, and *Leiophron uniformis)*, and predators (*Geocoris
punctipes*, *Orius insidiosus*, and *Chrysoperla* spp.) can be used for
biological control of *Lygus* spp. *Anaphes iole* is a solitary egg
parasitoid of *Lygus* bugs in North America. *Phasia obesa* is parasitic
on *Lygus* adults, and *P. pallipes*, *P. relictus*, and *P. varisae*
are parasitic on nymphs. *Peristenus digoneutis* and *P. stygicus* have
been used for biological control of *Lygus* species in North America.

**Host plant resistance.** Host plant resistance will be useful for
increasing the effectiveness and longevity of other *Lygus* control
measures. Although the *Lygus* spp. have not been reported as serious
pests of lentil, host plant resistance could be an important tool for
the management of this pest in crops where it has been reported as a
major pest such as cotton. Cotton genotype Arkot A306 has been reported
to be resistant to *L. lineolaris*. The use of *Bacillus thuringiensis*
(*Bt*) transgenic cotton has reduced insecticide application against
bollworm complex, and has also reduced *Lygus* populations.

**Chemical control.** Malathion, dimethoate, trichlorfon, and
fluvalinate are effective for the control of *Lygus* spp. Oxadiazine has
been reported to be effective in conservation of beneficials in a cotton
ecosystem and is safer to farm labor and the environment.

**Biopesticides.** The entomopathogenic fungus, *Beauveria bassiana*
isolate MK 2001 (1 x 10^7^ conidia ml^-1^) is highly pathogenic to
adults of *L. lineolaris* (84% mortality). Application of fungal isolate
MK 2001, 2 to 4 days before applying the fungicides metalaxyl, mancozeb,
or copper oxide synergized the insecticidal effect of this fungus.
However, application of fungicides prior to MK 2001 fungal isolate
antagonized the insecticidal effect. *Beauveria bassiana* can be used
for the control of *L. rugulipennis* in the greenhouse. A combination of
*B. bassiana* and the antifeedant chemical, pymetrozine reduced the
numbers and damage by *L. rugulipennis* in cucumber. Some of these
measures can be adapted for controlling *Lygus* bugs in lentil.

**Integrated pest management.** Adoption of cultural approaches to
manage *Lygus* spp. will be largely influenced by the availability and
efficacy of management practices, including insecticides. Sprays of *B.
thuringiensis* in alfalfa have been found to be effective against the
first three nymphal instars of *L. hesperus*, but the adults are
tolerant to *Bt* sprays. Traps baited with live *L. rugulipennis*
females attract conspecific males, which can form the basis for timely
control measures. Use of entomopathogenic fungus *B. bassiana* is quite
effective against the tarnished plant bugs. Mowing of alternate rows for
weed management, and preventing mirids from migrating from adjoining
areas is effective for managing *L. rugulipennis*. Although several
parasitoids and predators can be used for biological control of *Lygus*
bugs, there is a need to develop mass rearing techniques for these
natural enemies. A combination of sanitation practices and insecticide
applications is quite effective to control the *L. lineolaris*. Use of
insect-nets and yellow chromotropic traps is another effective method of
preventing the damage by *Lygus* bugs. These methods can be modified for
use for the control of *Lygus* bugs in lentil.

### Selected References

Bourland, F. M., and Smith, C. W. 2001. Registration of Arkot A306 and
Arkot A314 germplasm lines of cotton. Crop Sci. 41:2008-2009.

CABI (Commonwealth Agricultural Bureau International). 1954. *Lygus
pratensis*. *Distribution Maps of Plant Pests.* Map 39. Commonwealth
Agricultural Bureau International, Wallingford, Oxfordshire, UK.

CABI (Commonwealth Agricultural Bureau International). 2000. *Lygus
lineolaris*. *Distribution Maps of Plant Pests*. Map 604. Commonwealth
Agricultural Bureau International, Wallingford, Oxfordshire, UK.

Godfrey, L. D., and Leigh, T. F. 1994. Alfalfa harvest strategy effect
on *Lygus* bug (Hemiptera: Miridae) and insect predator population
density: implications for use as trap crop in cotton. Environ. Entomol.
23:1106-1118.

Jacobson, R. J., and Enkegaard, E. 2002. *Lygus rugulipennis* Poppius
(Het., Miridae): options for integrated control in glasshouse-grown
cucumbers. Bulletin OILB SROP 25:111-114.

Ramert, B., Hellqvist, S., and Petersen, M. K. 2005. A survey of *Lygus*
parasitoids in Sweden. Biocontr. Sci. Tech. 15:411-426.

Smith, R. A., and Nordlund, D. A. 2000. Mass rearing technology for
biological control agents of *Lygus* spp. Southwestern Entomol.
(Supplement) 23:121-127.

Tanigoshi, L. K., Mayer, D. F., Babcock, J. M., and Lunden, J. D. 1990.
Efficacy of the beta-exotoxin of *Bacillus thuringiensis* to *Lygus
hesperus* (Heteroptera: Miridae): laboratory and field responses. J.
Econ. Entomol. 83:2200-2206.

(Prepared by Margaret L. Allen)

![](media/image41.jpeg){width="2.0in" height="1.179861111111111in"}

Fig. 1. *Lyzus lineolaris* adults. (Courtesy Cathe L. Smith)

![](media/image42.jpeg){width="2.0in" height="1.1333333333333333in"}

Fig. 2. *Lyzus hesperus* adults. (Courtesy Cathe L. Smith)

![](media/image43.jpeg){width="2.0in" height="1.6in"}

Fig. 3. Eggs of *Lygus* *lineolaris.* (Courtesy Cathe L. Smith)

![](media/image44.jpeg){width="2.0in" height="1.5534722222222221in"}

Fig. 4. *Lygus* *lineolaris* nymph hatching from the egg. (Courtesy
Cathe L. Smith)

![](media/image45.jpeg){width="2.0in" height="1.6in"}

Fig. 5. Young nymph of *Lygus* *lineolaris* feeding on host plant.
(Courtesy Cathe L. Smith)

**Thrips: *Frankliniella occidentalis, Kakothrips robustus, and Thrips
angusticeps spp.***

Several species of thrips including *Frankliniella occidentalis,
Kakothrips robustus, Thrips angusticeps,* and *Thrips tabaci*
(Thysanoptera: Thripidae) damage grain legumes, including lentil.

**Geographic distribution**

*Frankliniella occidentalis* (Fig. 1) is commonly known as western
flower thrips. The majority of the species under *Frankliniella* are
present either in North or South America, but a few are cosmopolitan.
*Frankliniella* *schultzei* is known as cotton bud thrips, and is
important in Asia as a vector of tomato spotted wilt virus (TSWV).
*Kakothrips robustus* is known as pea thrips, which is widely
distributed in Europe. It attacks several crops such as lathyrus,
lucerne, mustard, and horse-bean in Poland, Hungary, Romania, Bulgaria,
and Czechoslovakia. *Thrips tabaci* and *Thrips angusticeps* are
commonly known as onion or potato thrips. The genus *Thrips* comprises
of several hundred species, which are polyphagous, and cosmopolitan in
distribution. *Thrips tabaci* and *T. angusticeps* have been reported as
key pests of lentil in Central Spain.

**Host range**

*Frankliniella* *occidentalis* is a polyphagous pest, and has a wide
host range including cereals, legumes, ornamentals, and fruit trees
(grapes, citrus, peach, plum, strawberries, raspberries, etc.). This
species is of worldwide importance as a vector of TSWV in a number of
crops. *Kakothrips robustus* is polyphagous, and infests a number of
cultivated and weed hosts. *Thrips tabaci* and *T. angusticeps* infest
onion, tobacco, cotton, tobacco, peas, brassicas, watermelon, mango,
gladiolus, banana, cucurbits, oil palm, etc. Besides infesting
cultivated crops, these species also feed on several weed hosts. *Thrips
tabaci* and *T. angusticeps* are important pests of lentil in Central
Spain, and affect the productivity and quality of seeds.

