Insect Pests of Mubgbean and Their Control N.S. Talekar...

Insect Pests of Mubgbean and Their Control

N.S. Talekar Entomologist

Asian Vegetable Research and Development CenterShanhua, Tainan, Taiwan

1. Introduction

Mungbean, Vigna radiata (L.) Wilczek, is a major legume crop grownwidely in south- and southeast Asia, mostly on small-scale family-owned farms.This low input, short duration crop is prized for its seeds, which are high in protein,easily digested, and consumed as food. It is an important source of dietary protein,especially in Indian Subcontinent where consumption of animal protein is very low.Because of its short duration, mungbean is easily adopted to multiple croppingsystem in the drier and wanner climates of lowland tropics and subtropics. Despiteits short duration, large number of insect pests attack mungbean from soon aftergermination to harvest limiting the yield and some pests destroy seeds in storage(Table 1). Since mungbean is grown mainly in the tropical climates, insect pests playimportant role in the economic production of the crop.

The insect pests that attack mungbean can be classified based on theirappearance in the field as it relates to the phenology of mungbean plant. They arethus: 1. stem feeders, 2. foliage feeders, 3. pod feeders, and 4. storage pests. Thisclassification is convenient in judging the economic importance of the pest,especially their influence on seed yield, and in devising control measures. In thischapter, therefore, the pests will be discussed according to this chronological order.

The major stem feeders, especially in seedling stage are the agromyzid flies,so call beanflies, consisting of at least three species. In India the girdle beetle,Oberia brevis (Swedenbord), a major pest of soybean, sometimes attack mungbean.Its damage, however, is minor and localized. Large number of foliage feedersbelonging to orders Lepidoptera and Coleoptera feed on the foliage of mungbeanand several other related legumes. These include armyworms (Spodoptera exigua(Huebner), Spodoptera litura (F.)) hornwonn; (Agris convolvuli (L.)), cottonleafhopper (Amrasca biguttula biguttula Nishida), Bihar hairy caterpillar (Spilosomaobliqua (Walker)), Epilachna spp. and flea beetles, grasshoppers etc. Most of theseinsects are highly polyphagous and feed on wide variety of legumes and non-legumes. Their damage to mungbean is highly localized and most case minor.However, two groups of insects, aphids especially black bean aphids, Aphiscraccivora Koch, and thrips belonging to genus Megalurothrips are especially

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damaging to mungbean and other related legumes They will be discussed here.Major pod feeders are the hemipteran bugs especially the'most widespread southerngreen stink bug, Nezara viridula L., and two species of lepidopterous podborer, theMaruca podborer, Maruca testulalis (Geyer) and Limabean podborer Etiellazinckenella Tretsche. Although other species such as the tomato fruitworm,Helicoverpa armigera (Huebner), Asiatic cornborer, Ostrinia furnacalis (Guenee)and others attack mungbean and other legumes, these legumes are secondary orminor hosts of these pests. Information on these pests can be more easily obtainedfrom study on other crops. The storage pests include species of bruchids belongingto genus Callosobruchus are primary pests of mungbean, especially on stored seeds.Their control is especially important to reduce avoidable losses.

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Table 1. List of insect pests recorded on mungbean

Plant Referencepartsattacked b

Acrocercops phaseospora Meyr. Lep: Gracillarridae L Vyas, 1978

Actinomorpha psittacina (Haan) Ort: Acrididae L Litsinger et al., 1978

Agris convolvuli L Lep: Sphingidae L Nayer et al. 1976

Alcidodes collaris Pasc Col: Curculionidae S Nair, 1975

Alcidodes fabrici F. Col: Curculionidae S Nair, 1975

Amsacta albistriga Walker Lep: Arctiidae L Anonymous, 1970

Amsacta lactinea (Cam) Lep: Arctiidae L Subba Rao et al., 1976

(= Estigmene lactinea) Lep: Arctiidae

Amsacta roorei Butler Lep: Arctiidae L Lai, 1985

Anarsia ephippias Meyr. Lep: Gelechidae L Fletcher, 1914

Anoplocnemis phasiana F. Hem: Coreidae P Nayer et al. 1976

Antricarsia irrolata Lep: Noctuidae L Nayer et al., 1976

Aphis craccivora Koch Hem: Aphididae L, S, F, P Nayer et al. 1976

Apion ampulum Fst. Col: Apionidae Subramaniam, 1959

Aulocophora similis Oliver Col: Chrysomelidae P Litsinger et al., 1978

Autographa nigrisigna Walker Lep: Noctuidae L Nair, 1975

Bemisia tabaci Gennadius Hem: Aleyrodidae L Nene, 1972

Bruchus phaseoli Gyllanhae Col: Bruchidae Sd Anonymous. 1970

Caliothrips indicus (Bagnall) Thy: Thripidae L, F Lal et al. 1981

Callosobruchus analis (F.) Col: Bruchidae Sd Raina, 1970

Callosobruchus chinensis (L.) Col: Bruchidae Sd Gujar & Yadava, 1978

Callosobruchus maculatus (F.) Col: Bruchidae Sd Nair, 1975

Catochrysops cnejus F. Lep: Lycaenidae F, P Litsinger et al., 1978a

Ceococcus coffeaie Green Hem: Coccidae L, S Subba Rao et al. 1976

Ceratina binghanzi Ck1l. Hym: Apidae Nayer et al., 1976

Chauliops fallax Scott Hem: Lygaeidae L, P Rawat and Sahu, 1968

Pest species Order: Familya

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Pest species Order: Family' Plant Referencepartsattackedb

Chrotogonus trachypterus Blanch Ort: Acridiidae L, F, P Lal 1983

Chrysodeixis chalcites (Esper) Lep: Noctuidae L Litsinger et al., 1978a

Clavigralla gibbosa (Spinola) Hem: Coreidae L, P Nayer et al. 1976

Colemania sphenarioides Bol. Ort: Acridiidae L, F, P Nair, 1975

Coptosoma cribaria F. Hem: Pentatomidae P Nayer et al. 1976

Cydia ptychora Meyr. Lep: Pyralidae P Lal et al. 1980

Cvrtozemia coqnata Marshall Col: Curculionidae L Pal, 1972

Diachrysia orichalcea (F.) Lep: Noctuidae L Babu et al., 1978

Empoasca biguttula Shiraki Hem: Jassidae L Litsinger et al., 1978a

Empoasca kerri Pruthi Hem: Jassidae L Pruthi, 1940

Empoasca moti Prughi Hem: Jassidae L ' Chabra et al. 1981

Empoasca terminalis Distant Hem: Jassidae L Chabra et al. 1981

Epilachna philippinensis Dreke Col: Coccinelidae L Litsinger et al., 1978a

Epilachna spp. Col: Coccinelidae L Lal et al., 1980

Etiella zinckenella Tretsche Lep: Pyticidae P Litsinger et al., 1978a

Eublema hemirhoda Walker Lep: Noctuidae L Nayer et al., 1976

Euchrysops cnejus F. Lep: Lycaenidae P Nayer et al. 1976

Eucosma melanaula Meyr. Lep: Eucosmidae L Nayer et al. 1976

Euproctis scintillans (Walker) Lep: Lymantridae L Subba Rao et al., 1976

Gracillaria soyella V.D. Lep: Gracillariidae L Nair, 1975

Helicoverpa armigera Huebner Lep: Noctuidae L, P Nayer et al., 1976

Homona coffearia Nietner Lep: Torticidae L Litsinger et al., 1978a

Lampides boeticus L. Lep: Lycaenidae P Nayer et al. 1976

Lamprosema indicata F. Lep: Pyralidae L Nair, 1975

Leucopholis irrorata (Chev.) Col: Scarabaeidae R Litsinger et al., 1978a

Locusta migratoria malinensis (Meyer) Ort: Acrididae L Litsinger et al., 1978a

Longitarsus manilensis Weise Col: Chrysomelidae L Litsinger et al., 1978a

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Order: Family Plant Referencepartsattackedb