**Nature of damage**

Most of the *Frankliniella* spp. prefers flowers (Fig. 2), but in the
absence of flowers, they also feed on foliage (Fig. 3). When the
populations of thrips are high, the plant terminals may blacken and
wither. Feeding by thrips on young leaves results in silvery streaks on
the opened leaves, distortion, or curling of leaves. In case of severe
infestation, the leaf area is reduced, which indirectly affects
photosynthesis and grain yield.

**Life cycle**

The life history of thrips involves an egg, two active larval instars
that feed, followed by two (or three) relatively inactive pupal instars
that probably do not feed, and the adults. One or both sexes may be
wingless. In most phytophagous thrips, the eggs are inserted into the
tissue of green plants. The males are generally haploid and derived from
unfertilized eggs. Despite this sex-determination process known as
haplodiploidy, many unrelated species have evolved the ability to
reproduce in the absence of males. The cytological mechanisms involved
are not fully understood, but parthenogenesis is important when the pest
species are introduced to new areas. Sexual dimorphism is usually
evident and is sometimes remarkable. Males of flower- and leaf-living
species are usually smaller than the females, but males of
fungus-feeding species are often much larger than females. Many
phlaeothripid species produce different sized morphs, with and without
wings. These forms often exhibit bizarre patterns of allometric growth,
particularly of the forelegs, and as a result, different morphs are
considered to represent unrelated species. The adults and larvae of many
species of thrips are found only in flowers, where they feed by sucking
out the contents of pollen grains, and probably, the cell sap of other
flower tissues around the bases of the anthers and on the developing
fruits. Some flower-living species also feed on leaves, but most
leaf-feeding species are restricted to leaves. Some thrips are
associated with very young leaves, and these insects are often very
small and active. Under optimum conditions, one generation may be
completed in 12 to 15 days. *Kakothrips robustus* is known to have only
one generation a year in England with established diapause. Fully-grown
larvae usually enter the soil in mid-summer, pupate early in the
following spring, and emerge as adults in late spring. *Thrips tabaci*
and *T. angusticeps* have 1 or 2 generations in temperate regions and
undergo diapause. However, there are multiple generations and no
diapause in the tropics.

**Management**

**Cultural practices:** Agronomic practices such as adjusting planting
date, seed rate, and intercrops play a significant role in suppressing
thrips populations. Climatic conditions such as mild winter (10 to
12^0^C night temperatures as in case of southern India), which are
unfavorable to crops, are favorable to thrip population build up.
Avoiding planting thrip susceptible crops during the vulnerable phase
would help in minimizing inputs on plant protection.

**Host plant resistance:** Resistant sources can play a significant role
in the management of thrips in several crops. However, there is no such
information on host plant resistance to thrips in lentil.

**Biological control:** *Entomophthora* spp. epizootic on *T. tabaci*
were noticed in the Netherlands in glasshouse situation on eggplants.
*Entomophthora parvispora* occurs widely throughout central and southern
Europe. However, the fungus has not been successfully multiplied. Since
bio-agents have not successfully kept the pest under manageable levels,
it has been recommended to use selective chemicals for the management of
thrips. Application of the nematode, *Steinernema feltiae* against *F.
occidentalis* decreased the population, but did not provide adequate
control. Introduction of *Amblyseius* spp. shortly after thrip invasion
in cucumber provides satisfactory control of *T. tabaci,* suggesting the
feasibility of biocontrol.

**Chemical control:** Several researchers have shown efficient
management of thrips as well as diapausing populations by foliar and
soil application of insecticides. Systemic insecticides such as
dimethoate, monocrotophos, and imidacloprid; and insecticides with
fumigant action such as dichlorvos have been found to be effective
against a range of thrips species. However, several thrips species have
been reported to develop resistance to insecticides. There is 30-fold
difference in resistance to malathion and 25-fold difference to
dichlorvos in western flower thrips in UK. There is no resistance to
insecticides in *F. occidentalis* to abamectin, methiocarb, and
pyrazophos in Australia. However, build-up of resistance to
chlorpyriphos, dichlorvos, and malathion has been recorded. Acephate,
dimethoate, endosulfan, fipronil, methamidophos, methidathion, and
spinosad are quite effective for controlling thrips. Resistance to
imidacloprid, dimethoate, and thiamethoxam has been observed in *T.*
*tabaci* in India.

**Integrated pest management:** Several management strategies have been
developed for the management of thrips on several crops such as
manipulating sowing dates, proper spacing, intercropping, utilizing
resistant sources, use of biocontrol agents, disposal of plant residues,
and need based application of pesticides. However, most of these are
location specific.

**Selected References**

Herron, G. A., and James, T. M. 2005. Monitoring insecticide resistance
in Australian *Frankliniella occidentalis.* Pergande (Thysanoptera:
Thripidae) detects fipronil and spinosad resistance. Australian J.
Entomol. 44:299--303.

Lewis, T. 1997. Pest thrips in perspective. Pages 1-13 in: Thrips as
Crop Pests, T. Lewis, ed. Commonwealth Agriculture Bureau International,
Wallingford, UK.

McDonald, C. O. 1995. Response of western flower thrips to dichlorovos
and malathion in the United Kingdom. Pages 347-350 in: Thrips Biology
and Management, B. L. Parker, M Skinner and T. Lewis, eds. Plenum Press,
New York.

McLeod, D. M., Tyrrell, D., and Carl, K. P. 1976. *Entomophthra
provispora* sp. nov., a pathogen of *Thrips tabaci*. Entomophaga
21:307-312.

Mound, L. A. 1976. The identity of the greenhouse thrips, *Heliothrips
haemorroidalis* (Bouche) (Thysanoptera) and the taxonomic significance
of spanandric males. Bull. Entomol. Res. 66:179-180.

Palmer, J. M., and Mound, L. A. 1978. Nine genera of fungus-feeding
Phlaeothripidae (Thysanoptera) from the Oriental region. Bull. British
Mus. Nat. Hist. Entomol. 37:153-215.

Perez Andueza, G., Mozos, P.M. de los, and Portillo Rubio. M. 2004. Main
pests of lentil in Castilla La Mancha: crop losses and influence on
yield parameters. Boletin de Sanidan Vegetal Plagas 30:763-772.

### (Prepared by G. V. Ranga Rao)

![](media/image46.jpeg){width="2.625in" height="1.8076388888888888in"}

Fig. 1. Western flower thrips, *Frankliniella occidentalis* adult.
(Courtesy ICRISAT)

![](media/image47.jpeg){width="2.0694444444444446in"
height="2.6368055555555556in"}

Fig. 2. Western flower thrips, *Frankliniella occidentalis* feeding on
flowers. (Courtesy ICRISAT)

![](media/image48.jpeg){width="2.3756944444444446in"
height="1.8180555555555555in"}

Fig. 3. Western flower thrips, *Frankliniella occidentalis,* feeding on
foliage. (Courtesy ICRISAT).

**Green Stink Bug: *Nezara* *viridula***

Green stinkbug, *Nezara viridula* (Hemiptera: Pentatomidae), is a highly
polyphagous pest on several grain legumes including lentil. The nymphs
and adults suck the sap from leaves and developing pods.

**Distribution**

Green stinkbug, *N. viridula* is distributed all over the world. Several
authors have depicted eastern Palearctic or Indo-Malayan region as the
ancestral home of *N. viridula.* The *N. viridula* has also been
recorded on soybean in north eastern Brazil, and is spreading in the
Neotropical region. It is also known as green vegetable bug, southern
green stinkbug, eastern green stinkbug, and pentatomid bug.

## Host range

The green stinkbug, *N. viridula,* is a highly polyphagous pest of many
important agricultural crops and weeds. It feeds on several plant
species including cotton, sunflower, soybean, mungbean, clover,
fababean, lentil, wheat, barley, flax, sugarbeet, okra, aubergine,
pepper, tomato, cowpea, cabbage, cauliflower, potato, beans, and peas.
Wild crucifers and leguminous plants are alternate hosts of *N.
viridula*. This species has been reported as a minor pest on lentil.

**Nature of damage**

The nymphs and adults suck the sap from leaves, stems, and pods,
resulting in malformation or drying up of the pods (Figs. 1 and 2). In
lentil, it sucks the sap from shoots and pods. The *N. viridula* bugs
cause varying degrees of damage from seedling stage where the young
growing tips of plants dry up, until crop harvest, especially during
reproductive phase, when they feed on the pods.

## 

## Life cycle

The adults are 14 to 18 mm long, green and shield-shaped with a small
head (Fig. 2). There are five nymphal instars. The first instar nymphs
cluster on or near the egg masses, and apparently do not feed. After the
first moult, the nymphs begin to disperse and feed on leaves and pods.
During the summers, the development period from egg to adult varies
between 35 to 37 days, depending on temperature. A constant temperature
of 28°C has been reported to be optimal for the growth and development
of *N. viridula*. The egg incubation period ranges from 5 to 7 days. The
1^st^, 2^nd^, 3^rd^ 4^th^, and 5^th^ nymphal instars complete
development in 3 to 5, 5 to 7, 7 to 9, 8 to 11, and 11 to 13 days,
respectively. The entire life cycle is completed in 37 to 52 days. A
female lays an average of 215 eggs (20 to 425 eggs per female). The
host - weed phenology, host abundance and diversity, and natural enemies
influence the population dynamics of *N. viridula*. With a reduction in
flowering and fruiting stages of crop hosts/weeds, *N. viridula*
migrates to other hosts, and may cause heavy damage.

**Management**

The green stinkbug is not a serious pest of lentil, and does not require
pesticide application to reduce the damage. Management practices would
be helpful in minimizing *N. viridula* damage in lentil when the
population density exceeds economic threshold levels.

**Cultural practices.** The stinkbugs prefer soybean over cotton, and
therefore, soybean can be used as a trap crop for attracting stinkbugs.
Narrow-row plantings harbor more *N. viridula* than the wide-row
plantings. The nymphal population of *N. viridula* has been found to be
greater in crops with high phosphorus levels, while potassium or
magnesium did not affect the population density.

**Biological control.** The most successful biological control agents
are the scelionid egg parasitoids, *Trissolcus basalis*, and *Trichopoda
pilipes*, which attack adults and large nymphs of *N. viridula*. Of the
egg parasitoids, *T. basalis* is most important and distributed widely.
*Trichopoda giacomellii* has also been reported to be an important
parasitoid on the eggs of *N. viridula* in Australia. There are a number
of predators on *N. viridula*, but they are nonspecific to this pest.
*Solenopsis invicta* is the dominant predator on eggs and early stage
nymphs. Grasshoppers are important egg predators during the reproductive
stages of the plant. *Trissolcus megallocephalus* has been reported as
an important egg parasitoid of *N. viridula* from Egypt. Parasitism by
*T. giacomellii* plays an important role in regulation of the *N.
viridula* populations in Argentina.

**Biopesticides and natural plant products.** The entomopathogenic
fungi, *Beauveria bassiana*, *Metarrhizium anisopliae*, and
*Paecilomyces lilacinus* result in high mortality of this insect under
laboratory conditions, but are not very effective under field
conditions. Sprays of *Aglaia odorata,* *Annona squamosa*, *A.
muricata*, and *Melia azedarch* leaf extracts are moderately effective.
Application of the *Carica papaya* leaf extract results in a significant
reduction in *N. viridula* populations.

**Host plant resistance.** The hemipterans are difficult to control with
insecticide sprays, and host plant resistance may help solve this
problem. Since *N. viridula* is a serious pest at fruiting/podding
stage, identification of genotypes with resistance to feeding on pods is
important along with the plant traits that make the insects restless on
the host plant. At the moment, there are no reports on identification of
resistance to green stinkbug in lentil.

**Chemical control.** Prophylactic insecticide applications result in
superior economic outcomes as compared with a strategy based on economic
thresholds. Cypermethrin is quite effective for controlling the
stinkbugs. Foliar sprays of cypermethrin + dimethoate result in better
control of *N. viridula* as compared to application of neem or
*Pongamia* kernel extracts. Endosulfan and λ-cyhalothrin are also
effective for controlling this pest.

**Integrated pest management.** Field scouting, identification of
different stages of the pest and its natural enemies, and knowledge of
injury thresholds at different crop stages are important for developing
effective management strategies. A management package involving early
planting, close spacing, and need-based insecticide application at
budding, flowering, or podding stages have been found to be effective in
controlling this pest on cowpea, and possibly, can be adapted on lentil
with suitable modifications. Biological, cultural, and chemical
approaches have been adopted in Brazil to control *N. viridula* on
soybean. No-till system, early-maturing cultivars, and early-planting
have been recommended to reduce pesticide use. Use of natural plant
products and biopesticides may be explored as a safer alternate to
enhance the activity of natural enemies, in addition to identification
of genotypes with resistance to *N. viridula.*

### Selected References

Anyim, A. 2003. Effects of insecticidal treatment on the yield and
control of major insect pests of soybean (*Glycine max* (L.) Merrill) in
South-eastern Nigeria. Int. J. Agric. Rural Dev. 4:100-109.

CABI (Commonwealth Agricultural Buraux International). 1970. *Nezara
viridula*. In: Distribution Maps of Plant Pests. CAB International,
Nosworthy Way, Wallingford, Oxfordshire, OX10 8DE, UK.

Harris, V. E., and Todd, J. W. 1980. Temporal and numerical patterns of
reproductive behavior in the southern green stink bug, *Nezara viridula*
(Hemiptera: Pentatomidae). Entomol. Exp. Applic. 27:105-116.

McPherson, R. M., and Bondari, K. 1991. Influence of planting date and
row width on abundance of valvetbean caterpillars (Lepidoptera:
Noctuidae) and southern green stink bugs (Heteroptera: Pentatomidae) in
soybean. J. Econ. Entomol. 84:311-316.

Nabirye, J., Nampala, P., Ogenga Latigo, M. W., Kyamanywa, S., Wilson,
H., Odeke, V., Iceduna, C., and Adipala, E. 2003. Farmer-participatory
evaluation of cowpea integrated pest management (IPM) technologies in
Eastern Uganda. Crop Prot. 22:31-38.

Pitan, O. O. R., and Odebiyi, J. A. 2001. The effect of intercropping
with maize on the level of infestation and damage by pod-sucking bugs in
cowpea. Crop Prot. 20:367-372.

Sands, D. P. A., and Coombs, M. T. 1999. Evaluation of the Argentinian
parasitoid, *Trichopoda giacomellii* (Diptera: Tachinidae), for
biological control of *Nezara viridula* (Hemiptera: Pentatomidae) in
Australia. Biol. Cont. 15:19-24.

Todd, J. W. 1989. Ecology and behavior of *Nezara viridula*. Annu. Rev.
Entomol. 34:273-292.

(Prepared by M. K. Dhillon and R. Singh)

![](media/image49.jpeg){width="3.0in" height="1.9930555555555556in"}

Fig. 1. Nymph of *Nezara viridula* feeding on the pod. (Courtesy
ICRISAT)

![](media/image50.jpeg){width="3.0in" height="1.9930555555555556in"}

Fig. 2. Adult of *Nezara viridula* feeding on the pod. (Courtesy
ICRISAT)

### Integrated Pest Management

**Storage pest: Bruchids *Callosobruchus maculatus* and *Callosobruchus
chinensis***

Bruchids, *Callosobruchus chinensis* and *C.* *maculatus* (Coleoptera:
Bruchidae) are one of the most important pests of grain legumes in
storage, including chickpea and lentil. Bruchid infestation commences in
the field even before the crop harvest, and then they multiply quite
fast in storage, resulting in heavy losses. The *Callosobruchus* species
are commonly known as spotted pulse beetle, Oriental pulse beetle,
bruchid, bean weevil, bruchid seed beetle, gram bean weevil, southern
cowpea weevil, cowpea weevil, etc.

**Distribution**

*Callosobruchus chinensis* and *C.* *maculatus* are cosmopolitan in
distribution, encompassing Australia and Oceania, Europe, Asia, Africa,
and the Americas.