Luperodes sp. Col: Chrysomelidae Lal et al., 1980Madurasia obscurella Jacoby Col: Chrysomelidae L Menon & Saxena, 1970Maruca testulalis (Geyer) Lep: Pyralidae F, P Nair, 1975Megalurothrips distalis (Karny) Thy: Thripidae L, F Lal et al. 1981Megalurothrips usitatus (Bagnall) Thy: Thripidae L, F Chang, 1992Melanagromyza sojae (Zehntner) Dip: Agromyzidae S Chiang & Talekar, 1980Mylabris pustulata Th. Col: Meloidae F Nayer et al., 1976Myllocerus maculosus Desbr. Col: Curculionidae Srivastava et at., 1977Mythimna separata (Walker) Lep: Noctuidae L Litsinger et at., 1978aNacoleia vulgalis GN. Lep: Pyralidae Nair, 1975Nezara viridula L. Hem: Pentatomidae P Nayer et a1.1976Oberea brevis S. Col: Cyranbycidae S Nair, 1975Ophiomyia centrosematis (de Meijere) Dip: Agromyzidae S Chiang & Talekar,1980Ophiomyia phaseoli (Tryon) Dip: Agromyzidae S. Saxena, 1973Ostrinia furnacalis (Guenee) Lep: Pyralidae S, P Talekar et at., 1991Oxya velox F. (= O. chinensis Thumb.) Ort: Acridiidae L, F, P Singh & Singh 1977Pachytychius mungonis Marshall Col: Curculionidae Fletcher 1917Phaneropteraftrcifera Stal Ort; Tettigoniidae L Litsinger et at., 1978aPhytomyza horticola Gour Dip: Agromyzidae L Singh & Singh, 1978Plusia chalcytis F. Lep: Noctuidae L Nair, 1975Plusia daubei F. Lep: Noctuidae L Nair, 1975Plusia peponis F. Lep: Noctuidae L Nair, 1975Riptortus fiuscus F. Hem: Coreidae P Nayer et at., 1976Riptortus linearis F. Hem: Coreidae P Nayer et al., 1976Riptortus pedestris F. Hem: Coreidae P Nayer et al., 1976Spilosoma oblique Walker Lep: Arctiidae L Nair, 1975Spodoptera exigua (Huebner) Lep: Noctuidae L Singh & Singh, 1978--------------------------------------------------------------------------------------------------------------------------

Pest species Order: Fainilya Plant Referencepartsattacked b

Spodoptera litura (F.) Lep: Noctuidae L Nair, 1975Stonzopteryx subsecivella (Zeller) Lep: Gelechiidae L Litsinger et al., 1978aSylepta sabinusalis Walker Lep: Pyralidae L Litsinger et at., 1978aTaeniothrips longistylus Karny Thy: Thripidae L, F Litsinger et at., 1978aTricentrus bicolor Dist. Hem: Membracidae L, F Nayer et al., 1976Zaphanera publica (Singh) Hem: Alyrodidae L, P Nayer et at., 1976

aOrder: families: Col = Coleoptera, Dip = Diptera, Hem = Hemiptera, Hym:

Hymenoptera,Lep = Lepidoptera, Ort = Orthoptera, Thy = ThysanopterabPlant parts attacked: F = flowers, L = leaves, P = pods, R = roots, S = stem, Sd = seeds

Pest species

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2. Stem feeders

Beanflies

Three species of flies belonging to family Agromyzidae are important pestsof mungbean and several other legumes throughout Asia. The species that attackmungbean are: Ophiomyia phaseoli (Tryon); Melanagromyza sojae (Zehntner)and Ophiomyia centrosematis (de Meijere). Their importance lies in the fact thatthey attack plant soon after germination when it is most vulnerable to insect pestattack. If these pests are not controlled, at times the whole -crop can be destroyedor severely damaged requiring re-sowing of the crop. The nature of damage andseasonality of these agromyzids is similar. Therefore same control measures cancombat all three pests. The biology, nature of damage and control measures aredescribed in details in this section.

Ophiomyia phaseoli

Morphology and Identification

Identification of adults of O. phaseoli in the field difficult because theydo not cause significant damage; they are agile and thus difficult to observe inthe field and, to an inexperienced person, they can be easily confused withadults of other agromyzid species since at least two other species of agromyzids(Ophiomyia centrosematis and Melanagromyza sojae) that attack mosteconomically important legumes simultaneously with O. phaseoli. Spencer(1973) gives details of morphology of adults of several agromyzids, including O.phaseoli. For practical purpose, therefore, it is much easier to identify O.phaseoli and other agromyzids by observing larvae and pupae especially anteriorand posterior spiracles of these immature stages (Figure 1). Besidesmorphological differences, their feeding and oviposition sites within the hostplants give a fairly accurate idea of their identity (Figure 2). These details areexplained by Talekar (1990).

Ophiomyia phaseoli larva is a cortex feeder and pupates in the cortexmostly at root shoot junction. Sometimes pupae can be seen sticking under themembranous epidermis (Talekar, 1990). In all plants the oviposition takes placein unifoliate or early trifoliate leaves. A biotype of this insect in Indonesia layseggs in cotyledons of soybean. Both larvae and pupae can be identified byobserving their spiracles. In both stages anterior spiracles are small, with acircle of six minute bulbs. Posterior spiracles closely adjoin on the conicalprojections usually with about 10 minute bulbs. The puparium is pale yellow,straw colored or light brown.

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A closely related species, Ophiomyia spencerella (Greathead) cannot beeasily distinguished from O. phaseoli in its larval or even adult stages. Theposterior spir acles of larvae or pupae are identical in both species, except thatimmatures in O. spencerella have 9.9 ± 1.2 openings on an average as against 10

Ophiomyia phaseoli Ophiomyia centrosematis Melanagromyza sojae

Larva

Last instar

Anterior

sp i rac l es a

Fa

I;la vat *:

Posterior

s p iracles

iC

a;

Pupa: ;.

Anterior ae*.: .......... *0*. .

spiracles **::

Posteriorspiracles

Figure 1. Morphological characters of immature stages of threeimportant species of agromyzids attacking mungbean inAsian tropics.

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Figure 2. Location of ovipositional and larval feeding sites insoybean plant and morphological characters of three stemfeeding agromyzids.

Ms. = M. sojae, O.p. = O. phaseoli, 0.c. = O. centrosematis(Please note, O. phaseoli does not lay eggs in the cotyledons of

mungbean)

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in O. phaseoli (Cn-eathead 1968). Such minute differences, however, are toodifficult to utilize in quick identification of the insect in the field. The mostprominent character that distinguishes O. spencerella from O. phaseoli is theshiny black pupae of the former as against pale yellow to brown of O. phaseoliand O. centrosematis, another cortex feeding agromyzid. The shiny black colorof the pupae of O. spencerella can be seen even without removing the steinepidermis.