**Host range**

The members of the family Bruchidae have long been reported to destroy
the seeds of leguminous plants. They also feed on seeds and flowers of
non-leguminous plants belonging to the families Compositae, Malvaceae,
Convolvulaceae, Anacardiaceae, Rosaceae, Umbelliferae, Papavaraceae, and
Palmae. Among the several species of bruchids attacking edible legumes,
*C. maculatus* and *C. chinensis* are most destructive, and attack
almost all edible legumes, including chickpea and lentil.

**Nature of damage**

Females of *C. maculatus* and *C. chinensis* lay eggs singly on seeds,
which are visible to the naked eye (Fig. 1). Bruchids tend to lay eggs
singly on a given host and if all the seeds are occupied, then the
female starts laying eggs on already egg-laden seeds. The neonate larva
bores into the seed beneath the oviposition site, and completes its
development within a single seed. Damaged seeds are riddled with adult
emergence holes (Fig. 1), which are unfit for human or animal
consumption.

**Life cycle**

The life cycle of bruchids passes through five larval instars (Fig. 2),
pre-pupal, pupal (Fig. 3), and adult (Fig. 4) stages. The egg incubation
period of *C. maculatus* in green gram lasts for 3 to 5 days, and the
combined larval and pupal period lasts for nearly 19 days. Total
development is completed in about 24 days. The adults of *C. chinensis*
and *C. maculatus* are easily distinguishable with the naked eye (Fig.
5).

**Management**

**Pre-harvest control.** Monocrotophos (0.04%), fenvalerate (0.02%), and
dimethoate (0.03%) sprays at 45 and 50 days after flowering of pulses
reduces *C. chinensis* damage in stored pigeonpea. Similarly, cowpea
protected by spraying malathion (0.05%) at maturity reduces the bruchid
damage. At pod maturity, the dehisced pods with exposed seeds are more
vulnerable to oviposition by bruchids, and this problem can be avoided
by harvesting the crop at physiological maturity.

**Hermetic storage.** Storing grain legumes in polythene bags with a
cotton lining has been found effective against *C. maculatus*. Even
though the adult females could penetrate the polythene bags, the cotton
lining posed hindrance for oviposition, leaving the grains safe. Bagged
grains encased in polythene sacks are also less damaged by the bruchids.

**Solar treatment.** High temperatures due to solar radiation have been
found to kill developing bruchid larvae in the seeds. Exposing grain
legumes to 70 to 80ºC temperature for short durations has been found to
be effective in reducing the bruchid infestation. Solar heaters operated
at 65°C for 5 min provide 100% control of *C. maculatus.* Solar heat
treatment is being used in India for disinfesting the grains.

**Use of inert dusts:** Dusts remove the epicuticular lipid layer of the
insects and have been used effectively against bruchids. Some of the
dusts in use include bentonite clay, hydrated lime, attapulgite dust,
limes, clays, synthetic silica, and sand. Treatment of pulses with fly
ash also hinders emergence of *C. maculatus* adults up to 12 months.

**Use of traps.** Pitfall traps can be used for capturing insects that
are active on the grain surface, and in other layers of grain. It also
serves as a monitoring or mass trapping tool. Two models are available:
standard model (Fig 6a) and the TNAU model (Fig 6b). A standard model
has 2 parts, perforated lid (2 to 3 mm) and a cone-shaped bottom
portion. A special coating with sticky material on the inner side of
cone to hold the trapped insects is essential. The new model has a
perforated lid and a cone-shaped bottom, which tapers into a funnel
shaped trapping tube. It is made of plastic, and is simple and
economical. Another two-in-one trap (Fig 6c) is a combination of probe
and pitfall traps, and is designed to increase the trapping efficiency.
This trap is also suitable for pulse beetles as they are seen only on
grain surface. It does not require coating on the inner surface with
sticky materials. Beetles are captured alive in this trap. The release
of pheromone by the trapped insects attracts more insects.

**Chemical control.** Chemical methods such as fumigation with
phosphine, methyl bromide, or dusting with primiphos methyl and
permethrin are effective against bruchids, but have certain
disadvantages such as increased costs, handling hazards, pesticide
residue, and possibility of development of resistance. With the proposed
ban of fumigants by 2015, there is an urgent need to develop safer
alternatives to conventional insecticides and fumigants to protect
stored grain from insect pests.

**Host plant resistance.** The chickpea genotypes CPI 29973, CPI 29975,
CPI 29976, NCS 960003, K 902, CM 72, CMN 122, and BG 372 have been
reported to be resistant to *C. maculatus*. Apart from cultivated legume
species, wild relatives of several grain legumes have shown high levels
of resistance to bruchids. Several varieties have been developed by
crossing wild species with that of the cultigens, and the results have
been highly promising.

**Natural plant products.** Neem, *Azadirachta indica* possesses
antifeedant, oviposition repellant, ovicidal, and adulticidal properties
against bruchids. Seed treatment with neem leaf powder at 0.5 to 2.0 mg
100^-1^ g of grain, neem seed kernel powder, neem bark powder, and neem
seed oil at 3% have been reported to be effective against bruchids. Neem
oil and *Pongamia* oil reduce seed damage by *C. maculates.* Use of
coconut, mustard, and groundnut oil (12 ml^-1^ kg) to suppress adult
emergence for two consecutive generations has also been recpmmended.
Volatile oils of *Cymbopogan nardus*, *C. schoenanthus*, *Clausena
anisata, C. citratus,* and *Ocimum basilica* have oviposition repellent
and adulticidal effects. *Lantana camara* and *Parthenium hysterophorus*
have also been reported to be repellent to pulse beetle in chickpea.

**Biological control.** Bruchids are attacked by parasitoids belonging
to ten families of Hymenoptera and one of Diptera. Important biocontrol
agents of bruchids are listed in Table 1.

**Table 1. Natural enemies of bruchids.**

+-------------------------------+---------------------------------------+
| **Biocontrol agent**          | **Species attacked**                  |
+-------------------------------+---------------------------------------+
| *Anisopteromalis calandrae*   | *C. maculatus*                        |
|                               |                                       |
|                               | *C. chinensis*                        |
|                               |                                       |
|                               | *C. analis*                           |
+-------------------------------+---------------------------------------+
| *Dinarmus basalis*            | *Callosobruchus* spp.                 |
+-------------------------------+---------------------------------------+
| *D. acutus*                   | *C. maculatus*                        |
+-------------------------------+---------------------------------------+
| *Uscana lariophaga*           | *Callosobruchus* spp.                 |
+-------------------------------+---------------------------------------+
| *U. mukerjii*                 | *C. chinensis*                        |
+-------------------------------+---------------------------------------+
| *Pteromalus* spp.             | *C. maculatus*                        |
+-------------------------------+---------------------------------------+
| *P. cerealellae*              | *C. maculatus*                        |
|                               |                                       |
|                               | *C. chinensis*                        |
+-------------------------------+---------------------------------------+
| *Eupelmus* spp.               | *C. maculatus*                        |
+-------------------------------+---------------------------------------+
| *Pyemotes ventricosus*        | *Callosobruchus* spp.                 |
+-------------------------------+---------------------------------------+

**Integrated pest management.** Prevention of infestation in the field
through timely harvest of the crop or insecticide use can be quite
useful for reducing bruchid infestation. Optimum drying of the grain,
use of solar radiation to kill the bruchids infesting the grain, and
storing the grain in polyethylene bags can be used to reduce bruchid
infestation. Dusts, neem leaf or kernel powder, and treatment of seed
with neem, *Pongamia*, or other vegetable oils can be used for
minimizing the losses due to bruchids. Under severe infestation, the
grain should be fumigated to get rid of bruchids infesting the grain. An
ideal management schedule would be as follows:

-   Spraying monocrotophos or dimethoate at pod formation stage and just
    before pod maturity to infestation by bruchids in the field.

-   Reducing the moisture content of the grains to less that 12% before
    storage. This can be accomplished by sun drying.

-   Using polythene lined bags for storage.

-   Treatment of grains with inert dusts such as clay, silica, fine
    sand, or fly ash.

-   Use of pitfall traps as a low cost device for use in households and
    medium scale storage.

-   Treatment of grain with botanicals such as neem oil, neem leaf
    powder, etc.

Though the above methods are economical, developing varieties with
resistance to bruchids through interspecific hybridization and genetic
transformation would be helpful in the long run. Use of biocontrol
agents, pheromones, and controlled atmosphere storage (CAS) has been
found to be very effective, and can be used for minimizing the losses
due to bruchids.

## Selected References

Durairaj, C. and Muthiah, A. R. 2003. Evaluation of plant oils against
pulse beetle in mungbean. Page 246 in: Proceedings of the National
Symposium on Pulses for Crop Diversification and Natural Resource
Management, 20-22 Dec, 2003, Indian Institute of Pulses Research (IIPR),
Kanpur, Uttar Pradesh, India.

Gonzalez-Valenzuela, M., Roche, R., and Simanca, M. E. 1988.
Parasitation capacity of *Pteromalus* sp. in laboratory conditions.
Cien. Agric. 33:19-22.

Gupta, H. C., Gupta, I. J., and Sharma, S. N. 1997. Parasitic wasp,
*Dinarmus acutus* (Thompson) and *D. basalis* (Rondani) (Hymenoptera:
Pteromelidae) parasitizing *Callosobruchus maculatus* (F.). J. Biol.
Control. 11:77-78.

Islam, W. 1992. Mating and egg-laying behaviour of *Dinarmus basalis*
(Rondani) (Hymenoptera: Pteromalidae), A larval-pupal parasitoid of
*Callosobruchus chinensis* (Linnaeus) Ann. Entomol. 10:13-16.

Mohan, S., and Fields, P. 2002. A simple technique to assess compounds
that are repellant (or) attractive to stored product insects. J. Stored
Prod. Res. 38:23-31.

Murdock, L. L., and Shade, R. E. 1991. Eradication of cowpea weevil
infestations in cowpea by solar heating. American Entomol. 37:228-231.

Riaz, A., Aslam, M., and Suleman, N. 2000. Evaluation of resistance in
different chickpea strains to *Callosobruchus chinensis* (Linn.) under
laboratory conditions. Pak. J. Biol. Sci. 3(6):1033-1036.

Singh, D., and Mehta, S. 1998. Screening of plant materials for
repellant and insecticidal properties against pulse beetle
(*Callosobruchus chinensis*) and housefly (*Musca domestica*). J. Med.
Aromatic Pl. Sci. 20:397-400.

Wilkin, D. R., and Green, A. A. 1970. Polythene sacks for the control of
insects in bagged grain. J Stored Prod. Res. 6: 97-101.

(Prepared by C. Durairaj and T. Srinivasan)

![](media/image51.jpeg){width="2.75in"
height="1.59375in"}![](media/image52.jpeg){width="1.6215277777777777in"
height="1.6131944444444444in"}![](media/image53.jpeg){width="1.6256944444444446in"
height="1.6145833333333333in"}

Fig 1. Bruchid damage in chickpea. A: a severely damaged seed lot
(Courtesy C. Durairaj); B: solitary white eggs on damaged seeds
(Courtesy W. Chen), and C: a beetle emerges from a damaged seed
(Courtesy W. Chen).

![](media/image54.jpeg){width="2.0in" height="0.9in"}

Fig. 2. Fourth instar grub of *Callosobruchus chinensis.* (Courtesy C.
Durairaj)

![](media/image55.jpeg){width="1.5in" height="0.9798611111111111in"}

Fig. 3. Pupa of *Callosobruchus chinensis.* (Courtesy C. Durairaj)

![](media/image56.jpeg){width="1.0in" height="1.4736111111111112in"}

Fig. 4. Adult *Callosobruchus chinensis.* (Courtesy C. Durairaj)

![](media/image57.jpeg){width="2.0in" height="0.7666666666666667in"}

Fig 5. Species identity of bruchids. (Courtesy C. Durairaj)

![](media/image58.jpeg){width="2.0in" height="2.073611111111111in"}

Fig 6. Traps for bruchid management. (Courtesy C. Durairaj)

**Pest Management in Chickpea**

Chickpea is the third most important legume crop in the world, after dry
beans and peas. It is cultivated in 42 countries in South Asia, North
and Central America, the Mediterranean Region, West Asia, and North and
East Africa. In recent years, it has become an important crop in
Australia, Canada, and the USA. Nearly 60 insect species are known to
feed on chickpea. The important insect pests damaging chickpea in
different regions are: wireworms \[false wireworms - *Gonocephalum*
spp.\], cutworms \[black cutworm - *Agrotis ipsilon* and turnip moth -
*Agrotis segetum*\], termites \[*Microtermes obesi* and *Odontotermes*
sp.\], leaf feeding caterpillars \[cabbage looper - *Trichoplusia ni*,
leaf caterpillar - *Spodoptera exigua*, western yellow striped
armyworm - *Spodoptera praefica*, and hairy caterpillar - *Spilarctia
obliqua*\], semilooper \[*Autographa nigrisignia*\], leaf miners
\[*Liriomyza cicerina* and *L . congesta*\], pea leaf weevil - *Sitona
lineatus*, aphids \[*Aphis craccivora*\], nodule damaging flies
\[*Metopina ciceri*\], pod borers \[cotton bollworm - *Helicoverpa
armigera*, and native budworm - *Helicoverpa punctigera*\], and bruchids
\[Chinese bruchid - *Callosobruchus chinensis*, bean bruchid -
*Acanthoscelides obtectus*, pulse bruchid - *Callosobruchus analis*, and
pulse bruchid - *C. phaseoli*\]. Among the above pests the pod borer,
*H. armigera* and the aphid, *A. craccivora* are the major pests of
chickpea in the Indian subcontinent. In the Mediterranean region, the
most important pest is the leaf miner, *L. cicerina*. *Aphis craccivora*
is important as a vector of the chickpea stunt disease, while *C.
chinensis* is the most dominant pest species in storage. Various
components of pest management in chickpea are discussed below.

**Population monitoring and forecasting.** Efforts have been made to
develop a forecasting system, particularly for *H. armigera* in India. A
rule of thumb has been developed to predict *H. armigera* population
using surplus/deficit rainfall in different months in southern India. A
combination of surplus rains during the monsoon and deficit rainfall
during November indicated low incidence, while deficit rains during the
monsoon and surplus rains during November indicated severe attack.
Additional information on November rainfall gives precise information on
the level of attack (low, moderate, or severe). In Australia, population
monitoring with sex pheromone-baited traps is used to detect the onset
of immigration or emergence from local diapause. Abundance of *H.
armigera* and *H. punctigera* as measured by light traps has shown that
seasonal rainfall and local crop abundance give a reasonable prediction
of the timing of population events, and the size of subsequent
generations. Timing of control is determined by field monitoring of
larval densities in crops through the period of crop susceptibility.
Control is only recommended when larval populations in crops at the
post-flowering stage exceed the threshold of 2 to 4 larvae per meter of
row. There have been no efforts to study and develop population
prediction models for other insects infesting chickpea. There may be
need to know the conditions under which the leaf miner, *L. cicerina*
infestations cross the economic threshold levels, and the climatic
factors that may predict the onset of heavy infestations.

**Host plant resistance.** The development of chickpea cultivars
resistant or tolerant to insects has a major potential for use in
integrated pest management, particularly under subsistence farming
conditions in developing countries. More than 14,000 chickpea germplasm
accessions have been screened for resistance to *H. armigera* at
ICRISAT, India, under field conditions. Several germplasm accessions
(ICC 506EB, ICC 10667, ICC 10619, ICC 4935, ICC 10243, ICCV 95992, and
ICC 10817) with resistance to *H. armigera* have been identified, and
varieties such as ICCV 7, ICCV 10, and ICCL 86103 with moderate levels
of resistance have been released for cultivation. However, most of these
lines are highly susceptible to Fusarium wilt. Therefore, concerted
efforts are being made to break the linkage by raising a large
population of crosses between the lines with resistance to *H. armigera*
and the lines resistant to wilt.