So far O. spencerella is confined to countries in East-Southeast Africaand Nigeria. This insect does not occur outside Africa.

Biology

Eggs

Fertilized females are most active on warm clear days. They are activeflier and seek tender leaves on the host-plant for oviposition. Adults havedistinct preference for younger legume hosts for oviposition and feeding. Theytend to lay eggs during the morning hours on the upper side of the leaves, oftennear the midrib close to the petiole. The eggs are inserted between the epidermisand spongy parenchyma. In all legumes . 0. phaseoli lays eggs in leaves,especially the unifoliate leaves. However, a biotype of this species found inIndonesia oviposits in addition to unifoliate leaves, in cotyledons of soybeanonly. It does not lay eggs in cotyledons of other legumes.

The egg is oval, milky white, opaque or translucent measuring 0.30-0.39mm long and 0.10 to 0.17 mm wide (Talekar, 1990): About 10 to 15 % of leafpunctures contain eggs; remaining punctures are made by adults to feed on plantsap oozing out from the puncture injury. Number of eggs per female varyconsiderably. In common bean (Phaseolus vulgaris), van der Goot (1930) foundthat a female, on an average, laid 94 eggs, with a range of 16 to 183 in herlifetime. Morgan (1938) found a single female laid a maximum of 314 eggs andthat the average for 17 females was 99 eggs/fly. Burikam (1980) found a singlefemale laid an average of 77 eggs in cowpea during her life time. Earlier Raros(1975) reported a mean of 1106 eggs laid by a single multimated female and 416by unimated ones throughout her lifespan. Oviposition period lasts from 3-4days after adult emergence and continues for 10-15 days (Morgan 1938).Incubation period of eggs varies 2 to 4 days depending upon temperature(Agarwal and Pandey, 1961, Singh, 1982; Taylor, 1958).

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Larvae

The egg hatches in its capsule at any time of the day. Ophiomyiaphaseoli undergoes three larval instars. The newly hatched, pale, yellowishwhite first instar larva remains motionless for 1-2 hours before beginningfeedings. The first instar feeds mainly in leaf blade tissue before enteringmindrib eventually entering the stem. The first larval stadium lasts between 1.7to 2 days with a mean of 1.9 days (Raros, 1975). Second instar larva initiallystill feeds inside midrib but soon enters petiole and usually molts into third instarat petiole stem junction. The duration of second instar lasts between 2 to 2.4days (Raros, 1975; Burikam, 1980; Singh 1982). The third instar feedsvoraciously in stem just beneath the epideimis. In young seedling the larvalfeeding can extend up to roots but in most ceases at just below soil surface. Theduration of the third instar varies from 4.5 to 5.5 days with a mean of 4.7 days incommon bean (Raros, 1975), 3 to 4 days (mean 3.33 day) in cowpea (Burikam,1980). Before pupation, the fully grown larva ceases feeding for 6-10 hours,constructs a semicircular window in epidermis for escape of adult emerging fromthe pupae. The prepupal period lasts 1.5 to 2 hours. The freshly formed pupabecomes opaque.

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Pupa

The site of pupation varies depending upon the stage and condition of thehost plant. During the seedling stage, pupation normally takes place beneath theepidermis of the stem, near the soil surface. In the later stages of the host plant,pupation can take place at the junction of the leaf lamina and petiole. In someinstances, pupation is observed in the midrib of the leaflet (Raros, 1975;Burikam, 1980).

The pupa is barrel shaped (Figure 1). The cephalic end is somewhatpointed and the posterior end is slightly rounded. There are 12 visible segments.When newly emerged, it is yellow with a brownish tinge and distinctly darkerends. The segments are well defined and the anterior and posterior spi racles areblack. Shortly before the adult emergence, the color of the whole pupariumbecomes dark brown: Puparium measures between 2.02 to 2.30 mm long and0.81 to 1.05 mm wide (Talekar, 1990).

Pupal period varies according to temperature. Ali-Nasr and Assem(1968) found that below 22°C, the pupation period ranged between 11 to 14 dayswith an average of 13 days. At 28°C, it shortened to 10 to 12 days with anaverage of 11 days, and at 32°C, the pupal period lasted 8 to 9 days with anaverage of 8 days. Morgan (1938) in New South Wales, Australia, observedsimilar influence of temperature under field condition. Pupal period was 2 to 3weeks when eggs are laid in late April (autumn), 4 weeks when the eggs are laidearly June (beginning of winter) and 9 to 10 days in warm summer weather.

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Adult

The fully developed imago emerges from the puparium via a transverse Tshaped slit or a crack made by its ptilinum. Immediately after emerging the softbodied and unpigmented imago remains motionless to allow the wings to unfoldand exoskeleton to harden and darken. The adult fly attains a metallic blackcolor in about an hour. The adult flies of 4 to 5 hours after its emergence frompupa. Agarwal and Pandey (1961) report an average measurement of flies to be2.07 mm long and 4.97 mm wide, including wing expanse. They found femaleto be slightly bigger than male. van der Goot (1930) found that females are 1.88to 2.16 mm long and 0.70 mm wide at the thorax, with a wing expanse of 4.45mm. In males, the body is 1.60 to 1.84 mm long and 0.60 mm wide at thethorax, with a wing expanse of 3.80 mm.

Raros (1975) found that female adults live for 23 to 42 days and males 31to 38 days under undefined laboratory condition. If no food is provided they diein 2=3 days. Burikam (1980) observed much shorter adult longevity; 7.13 ± 2.39and 15.42 ± 3.78 days, respectively, for males and females under laboratorycondition. Singh (1982) maintained three sets of newly emerged flies in thelaboratory. The life span averaged 49 hours for starved flies, 94 hours for fliesprovided with water only and 212 hours for flies provided with glucose solution.

Adults feed on three general food sources; water droplets on the leaves,natural secretions of plants, and host plant sap exuding from feeding andovipositional punctures made on the leaves by the females. Adults starts matingafter an average pre-mating period of 18 hours (Raros, 1975). Copulation takesplace only during day time. Copulation lasts from 4 to 94 minutes with anaverage of 18.5 minutes under laboratory conditions (Raros, 1975). van derGoot (1930) reports the duration of copulation to between 1 to 2 hours usuallyoccurring on the upper surface of the leaves. Males and females mate severaltimes during their life.

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Generations

In Java, Indonesia, van der Goot (1930) reported the maximum number ofgenerations a year to be 14, while in the Philippines, Otanes (1916) and inAustralia, Morgan (1938) found that there are 9 to 11 generations a year. InIndia, Agarwal and Pandey (1961) recorded 8 to 9 generations between July andfollowing April and in Egypt, Abul-Nasr and Assem (1968) found 10-12generation a year.

Five life table studies made by Yasuda (1982) revealed that in commonbean (Phaseolus vulgaris) the differences in the initial number of eggs laid wasnot great among four or five observations, however, the total survival rate fromeggs to adults was season dependent. The survival rate was much higher insummer (14-20%) than in winter (3 to 6%) or Spring (8%). The location ofpuparium within the host plant during different seasons appeared to cause thisvariation. In Summer, pupation takes place in the lower part of the stem orbeneath the soil, in winter it occurs in the upper part. Such seasonal changes inthe site of pupation affects the parasitism by pupal parasites (Pteromalids) andsummer pupae practically escape parasitism.

Economic Impact

In tropical to subtropical Asia, O. phaseoli remains a destructive pest ofmost food legumes; common bean, cowpea, mungbean, blackgram, lima beanand soybean (at least in Indonesia). The nature of extent of O. phaseoli damagein different hosts varies from crop to crop and season to season. In general,however, plants are more heavily damaged in the seedling stage than when theyare old. The consequences of insect attack in the seedling stage, if the plantsurvives, are manifested even in the older plants. In general, the yield lossduring rainy season is much less than in the dry season. In Java, Indonesia, in30 observations at Bogor, van der Goot (1930) found that up to 100% commonbean plants were damaged with high plant mortality and yield loss. In Tanzaniathe yield loss ranges from 30 to 50% (Wallace, 1939, Walker, 1960, Swaine,1968). In New South Wales, Australia, Morgan (1940) found it impossible togrow common bean, indicating thereby 100% plant damage and total yield lossif plants are not adequately protected. In Taiwan this pest causes 35% yield loss(Talekar 1990) in common bean and mungbean.

In Indonesia, a biotype of O. phaseoli attacks soybean. Whereas in restof the world, O. phaseoli lays eggs in leaves in all host plants, in Indonesia, thebiotype lays eggs in soybean cotyledons soon after these plant parts emergeabove ground. Larvae, after initial feeding in cotyledons, enter stems and inmost cases kill soybean plant. The extent of damage and subsequent yield loss

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varies from season to season. In dry season (June to October ) van der Goot(1930) found the plant mortality to be 80% compared to 13% in wet season(November to April). Ophiomyla phaseoli_causes very little if any loss insoybean in rest of the world.

In cowpea, O. phaseoli damage varies from location to location. InIndonesia, although the damage can reach up to 100% of the plant population,the plant mortality is rare (van der Goot, 1930). In the Philippines, however, O.phaseoli infestation is high throughout the year, especially in the dry seasonwhen plant mortality can reach 60%. Those plants that survive the attack remainstunted and produce few or no pods (Otanes,- 1918). In Taiwan O. phaseolidamage reduces cowpea yield by 32% (AVRDC, 1985).

In mungbean (Vigna radiata), van der Goot (1930) reported 100% plantmortality due to O. phaseoli infestation in South Sumatra in dry season. Similarplant.,mortality was observed in Malaysia (Ooi 1973). In Taiwan yield loss ofabout 20% occurs in autumn planting (Talekar 1990). In peas (Pisum sativum)Kooner et al. (1977) report plant mortality of 40% and yield loss of roughly 50%in India.

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Nature of Damage

The most serious damage by adults occur when plants are at the unifoliatestage. The unifoliate leaves show a large number of feeding and ovipositionpunctures on the upper side with corresponding light yellow spots, especially onthe basal portion of the leaf. The first and second trifoliate leaves show someegg holes, but leaves situated above this are practically undamaged. Larvalfeeding soon after hatching leads to numerous larval mines which are better seenon the underside of the leaves just under the epidermis and appear silvery,curved stripes. On the upper side, only few tunnels are visible. Later, both eggholes and larval mines turn dark brown and are clearly visible. In case of severeattack, infested leaves become blotchy and later turn down. These leaves maydry out and even fall down. When infestation comes late (when plants aremature), insect damage is confined to the leaf petioles in which case this plantpart gets swollen and, at times, the leaves may wilt.

The developing larvae in second and third instar mine downward intocortex just underneath the epidermis. Third instar continues to feed downwardsinto the tap root and returns to pupate still inside stem close to the soil surface.The feeding tunnels are clearly visible on the stems (Talekar, 1990). If insectpopulation is high, larval feeding leads to destruction of cortex tissue aroundroot-shoot junction. This initially leads to yellowing of leaves, stunting of plantgrowth and even plant mortality. If the damage is less severe, the root-shootjunction area appears swollen. In some cases plant produces adventitious rootsabove this swollen area on the stem.

In Indonesia, where a biotype of O. phaseoli attack soybeans soon afteremergency, larval tunnels in cotyledons are clearly visible (Talekar, 1990).Later damaged cotyledons turn yellow and drop off. In most cases the plant iskilled with 10-15 days after emergence.

Control Measures

Adequate control of O. phaseoli m tropical to subtropical Asia isnecessary if one is to get satisfactory yields in most economically importantlegumes such as common bean, soybean (especially in Indonesia), mungbean,cowpea and peas. This is especially true when the crop is grown in dry seasonas is the case with most field legumes such as soybean and mungbean which aretraditionally planted after wet-season rice crop. The fact that O. phaseoliinfestation only in seedling stage causes economic yield loss, it is essential tocontrol this pest only during the first 4 to 5 weeks after germination. Since O.phaseoli adults are tiny and agile and its major damage is hidden inside theplant stem, it is important to adopt control measures such as spraying of the

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chemicals immediately after germination or protection of the crop by applyingchemicals in soil simultaneously with sowing of the crop. It is, therefore,important to know the seasonality of the pest to undertake appropriateprophylactic control measures.

At present use of insecticides is the only control measure practiced byfarmers. This is mainly because of the absence of other reliable controlpractices. Considerable research has been done on developing alternate andsafer control measures but so far none of these measures have yielded practicalcontrol measures.

Host-plant resistance

Despite its versatility and potential, this approach has not yet beensuccessfully utilized in O. phaseoli control. This is mainly because the insect isconfined to tropical, and subtropical areas where research on host-plantresistance is virtually non-existent. Most of the significant research on thisaspect in recent years has been carried out at the Asian Vegetable Research andDevelopment Center (AVRDC), Taiwan. AVRDC has identified sources ofresistance to O. phaseoli in common bean (Talekar, 1990), soybean (Talekar andTengkano, 1993), mungbean (Chiang and Talekar, 1980) and cowpea (Talekar,1990). At present breeding for resistance is actively pursued in common beanby International Center for Tropical Agriculture (CIAT), Cali, Colombia at itsprogram in Afiica. However, no active breeding is pursued in other three crops.AVRDC has developed several mungbean breeding lines with moderate level of

resistance to O. phaseoli (AVRDC, 1990). However, the yield and level ofresistance in these lines needs further improvement.

Cultural control

Amongst various cultural practices occasionally attempted so far, only theuse of rice straw and similar other plant straw mulch gives some control of onlya biotype of O. phaseoli that attacks soybean in Indonesia (van der Goot, 1930).This biotype lays eggs in soybean cotyledons and second and third instar larvae

feed in the stems of newly germinated plant which invariably results in plantmortality. The rice straw mulch covers the cotyledons making them inaccessiblefor oviposition. Insect then lays eggs in unifoliate leaves, however, by thenplant has developed adequately and tolerates O. phaseoli damage although theyield could be reduced.

Similarly ridging of the crop reduce plant mortality caused by O. phaseoliboth in common bean and soybean (van der Goot, 1930). Ridging crop 2-3weeks after germination helps to cover the adventitious roots which areproduced by the O. phaseoli damaged plants and damaged area around root

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shoot junction. The soil support prevents lodging and improves survival of thedamaged plants.