Efforts are also currently underway at ICARDA to develop varieties with
resistance to the leaf miner, *L. cicerina*. Varieties with low trichome
density or devoid of trichomes are highly susceptible to aphid, *A.
craccivora* damage. A number of genotypes/lines have been reported to be
less susceptible to aphid damage. High levels of resistance have been
observed in desi type chickpeas to bruchids, *Callosobruchus* spp. Lines
showing resistance to bruchids usually have small seeds with a rough
seed coat. However, such grain is not acceptable to the consumers.
Chickpea seed that is split for *dhal* is unattractive to ovipositing
bruchid females, and therefore, processing the chickpea into split peas
or flour immediately after crop harvest can minimize the losses due to
these pests.

**Cultural manipulation of the crop and its environment**. A number of
cultural practices such as time of sowing, spacing, fertilizer
application, deep plowing, interculture, and flooding have been
documented by several workers to reduce insect damage. Inter-cropping or
strip-cropping with marigold, sunflower, linseed, mustard, and coriander
can minimize the extent of H. armigera damage to the main crop.
Strip-cropping also increases the efficiency of chemical control. Hand
picking of large-sized larvae can also be practiced to reduce pod borer
damage. However, the adoption of cultural practices depends on the crop
husbandry practices in a particular agro-ecosystem. Crop rotations do
not help manage the polyphagous and highly mobile insects such as
*Helicoverpa/Heliothis* species, although it has been noted that some
crops (e.g., alfalfa) are more attractive to the moths. Habitat
diversification to enhance pest control has been attempted in Australia.
An area-wide population management strategy has been implemented in
regions of Queensland and New South Wales to contain the size of the
local *H. armigera* population, and chickpea used as a trap crop plays
an important role in this strategy. Trap crops are managed in the same
way as commercial crops, but destroyed by cultivation before larvae
begin to pupate. The trap crops reduce the size of the local *H.
armigera* population before it can infest summer crops and start to
increase in size. As a result, the overall *H. armigera* pressure on
summer crops is reduced, resulting in greater opportunity for the
implementation of softer control options, reduced insecticide use, and
greater natural enemy activity. Plowing the fields before planting and
after crop harvest reduces damage by the *Helicoverpa* and cutworm,
*Agrotis ipsilon*. The plants at times are able to recover from foliar
damage by *H. armigera* and the cutworms. The black aphid, *A.
craccivora* incidence is greater under drought conditions and wider
spacing. Early sowing leads to early canopy closure, which also helps to
reduce virus spread in chickpea. Therefore, early sowing and optimum
planting densities can be used to minimize aphid infestation. Early-sown
crops also escape leafminer damage in West Asia.

**Biological control.** The importance of both biotic and abiotic
factors on the seasonal abundance of *H. armigera* is poorly understood.
Some parasitic wasps have been reported to avoid chickpea probably due
to their dense layer of trichomes and the acidic exudates. The egg
parasitoids, *Trichogramma* spp. are almost absent from chickpea
ecosystem. The ichneumonid, *Campoletis chlorideae* is probably the most
important larval parasitoid on *H. armigera* in chickpea in India.
*Carcelia illota*, *Goniophthalmus halli*, and *Palexorista laxa* have
also been observed to parasitize 3 to 54% larvae on chickpea. Predators
such as *Chrysopa* spp*., Chrysoperla* spp.*, Nabis* spp*., Geocoris*
spp*., Orius* spp*.,* and *Polistes* spp. are the most common in India,
attacking *Helicoverpa* spp. on chickpea and other crops. Provision of
bird perches or planting of tall crops that serve as resting sites for
insectivorous birds such as Myna (*Acridotheris tristis)* and Drongo
(*Dicrurus macrocercus)* helps to reduce the numbers of caterpillars.

Braconid wasps such as *Microgaster* sp., *Bracon kitcheneri*, and
*Fileanta ruficanda* parasitize the black cutworm, *A. ipsilon* larvae,
while *Broscus punctatus* and *Liogryllus bimaculatus* are common
predators. A parasitic wasp, *Opius* sp. feeds on the leafminer larvae
in West Asia, but further research is required before this insect can be
used for biological control in the field. Coccinellids and chrysopids
are common aphid predators, while *Trixys indicus* and *Lipolexix
scutellaris* are important parasitoids.

**Bio-pesticides and natural plant products.** The use of microbial
pathogens including *H. armigera* nuclear polyhedrosis virus (HaNPV),
entomopahogenic fungi, *Bacillus thuringiensis* (*Bt*), nematodes, and
natural plant products such as neem, custard apple, and karanj kernel
extracts have shown some potential to control *H. armigera*. HaNPV is a
viable option to control *H. armigera* in chickpea. Jaggery (0.5%),
sucrose (0.5%), egg white (3%), robin blue (0.1%), and chickpea flour
(1%) are effective in increasing the efficacy of HaNPV. The efficacy of
HaNPV in chickpea is increased by the addition of milk powder, and the
additive Aminofeed®. Entomopathogenic fungus, *Nomuraea rileyi* (10^6^
spores per ml) results in 90 to 100% larval mortality, while *Beauveria
bassiana* (2.68 x 10^7^ spores per ml) resulted in 6% damage in chickpea
compared to 16.3% damage in the untreated control plots. Formulations of
*Bt* used as a spray are effective to control the *H. armigera* and
semilooper, *A. nigrisigna*. Spraying *Bt* formulations in the evening
results in better control than spraying at other times of the day.
Spraying with neem seed kernel extract (5%) is effective for controlling
the leaf miner, *L. cicerina*, but the persistence is limited.

**Chemical control.** Management of *H. armigera* on chickpea and other
high-value crops relies heavily on insecticides. Cypermethrin,
profenophos, spinosad, methomyl, and indoxacarb are quite effective for
*H. armigera* control on chickpea. Spray initiation at 50% flowering is
most effective for the control of *H. armigera*. Use of indoxacarb in
chickpea in Australia is limited to one application with a cut-off date
for application to ensure that at least one generation of *H. armigera*
is not exposed to this insecticide before the commencement of its use in
summer crops such as cotton and mungbean.

Endosulfan dusts, sprays, or baits are effective for cutworm control.
Generally, there is no need for aphid, *A. craccivora* control on
chickpea, but chemical control may become necessary to prevent the
spread of the chickpea viruses. A number of insecticides such as
methomyl, oxy-demeton methyl, and monocrotophos are effective for aphid
control. Endosulfan, dichlorvos, and malathion have been recommended for
controlling the semilooper, *A. nigrisigna*.

**Integrated pest management (IPM).** The important insect pests
damaging chickpea that need to be controlled during different stages of
crop growth are cutworms (*A. ipsilon*), leaf feeding caterpillars (*S.
exigua*, *S. praefica, S. obliqua, A. nigrisignia*), leaf miners (*L.
cicerina* and *L. congesta*), aphids (*A. craccivora*), pod borers (*H.
armigera* and *H. punctigera*), and bruchids (*Callosobruchus* spp. and
*A. obtectus*). Control measures may also be undertaken based on pest
forecasting by pheromone or light trap catches (5 to 10 males per trap
per day). Pheromone or light traps, booby traps, visual scouting, and
sweep nets can be used to obtain an estimate of insect infestations to
decide the need to initiate control measures. For pod borers,
*Helicoverpa* spp., control measures may be undertaken based on economic
thresholds (one egg per plant or 2 - 3 small larvae per meter row).
Chickpea cultivars with resistance to insects can play major role in
integrated pest management, particularly under subsistence farming.
Varieties such as Vijay, Vishal, and ICCV 10 with low to moderate levels
of resistance to pod borers can be cultivated in India. Varieties with
resistance to leafminer and aphids have also been identified for use in
West Asia. High levels of resistance have been observed in *desi* type
chickpeas to bruchids, *Callosobruchus* spp.