Intercropping with 60 crop plants belonging to 14 botanical familiesfailed to protect common bean, soybean and mungbean in tests carried out inTaiwan (AVRDC, 1981a; 1981b).

Biological control

In practically all over tropical to subtropical Asia where O. phaseoli_ is aserious pest, a large number of parasites are also present. However, theseparasites do not seem to play any role in checking the bean fly population. InIndonesia, for examples, where this and other species of agromyzids areespecially serious several species of parasites are known exists (van der Goot,1930) for at least 70 years. However, agromyzids continue to cause unabateddamage. This is particularly true for pest control within 4 weeks after cropgermination when O. phaseoli is particularly devastating. In East Africa twomajor parasite species Opius phaseoli and Eucoilidea sp. are chief biotic factorsaffecting the population of O. phaseoli and a closely related agromyzid, 0.spencerella. However, sufficient agromyzids still survive to cause heavyinfestations in subsequent generations of plants. In Hawaii where O. phaseoliwas accidentally introduced in 1968, Opius phaseoli and Opius importatus wereintroduced in 1969 (Davis, 1971) specifically to control the newly introducedpest. Initial studies showed 100% parasitism of the agromyzid in Kauai and 25to 83% on Maui islands. However, surveys conducted in 1973-74 revealedparasitism to ranges from 8.3 to 23.5% only. Ophiomyia phaseoli still remains aproblem in Hawaii. The hidden nature of larval and pupal stages of O. phaseolireduces the effectiveness of most natural enemies in controlling this pest.

Chemical control

Insecticides are relatively quick acting and give immediate results. Bothpreventive and early curative insecticide treatments show promise in controlling0. phaseoli. However, due to concealed nature of larval feeding and damage,preventive measures have better chances of succeeding than curative. Talekar(1990) gives details of various chemical control measures from the earliest useof inorganics and botanicals to modern insecticides including latest insectgrowth regulars.

Pre-sowing application of commonly used systemic insecticides such ascarbofuran, aldicarb, phorate to soil alongside of seeds helps protect youngseedlings when they are most vulnerable to 0. phaseoli infestation. Thistreatment at the rate of 1 to 2 kg/ha can protect the crop through the vulnerableperiod of 4 weeks after germination only if the soil is acidic. In alkaline soil,

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these chemicals breakdown rapidly and post-sowing application of chemicalsprays 2 weeks after germination is essential to protect the crop through first 4weeks. Coating of systemic chemical like carbofuran on seed before sowing alsocontrol O. phaseoli in cowpea and mungbean (IRRI, 1981).

Spraying of chemical insecticide at weekly interval starting soon aftergemination through only first 4 weeks gives adequate control of 0.-phaseoli ona wide variety of crops. Among the chemicals that give consistent control aremonocrotophos, omethoate and dimethoate. These chemicals are structurallyclosely related and there is strong possibility that an insect strain developingresistance to one will have cross resistance to others (Talekar, 1990). A newlyintroduced trisect growth regular, cyromazine, is effective against wide range ofagromyzids. Neither neem extract nor Bacillus thuringiensis control O.phaseoli.

Integrated Control

At present only chemical control measure is effective at controlling O.phaseoli. Cultural control measure such as rice straw mulching is effective onlyin Indonesia that too on soybean which suffers damage from a specific biotypeof O. phaseoli. Ridging plants provide some control, however, this method istoo -laborious to be practical in commercial production of any legume. Despiteavailability of large number of parasites, their effectiveness is extremely limited.Potential exist for integration of only host-plant resistance and chemical control.Availability of even moderately resistant legume cultivar will cut down the useof chemical insecticides. Sources of resistance in major legumes are alreadyavailable, however, there are no breeding programs to develop agromyzidresistant legumes. Until that happens, we may have to depend on chemicalpesticides to protect legumes from O. phaseoli for commercial production.

Ophiomyia centrosematis


The eggs of Ophiomyia centrosematis are laid in hypocotyl of plant justunderneath the epidermis. The practically transparent eggs, on the average, are0.413 ± 0.023 mm long and 0.163 + 0.025 mm wide.

Larvae have three instars. The first instar is practically transparent and thesecond and third instars are milky white. Larvae become opaque before pupation.The length of the cephalopharyngeal apparatus was 0.22 ± 0.03 mm in the first

instar. 0.44 ± 0.02 mm in the second, and 0.64 ± 0.02 mm in the third (Talekarand Lee, 1988). There was thus linear increase in the length of cephalopharyngeal

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apparatus from first through the third instar.

The anterior spiracles are much longer than posterior ones (Figure 1). Thedistal end of the posterior spiracle is divided into three conical structures with oneopening on each (Talekar, 1990). This feature is retained in pupae.

Initially pupa is light yellow, becoming golden yellow and dark yellow justbefore adult emergence. Pupae on average are 2.30 ± 0.10 mm long and 0.89 ±0.07 mm wide and weigh 0.708 + 0.021 mg (Talekar and Lee, 1988).

Adult is a small, shinning black species. Spencer (1973) describes detailsof other morphological characters.

Biology

Egg

Ophiomyia centrosematis flies hover over soybean plant and alight on thefoliage but descend to stem to lay eggs just underneath the epidermis in the stembelow cotyledon (hypocotyl). The feeding puncture are absent on leaves but aremade in hypocotyl. Oviposition starts on the third day after adult emergence frompupae and peaks on the seventh day (Talekar and Lee, 1988). During theoviposition period one female, on the average, lays 63.25 ± 13.68 (range 45-85)eggs. Most eggs are laid between 1100 to 1700 hours. Egg incubation lasts 44.0± 0.33 days at 25°C.

Larva

Larvae emerging from the eggs fed on cortex just underneath the stemepidermis. There are three instars (Talekar and Lee, 1988). The first instar larvaeare practically transparent and second and third instar are milky white. Theduration of the larval period at 28°C was 10.88 ± 1.89 days (range 9-14).

Pupa

Pupation takes place in stem cortex at the root shoot junction. Initiallypupae are light yellow, becoming golden yellow and dark yellow just before adultemergence. Pupae are 2.30 ± 0.10 mm long and 0.89 ± 0.07 mm wide. The pupalperiod lasts on the average 11.03 ± 0.85 days; ranging 11-13 days (Talekar andLee, 1988). Although O. centrosematis laid up to 13 eggs per plant, there were,on the average, only two pupae per plant. Insect apparently suffers fromconsiderable mortality in egg and larval stage.

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Adult

Adult emergence from pupae takes place during day time. At 18-20°C thepeak emergence occurs at 1100 to 1300 hours whereas at 28-32°C, it takes placebetween 0700 and 0900 hours (Talekar and Lee, 1988). Most matings occurbetween 0500 and 0800 hours. Males live for 15.35 ± 5.63 days (range 6-24) andfemales 12.45 ± 4.06 days (range 6-21) at 28±1°C. Pre-mating period lasts 2.5days, pre-oviposition 3.5 days, and oviposition 12.2 ± 3.8 days at 28 ± 1°C.

Temperature influences the duration of the developmental stage. Withinthe range of 20 to 35°C, higher the temperature shorter were the egg incubationand duration of larval and pupal stage (Talekar and Lee, 1988).

Economic importance

Ophiomyia centrosematis is a minor pest of most legumes in Asia andAfrica. Its damages at times go unnoticed in the presence of more dominant O.phaseoli. In Uttar Pradesh, India, however, O. centrosematis is a destructive pestof peas (Pisum sativum). Singh et al (1981) found that more than 95% of thedamaged plants die when the crop is planted 1-8 October. Plant mortality isreduced in crop planted in November.

Nature of damage

Major damage comes from larval feeding inside stem cortex belowcotyledons. As a result of the feeding the cortex tissue is destroyed. Frass isaccumulated in this part. Upon opening larvae are found feeding in the tunnels.Pupae are found in the same layer but at root-shoot junction. In severe case ofdamage, plant looks wilted and eventually die. Adults make oviposition andfeeding punctures in hypocotyl but these punctures are too small, barely seen bynaked eye.

Control Measures

Because the seasonality and nature of damage of O. centrosematis issimilar to more predominant than O. phaseoli, special control measures for O.centrosematis are rarely undertaken. In fact, except for damage to peas in India(Singh et al., 1981) there are no reports of control measures undertaken to combatO:--centrosematis. No attempts have been made to search for legume varietiesresistant to this pest. Cultural control measures such as ridging and mulching usedto combat O. phaseoli are likely to give control of O. centrosematis. The

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chemical insecticides used to combat O. phaseoli are likely to be effective against0. centrosematis.

Some parasites that attack Ophiomyia centrosematis are primary parasitesof other, more important agromyzids such as Ophiomyia_phaseoli. All parasitesare native. They do not seem to be important in checking the pest population atthe beginning of the season when host-plants are in seedling stage and whenagromyzid infestation causes significant damage. No effort has been made tointroduce any parasite specifically to control O. centrosematis.

Melanagromyza sojae


The eggs which are laid in leaf tissue is whitish partly transparent andmeasures 0.34 ± 0.02 mm in length and 0.15 ± 0.01 mm in width (Lee, 19776,Wang, 1979).

Young larva, found in the pith, is nearly colorless. The peculiar shape, sizeand nature of sclerotization of posterior spiracular bulbs can be used inidentification of larvae of this species (Figure 1). The anterior spiracles are short,knob-like, with eight minute pores. Posterior spiracles are well separated andnormally consist of six raised pores around a central truncated horn. Thesecharacters are retained in pupae.

The pupa is cylindrical, golden yellow, and measures 2.75 mm long and1.00 mm wide (Singh, 1982). Pupa is always located in the pith tunnel, often atthe level of unifoliate leaves of younger plants and usually near the fly escapehole, as a dark depression.

Freshly emerged adults fly has moist crumpled wings and very faintpigmentation on the abdomen and legs. Progressive darkening and hardening ofthe body wall and legs occurs for about 30 minutes during which time the wingsalso become smooth and dry. Soon the fly develops its metallic black color with ametallic shiny abdomen. Antennae, legs, and bristles on head and thorax are allblack. The wings are transparent. Females are larger and have tube-shapedabdomen. In females body length is 1.88 mm, width at thorax 0.70 mm, wingexpanse 4.45 nun. In male body length is 1.60 mm, width at thorax 0.50 mm, andwing expanse 3.90 mm.

Spencer (1973) gives details of other morphological characters.

Eggs

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Biology

Eggs are always laid on the under side of the young leaves; unifoliate if theplant has only two leaves, or in fully opened trifoliate leaves at the basal part ofleaf lamina, near the petiole. Numerous feeding punctures are made on the upperside of the leaves. The egg measures 0.34 ± 0.02 mm in length and 0.15 ± 0.01mm in width (Wang, 1979). The egg is whitish, partly transparent. Usually oneleaflet receives 1 or 2 eggs, however, that number may reach 5 or 6 dependingupon adult population density. Eggs hatch commences in 2 days, peaks in 3 daysand can last up to 7 days after oviposition (Wang, 1979).

Larva

Immediately after emergence, the larva bur rows through the mesophylltissue into the closest vein disappearing downwards in the leaf eventuallytunneling through the petiole ending up in the stem. In the stem, larva burrowstunnel into the pith reaching root shoot junction. It burrows further into thickenedtap root, turns around, and moves upward into the pith, thus widening the originaltunnel. It gnaws through xylem and phloem tissues to the epidermis, making ahole to the outside, close it with debris and pupates in the stem (van der Goot,1930).

The larva is nearly colorless and attracts very little attention when the stemis cut open for observation. Larva undergoes three instars. Singh (1982) reportsduration of three instars at 32 ± 2°C and 70% RH as follows: first instar, 22 hoursand 3 minutes, second instar 42 hours and 52 minutes and third instar 98 hoursand 2 minutes. The total duration of larval stage was 7 days. Natural mortality oflarvae is very high. Despite large number of eggs a maximum of only two larvaewere found in van der Goofs (1930) study in Indonesia. Wang (1979) reports62.1, 24.1 and 20% mortality of larvae in 1st, 2nd and 3rd instars, respectively.

Pupa

The pupa is cylindrical, golden yellow, and measures 2.75 mm long and1.00 mm wide. Duration of the pupal period in the laboratory at 30 ± 2°C and70% RH was 189 hours and 36 minutes (Singh, 1982). At average temperature of27°C, the pupal stage lasts 6 to 9 days in June in northern Taiwan (Lee, 1976). InIndonesia, van der Goot (1930) reports a pupal period of 9 to 10 days.

Majority of pupae emerge into adults during the morning and early hours ofthe day. The total development time from egg to adult is 16 to 26 days with anaverage of 21 days, in lowland Indonesia.

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Adult

The freshly emerged fly has moist crumbled wings and very faintpigmentation on the abdomen and legs. Progressive darkening and hardening ofthe body wall and legs occurs for first 30 minutes, during which time the wingsalso become smooth and dry. Soon the fly develops its metallic black color andseeks soybean and other host plants. Melanagromyza sojae adults are weak fliersand their activity is strongly influenced by the weather. They feed on plant juicesfrom egg and food holes made in the leaves by females, dew drops and similarother moist materials. Copulation occurs 3 to 5 days after adult emergence. Theinsect copulates only in the morning hours from 0700 to 1000 hours. Ovipositionbegins soon after copulation and lasts for 19 days (Wang, 1979). Eggs are laid inthe leaves. Eggs are laid in the leaves. In Taiwan, the insect laid 171±115 eggsper female throughout its life. It laid 1 to 34 eggs per day, and 50% of eggs werelaid within the first 9 days (Wang, 1979).

In laboratory, van der Goot (1930) found life-span of adult to be 15 to 36days with an average of 23 days for females and 10 to 46 days with an average of26 days for males. This life span, according to the same study, was longer than itis under field condition. In Taiwan Wang (1979) reports the life span of 6 to 19days for adult flies. In India, Singh (1982) reports average life span of slightlymore than 4 days at 30 ± 2°C and 70% RH. .