Early plantings generally suffer low damage due to leafminer in West
Asia and the Americas. Early sowing leads to early canopy closure, which
also helps to reduce virus spread in chickpea. Therefore, early sowing
and optimum planting densities can be used to minimize aphid
infestation. Plowing the fields before sowing and after crop harvest and
flooding reduces the infestation and population carryover of pod borers
and soil dwelling insects. Inter-cropping or strip-cropping chickpea
with marigold, sunflower, linseed, mustard, and coriander can minimize
the extent of *H. armigera* damage to the main crop. Hand picking of
large-sized larvae can also be practiced to reduce pod borer damage.
Trap crops are managed in the same way as commercial crops, but
destroyed before the larvae begin to pupate.

The ichneumonid, *C. chlorideae* is the most important larval parasitoid
on *H. armigera*, while *Chrysoperla, Nabis, Geocoris, Orius,* and
*Polistes* are the common predators attacking *Helicoverpa* on chickpea
and other crops. Provision of bird perches or planting of tall crops
that serve as resting sites for insectivorous birds such as Myna and
Drongo helps to reduce the numbers of caterpillars. A number of natural
enemies have been reported in case of cutworms, aphids, and other
foliage feeders. However, except for aphids, natural enemies are not
very effective in reducing insect damage under field conditions.

Nuclear polyhedrosis virus (HaNPV) and formulations based on *Bacillus
thuringiensis* can be used for minimizing the damage by *Helicoverpa*,
and possibly other lepidopteran insects (*Spodoptera* spp.). Neem oil
(1%) and neem seed kernel extract (5%) (NSKE, 10 kg per ha) are also
effective against lepidopteran insects, leafminer, and the aphids. These
can be used alone in rotation with insecticides or in combination with
the commonly used insecticides. Endosulfan dusts or sprays or bait have
been found to be effective for the control of cutworms. Methomyl,
oxy-demeton methyl, malathion, and monocrotophos are effective for
controlling leaf feeders and aphids. Cypermethrin, fenvalerate,
methomyl, thiodicarb, profenophos, spinosad, and indoxacarb are
effective against pod borers and other leaf feeding insects.

**Conclusions**

Insect-resistant cultivars will form the backbone of integrated pest
management in future. The development and deployment of cultivars with
resistance to insects would offer the advantage of allowing some degree
of selection for specificity effects, so that pests, but not the
beneficial organisms are targeted. For pest management programs to be
effective in future, there is a need for in-depth understanding of the
population dynamics of insect pests to develop appropriate control
strategies, combine resistance to insects with resistance to important
diseases and cold tolerance, utilization of wild relatives to diversify
the genetic basis, and thus, increase the levels of resistance to the
target insect pests, identification of quantitative trait loci (QTLs)
associated with resistance to insects, develop insect-resistant
varieties through genetic transformation using genes with diverse modes
of action, and insecticide resistance management.

###### Selected References

Das, D. K., Trivedi, T .P., and Srivastava, C. P. 2001. Simple rule to
predict attack of *Helicoverpa armigera* on crops growing in Andhra
Pradesh. Indian J. Agric. Sci. 71:421- 423.

Dhingra, S. 1994. Development of resistance in the bean aphid, *Aphis
craccivora* Koch. to various insecticides used for nearly a quarter
century. J. Entomol. Res. 18:105-108.

Fitt, G. P. 1989. The ecology of *Heliothis* species in relation to agro
ecosystems. Annu. Rev. Entomol. 34:17-52.

King, A. B. S. 1994. *Heliothis/Helicoverpa* (Lepidoptera: Noctuidae).
Pages 39-106 in: Insect Pests of Cotton, G. A. Matthews and J. P.
Tunstall, eds. Commonwealth Agricultural Bureau International,
Wallingford, UK.

Lal, S. S., Yadava, C. P., Dias, C. A. R., and Nigam, R. 1989. Effect of
planting density and chickpea cultivars on the infestation of black
aphid, *Aphis craccivora* Koch. Madras Agric. J. 76:461-462.

Reed, W., Cardona, C., Sithanantham, S., and Lateef, S. S. 1987. The
chickpea insect pests and their control. Pages 283-318 in: The Chickpea,
M. C. Saxena and K. B. Singh, eds. Commonwealth Agricultural Bureau
International, Wallingford, UK.

Romeis, J., and Shanower, T. G. 1996. Arthropod natural enemies of
*Helicoverpa armigera* (Hübner) (Lepidoptera: Noctuidae) in India.
Biocont. Sci. Tech. 6:481-508.

Sharma, H.C. (ed.). 2005. *Heliothis/Helicoverpa* Management: Emerging
Trends and Strategies for Future Research. Oxford and IBH Publishers,
New Delhi, India. 469 pp.

Sharma, H.C., and Ortiz, R. 2002. Host plant resistance to insects: An
eco-friendly approach for pest management and environment conservation.
J. Environ. Biol. 23:11-35.

Zalucki, M. P., and Furlong, M. J. 2005. Forecasting *Helicoverpa*
populations in Australia: a comparison of regression based models and a
bioclimatic based modeling approach. Insect Sci. 12:45-56.

(Prepared by H. C. Sharma, C. L. L. Gowda, G. V. Ranga Rao, M. K.
Dhillon, and M. El Bouhssini)

**Pest Management in Lentil**

Lentil is an important pulse crop grown in southern Europe, North
Africa, Asia (Turkey, Syria, Lebanon, Jordan, and India), the former
USSR, and North America. Ethiopia contributes more than 66% of the area
and about 60% of the production of lentil in sub-Saharan Africa. Poor
crop management and abiotic and biotic stresses reduce grain yields.
Among the biotic constraints, insect pests play a major role in yield
reduction. About three-dozen insect pests have been reported to infest
lentil under field and storage conditions, out of which 21 species have
been reported from India. However, only a few of these are economically
important, and require control measures. The insect pests feeding on
lentil under field conditions include aphids, *Sitona* weevils, *Lygus,*
stinkbugs, cutworms, thrips, bud weevils, and pod borer. During storage,
several species of seed beetles such as *Bruchus* spp. and
*Callosobruchus* spp. cause severe damage. The pest status of each
species largely varies across locations. For example, *Aphis
craccivora*, *Etiella zinckenella*, and *Callosobruchus chinensis* have
been reported as the major pests of lentil in India, while *Sitona
crinitus*, *Bruchus lentis*, and *E. zinckenella* have been identified
as the most important harmful insects of lentil in Turkey. Aphids
(*Acyrthosiphon pisum* and *A. craccivora*), bruchid (*Bruchus lentis*),
thrips (*Thrips tabaci* and *T. angusticeps*), and leaf weevils (*S.
lineatus*) are the key pests of lentil in Castilla La Mancha (central
Spain). The *Lygus* bugs are major pests of lentils in the Pacific
Northwest USA. Control measures for each species or group of insects
(usually related species or genera) are described below.

**Cultural practices.** The stem weevils, *Sitona* species emerge from
the soil after aestivation, and hence, crop rotation can reduce the
likelihood of successful re-colonization and subsequent infestation.
However, it may be noted that *Sitona* weevils are strong fliers and can
migrate considerable distances. Weedy fields are more prone to *Sitona*
damage. Thus, early sowing combined with the control of *Sitona* and
weeds, and P application gives higher net returns with virtually no risk
of economic loss to the farmers. Cultural control programs for *Lygus*
bugs are only partially effective because the target insect is supported
by a variety of hosts. The continuity of plant hosts support *Lygus*
bugs throughout their life cycle. Disturbing habitat by disking near
fencerows and mowing roadsides can potentially lower *Lygus* bug
numbers, but may also injure over-wintering populations of beneficial
insects. Flooding fields has been recommended as a control measure for
the cutworm, *Agrotis ipsilon*. Deep plowing of fields between the crops
turns up larvae and pupae to the soil surface making them susceptible to
predators and sunshine. Weed hosts on outlying areas are often preferred
sites for oviposition and serve as food for the cutworms during the
off-season. Removing volunteer soybean plants from the lentil crop is
critical, since volunteer soybean plants serve as a source for thrip
outbreaks in lentil.