Economic Importance

Melanagromyza sojae is a pest of mainly soybean and to some extentmungbean and blackgram. In soybean, the insect infestation occurs in theunifoliate or early trifoliate leaf stage. By this time plants are well establishedand the insect infestation rarely results in plant mortality. Yield loss varies fromlocation to location and plant growth stage when infestation occurs. Yieldreduction .occurs only when the plant is damaged in seedling stage. The later thedamage, lesser is the yield loss. In Taiwan, yield loss among 163 soybeanvarieties was 31% (AVRDC, 1981). In Shandong Province of China, there arereports ofM. _sojae causing plant mortality in soybean (Anonymous, 1978). Theyield loss there amounts to about 70 to 90 kg seed yield/ha. In India,Bhattacharjee (1980) studied relationship between M. sojae infestation, plantheight and yield loss in soybean. According to his calculations, this insect, if notcontrolled, can cause up to 80% yield loss. This pest probably cause significantyield loss in soybean in Indonesia. However, in most cases, if the crop is notprotected, Ophionryia phaseoli causes severe damage before M sojae infestationbegins. Hence no independent information is available on the extent of plantdamage or yield loss by M. sojae in that country.

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Nature of damage

Melanagromyza sojae overwhelmingly prefers soybean. The adult flies layeggs in unifoliate or early trifoliate leaves and major larval feeding occurs in pithof the stem. On outside there are no symptoms of infestation except some minuteovipositionlfeeding punctures at the base of leaf lamina. When stem is cut open,feeding tunnels with larvae and pupae are visible. In slightly older plants, twoseparate tunnels are often found. The one in the lower half is older and hasdeveloped a dark brown color. It originates in the stem roughly at the junction ofthe unifoliate leaves, and extends downwards up to the soil surface, indicating thatthe infestation occur red earlier, from the eggs laid in the unifoliate leaf Thesecond tunnel starts just under the top the plant and extends downwards up to thefirst tunnel. Presuming that the plant at the unifoliate leaf escaped infestation, thistunnel can extend up to the soil surface. This feeding results from the laterinfestation of trifoliate leaves. If the plant is damaged very early, at times the laterinfesting larvae do not have enough pith tissue to feed on. Under suchcircumstances, the larva gnaws upwards resulting in the hallowing of the topwhich at times leads to weathering of the top.

Control Measures

In most legumes, the seasonality of occurrence of Melanagromyza_sojae issimilar to more predominant agromyzid Ophiomyia phaseoli which infests host-plant at the same time or even earlier than M. sojae. In most cases, the controlmeasures adopted for combating O. phaseoli also control M sojae. Since M.sojae prefers soybean over other host plants, the control measures described hereare for the control of this pest on soybean.

Since M. sojae damage in seedling stage only causes economic yield loss, itis essential to control this pest only during first 4 to 5 weeks after plantgermination. Since adult flies are too small and at times remain hidden in plantcanopy and larval damage is hidden inside plant stem, it is important to adoptcontrol measures such as spraying of chemicals immediately after cropgermination or protect the crop by applying chemicals in soil simultaneously withsowing of seeds. It is important to know the season when the pest is serious toundertake appropriate prophylactic control measures. In general pest is moreserious in dry season than in rainy wet season and precautionary control measuresare more important in dry than in wet season.

At present farmers do not control or use only chemical control to combatM sojae. This is mainly because of the absence of other reliable controlpractices. Some research has been going on in soybean to devise other control

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measures but the progress in Asia, where this pest is more serious, is very slow.


Chiang and Talekar (1980) found four wild Glycine soja accessions highlyresistant to M. sojae. The resistant accessions are viny plants with very thinstems. Breeding resistance into cultivated soybean failed because in order to havehigh yield, it was essential to have plants with thick and strong stems to supportlarge number of pods without lodging. Any increase in stem size beyond the vinystern of the resistant parent increased the susceptibility of the progeny to M. sojae.Since despite 100% plant infestation yield loss is barely 30% and the fact that thispest rarely kills plants, it is possible to develop soybean cultivar tolerant to thispest. Some soybean accessions indeed show tolerance - no yield loss despiteheavy infestation - to M. sojae (AVRDC, 1979).

Biological control

Although several species of parasites are present in areas such as Indonesia,India and Taiwan where M sojaeis endemic on soybean, their parasitism rarelyexceeds 50%. Most of this parasitism comes rather late in the season after insecthas attacked the plant and caused significant damage. During dry season in areasendemic to M. sojae the pest population is so high that enough insects escapeparasitism and continue to cause serious crop damage. Most of the reportedparasites are native and there is no introduction of any natural enemy anywherespecifically to control M. sojae.

Chemical control

Preventive and curative insecticide application both have potential in givingadequate. control of M. sojae. Since M. sojae damage in seedling stage causesyield loss, the earlier the chemical is applied, the better is its effectiveness.Among the more effective chemicals are monocrotophos, dimethoate, omethoate,pyrazophos and cyaromazine. These chemicals must be spray applied once aweek soon after germination to up to 4 or 5 weeks after germination. Systemicinsecticides such as aldicarb, carbofuran and phorate applied in band along withseeds at sowing time gives satisfactory control for up to 2-3 weeks aftergermination. Talekar (1990) gives details of various chemical control measuresfrom the earliest use of inorganics and botanicals to modern synthetic organicchemicals including latest insect growth regulators some of which are specific toagromyzids.

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Integrated Control

At present only chemical control measures are effective in controlling M.sojae. Despite availability of large number of parasite, their utility in giving adequatecontrol of M sojae is limited. Cultural control measures such as mulching which tosome extent is effective in controlling Ophiomyia phaseoli was not control M sojae.Potential exists for integrated of host-plant resistance and chemical control to reducethe pesticide use. However, there is no breeding program in Asia where soybeancultivars resistant to M. sojae are being developed.

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3. Foliage Feeders

Among numerous insects that are reported to feed on foliage of mungbeanin Asia (Table 1), only two groups, one species of aphid, Aphis craccivora Kochand three species of thrips: Megalurothrips usitatus (Bagnall), Megalurothripsdistalis (Karny) and Caliothrips indicus Bagnall are specific to legumes and causesevere damage to mungbean. Most other species are not endemic on mungbean orrelated Vigna species. They are highly polyphagous feeding on wide varietyplant species besides the members of Leguminoceae. Mungbean or related Vignaspecies are not their primary host. Tnformation on these pests can be gleaned frompublications on pests of other economically important crops. In this section wewill discuss the biology and control of A. craccivora and thrips.

Thrips

Three major species of Thysanoptera damage mungbean and other legumesin Asia. Megalurothrips dorsalis (Karny) and Caliothrips indicus Bagnall areprevalent in South Asia and Megalurothrips usitatus (Bagnall) mostly in SoutheastAsia, although there are reports of M usitatus damage, albeit minor, in SouthAsia. Despite the taxonomic differences, the nature of damage in most Vignahost-plant species is similar and most control measures devised for one specieswork for the other species. Despite their importance on legumes in South Asia,there is very little, if at all, published information available on the biology,ecology and natural enemies of M. distalis and C. indicus. In this section,therefore, we will include the information on biology and ecology of M. usitatusbut include information on control measures devised for all three species.

Biology of M. usitatus

Megalurothrips usitatus (Figure 1) is a damaging pest of adzuki bean,vegetable' soybean, grain soybean, and mungbean in Taiwan (Figure 1). Most ofthe biology and ecological research on this pest is done by scientists in Taiwan.The following description is based on publication of Chang (1987, 1988a, b, 1989,1990a, b, 1992).