**Host plant resistance.** Host plant resistance to aphids has a
potential for use in pest management in lentil. The cultivar Yerli
Kirmizi shows reduced nodule feeding by *Sitona*, and increased seed
yield over Sazak 91. Isoflavone reductase can be used to select for less
susceptibility of lentils to *Sitona*. However, no sources of resistance
to *Sitona* have yet been found in the lentil germplasm. The CryIII
toxin expression in nodules resulted in significant reduction in nodule
damage by *S. lineatus* on *Pisum sativum*, and *S. hispidulus* on
*Medicago sativa*, and could be used as a component to produce
*Sitona*-resistant lentils. Early maturing and small-seeded genotypes of
lentil are more susceptible to thrips in Bangladesh, but thrips in
general are not a major pest of lentil. Host plant resistance can also
be used for the control of *E. zinckenella* in lentil, since some
variation in genotypic susceptibility to pod borer has been reported.
Short-duration genotypes suffer more damage by *E. zinckenella*. The
genotypes P 927, P 202, and LH 90-39 are resistant to lima pod borer
damage, while LL 147 has shown a tolerant reaction.

**Biological control**. Aphids are attacked by a number of natural
enemies, especially coccinellids, which prevent rapid increase and
reduce infestation levels sufficiently. *Lygus* bugs have several
natural control agents including a fairy wasp, in the family Mymaridae,
that parasitizes the eggs of the bug.  The parasitic wasp, *Peristenus
pallipes* attacks *Lygus* nymphs, but its effectiveness is not well
documented. A European species, *P. digoneutis* has been introduced into
alfalfa fields in eastern North America where it parasitizes about 40%
of the tarnished plant bugs.  One of the few parasitoids of *Lygus*
adults is a tachinid fly, *Alophorella* sp., and nabid plant bugs,
big-eyed bugs, and spiders, which can also be used for biological
control of this pest in lentil.

**Bio-pesticides and natural plant products**. The water extract of
*Melia azedarach* kernels at 50 g L^-1^ significantly reduced *S.
crinitus* adult damage on lentil leaves for one week, and can be used
for minimizing the losses due to lepidopteran pests.

**Chemical control**. Insect damage in lentil, in general, is not high
enough to warrant application of chemical insecticides. However,
need-based application of insecticides may be undertaken under heavy
insect infestation. In the event of severe aphid infestations before or
at flowering, need-based application of dimethoate is quite effective.
Dimethoate can be applied for aphids and Lygus bugs at the same time.
Seed treatment with imidacloprid reduces bean leaf roll virus (BLRV),
faba bean necrotic yellow virus (FBNYV), and soybean dwarf virus (SbDV)
in faba bean and lentil. Seed treatment increases yields of susceptible
lentil varieties, but not that of the resistant ones. The products with
greatest potential for pest control on lentil include bifenthrin,
cyfluthrin, imidacloprid, and λ-cyhalothrin. In addition, efforts are
being made to develop thiomethoxam as a seed treatment for aphid
control.

Carbofuran increases nodule mass by significantly reducing *Sitona*
nodule damage. Chlorpyriphos (720 g a.i. ha^-1^, malathion 1,300 g a.i.
ha^-1^, or oxy-demeton methyl 265 g a.i. ha^-1^ can also be applied in
case of severe insect infestation. Carbofuran and aldicarb reduce
*Sitona* damage in nodules significantly, and yield increases due to
carbofuran are generally higher in early than in the late-sown crop.
Seed treatment with furathiocarb effectively controls the damage by
*Sitona*, increase grain and straw yield, and is less disruptive to the
environment than foliar insecticide sprays. Treatment for *Lygus* bugs
invariably takes place when treatment for pea aphid is made in the USA.
This usually occurs at 50 percent bloom, and the rate of dimethoate used
for aphid control is adequate for *Lygus* control.

Cutworms can be controlled by broadcasting a poison bait (10 kg ha^-1^)
prepared with wheat bran, cotton or groundnut cake, and moistened with
water and trichlofon, carbaryl, or parathion in the evening.
Insecticides such as phosalone and carbaryl effectively control thrips
on lentil. Application of an insecticide for sucking insects (e.g.,
deltamethrin, malathion, dimethoate, or endosulfan) also provides good
control of thrips. Control measures for pod borers,
*Helicoverpa/Heliothis*, *Etiella zincknella*, and *Cydia nigricans* are
rarely needed. These insects directly affect the seeds and consequently
yield, and insecticide application may occasionally be necessary.
Application of endosulfan, indoxacarb, methidathion, and deltamethrin.
at flowering/early pod setting is quite effective. Technologies for
managing *H. armigera* with biopesticides such as *Bacillus
thuringiensis*, HaNPV, and neem seed kernel extract are also available,
and can be used as a viable alternative to synthetic insecticides.

######## 

**Control of insects in storage.** The most serious and frequently
encountered insect pests of stored lentils are *Callosobruchus
chinensis*, *C. maculatus*, *Bruchus ervi*, and *B. lentis.* Store the
grain in a cool and dry place. The grain should be dried to \<12%
moisture content, and stored in airtight containers or sealed polythene
bags, which can be kept under sunlight for 2 to 3 days to kill the
insects that may have infected the seed before storage. Spread a layer
of neem leaves, wood ash, or dust on the top of the storage containers
to minimize bruchid infestation. The grain may be split before storage
(as *dhal*) to reduce the chances of infestation during storage.
Fumigate infested seeds with phosphine before storage, and do not plant
infested seeds. Application of two sprays (one before 50% flowering, and
second 15 days later) of either of endosulfan, methyl parathion, or any
other contact insecticide provides adequate control from bruchid
infestation in the field. Stores should be cleaned of all residues of
earlier stored products and de-infested with malathion. Stored seeds can
be enclosed in a polythene bag and fumigated with phosphine, which
controls all insect stages, leaves no residue, and does not affect taste
or germination. Mixing seeds with olive oil and salt (5 ml and 20 g
kg^-1^ seed) or neem seed oil (3 ml kg^-1^ seed) provides good control
for a period of 3 to 4 months. A dose of 1 kGy of gamma radiation
completely kills *C. chinensis* within a week, and is a suitable
alternative control measure for *C. chinensis*. The dose is also
sufficient for the management of other pests including *Tribolium
castaneum* and *Rhizopertha dominica*. There are several other
non-chemical approaches for controlling species of *Callosobruchus*.
Neem is probably the most accepted alternative to fumigation and
chemical control.

###### Selected References

Bragg, D., and Burns, J. W. 2001. Pea aphid and pea seed weevil control
in dry field peas: Effects on yield and yield components from
insecticide application timings
http://variety.wsu.edu/Updates/2001_Pea_Report.pdf .

El Damir, M., El Bouhssini, M., and Al Salti, M. N. 2000. Effect of
*Melia azedarach* (L.) (Meliaceae) fruit extracts on *Sitona crinitus*
(H.) (Coleoptera: Curculionidae) adults feeding. Arab J. Plant Prot. 18:
64-67.

Islam, M. S., Solh, M.B., and Saxena, M. C. 1985. Effect of
fertilization, inoculation, and carbofuran on nodulation, yield, and
protein content of lentil \[*Lens culinaris*\]. Lens Newslett. 12:32-36.

Lal, S. S. 1992. Insect pest of lentil and their management: review.
Agric. Rev. 13: 225-232.

Weigand, S., Lateef, S. S., El Din, N. E. S., Mahmoud, S. F., Ahmed, K.,
and Ali, K. 1994. Integrated control of insect pests of cool season food
legumes. Pages 679-694 in: Expanding the Production and Use of Cool
Season Food Legumes, Proceedings of the Second International Food Legume
Research Conference on Pea, Lentil, Faba bean, Chickpea, and Grasspea,
F. J. Muehlbauer and W. J. Kaiser, eds. Kluwer Academic Publishers
Group, Dordrecht, Netherlands.

(Prepared by H.C. Sharma, M.K. Dhillon, and M. El Bouhssini)