Megalurothrips usitatus has six distinct developmental stages: egg, larval I,larva II, prepupa, pupa, and adult. Eggs are laid in petals and sepals. This is thereason for occurrence of large number of larvae in coups in adzuki bean flowers.In laboratory, at constant temperatures between 14 and 30°C, the egg, larval, andpupal periods and adult longevity were 2-19, 5-10, 2-7, and 6-30 days,respectively. Female longevity is greater than that of the male at all temperatures.All M. usitatus died after hatching at upper (30°C) and lower

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Figure 1. Adults ofMegalurotrips usitatus.Top, male; bottom, female.

(14°C) temperatures. The mature larvae crawl downward and pupate 1-6 cmbelow the soil surface. Adults emerged in about 5 days.

During vegetative stage, M usitatus is found inside the top, unopened,trifoliate leaves. During reproductive stage when plants start bearing flowers,more thrips are found in flowers on the 7th and 8th node on main stem of soybeanplant. Within the flowers, male and female thrips were randomly distributed inthe initial blooming stage (Chang, 1992). Rainfall adversely affects the survivalof M usitatus. The water drops accumulated in the flowers and vegetative budsdrowns and kill the .thrips. This pest, therefore, is not important during rainy;season but can be devastating during dry season.

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Nature of Damage

Both larvae and adults prefer flowers over the leaves of adzuki beans. Inflowers, both larvae and adults feed on pollen and rasp other flower parts and suckthe plant juice oozing out from the injured plant parts. As a result of this type ofdamage, flowers drop of and no pods are formed (Figure 2). In mungbean plantscan form new flowers but they are also attacked and at times there is total yieldloss. In the absence of flowers, M. "'snafus feed inside

Figure 2. Severe trips damage in mungbeanresults in loss of flowers and plantsdo not produce any pods.

vegetative buds, rasping the unopened leave and sucking plant juice oozing out ofthe plant part. When the leaves open they all appear crinkled giving appearanceof virus damage.

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Control Measures

Biological control

A minute parasitic wasp, Ceranisus menes was first observed on M.usitatus in adzuki bean field in 1988 in Taiwan (Chang, 1990a). It hasoccasionally being observed in laboratory from thrips collected in the field andreared in laboratory. This parasite attacks larvae and emerges before or duringpupation of the thrips. In the Philippines, substantial predation of M usitatus wasobserved in potato. The most important predators were a myrid, Campylommalivida and a spider Argurodessp sp. In Japan Ceranisus femoratus and C. vinctusattack M. usitatus larvae. No information of worthwhile mention on the biologicalcontrol of M distalis and Caliothrips indicus which attack mungbean in SouthAsia where publications after publications describe wasteful research oninsecticide screening.

Cultural control

In study in Taiwan in adzuki bean field, in winter season, efficiency of bluePVC plate traps coated with sticky substance attracted significantly more M.usitatus than yellow or green traps (Chang 1990b). In spring blue trap alsoattracted more thrips than white, yellow or green. However, when the populationincreased, there was no difference in the number of thrips being attracted to blueand white traps. Megalurothrips usitatus was strongly attracted to light withwavelength of 450-480 nm. Potential exist to use these traps for reducing Musitatus damage to legumes grown at least on small scale farms.


In India, in the first of two field tests, Chhabra and Malik (1992) screened70 entries of mungbean germplasm for resistance to M. distalis. Deformity ofinflorescence due to thrips damage was used as a criterion for resistance. On tworesistant genotypes, SML77 and UPM82-4, insect development was prolongedand adult longevity was shortened. In the second test in which 20 genotypes ofmungbean were tested, SML99 and SML100 were the most resistant and SML117was the most susceptible to pest damage (Chhabra and Kooner, 1994).

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Chemical control

Chemical insecticides are widely used to control thrips on mungbean andother legumes. Chang (1991) found that none of the chemicals he used control M.usitatus in Taiwan. In India several insecticides were tested for the control of M.distalis and C. indicus on various crops (Acharya and Koshiya, 1991; Singh andSingh, 1991; Ghorpade and Thakur, 1989; Koshta et al. 1988; Mundhe, 1980;Awate et al. 1978; Awate and Pokharkar, 1977; Gargav and Vaishampayan,1978). Among large number of chemicals tested in these studies, sprayapplication of dimethoate, monocrotophos, chlorpyrifos, quinalphos, and gamma-HCH (Lindane) gave satisfactory control of the pest. Soil incorporation ofphorate, disulfoton, aldicarb and carbofuran at the time of planting also helped toreduce the thrips population.

Aphis craccivora

Aphis craccivora Koch is a polyphagous insect with marked preference tolegumes. Amongst legumes, mungbean alongwith cowpea and groundnut are mostdamaged by this pest.

Biology

Adult aphids are black or dark brown, shiny, abdomen with large, dark,practically solid dorsal plate (Figure 3). Winged parthenogenetic females are 1.5to 2.0 mm long, dark dorsal abdominal plate with cross markings of varying

Figure 3. Adultof Aphis craccivora.

number. Antennaeare about two thirdas long as the body.Nymphs arewingless, dark withfairly rounded body

0 1 2 mm shape. Nymphsappear on the cropsoon after

germination from adults having overwintered or spent dry season on nearbyleguminous plants. In tropics only females, winged or wingless, are found, and

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parthenogenetic reproduction occurs throughout the year. The aphid isovoviviparous, with females retaining eggs inside their bodies and giving birth tosmall nymphs. Males are winged and sexual forms are occasionally found.

The optimal development temperature is 24-28.5°C and relative humidity65%. The optimal daylength for nymphal development is 16 hours light and 8hours of darkness (Abdel Malek et al 1982). Photoperiod does not but plantchemistry, particularly reduction in the rate of translocation of photosynthates,influence formation of winged individuals (Mayeux, 1984). Aphis craccivora iscapable of rapid population multiplication. On groundnut Talati and Butani(1980) observed that on groundnut offsprings from a single gravid adult aphidranged 17 to 43 in 15 days. In the same laboratory study authors noted fournymphal- instars in A. craccivora when reared on cowpea. The total nymphalperiods averaged 5.6, 5.1, 5.15, and 4.86 days in May-June, August-September,October-November, and March-April, respectively, in India. The duration of thetotal life-cycle during the corresponding periods were 11.07, 11.15, 10.79, and10.42 days, respectively (Patel and Srivastava, 1989).

Aphis craccivora infestation of mungbean, cowpea or groundnut is seriousonly during cool-dry season. They do not survive periods of heavy rains. Duringdrought, the pest survives on leguminous weeds. It can also withdraw into cracksin soil. As soon as the rains come and aphids are pushed out by closing cracks insoil, they colonize aerial plant parts. Aphis cracivora often migrates over widedistances into dry zones where, given transient periods of sufficient rainfall, it cancause damage, especially be transmitting viruses. During drought, irrigated plantsare more heavily damaged than those that did not receive irrigation.

Young colonies of nymphs concentrate on young shoots and are regularlyvisited by ants and there is mutualism between ants and aphids.

Nature of Damage

Young aphids cluster over tender shoots and occasionally young pods ofmungbean and suck plant sap-from these plant parts. Heavy infestation weakensthe plant and entire plant can be destroyed. Severe attack at the time of floweringand seed formation affects yield and produce infestation weakens the plant andentire plant can be destroyed. Severe attack at the time of flowering and seedformation affects yield and produce wilt symptoms. In addition, abnormalitiesdue to virus diseases - rosetting, stunting, mosaic, mottle etc. - can be observed.

The greatest damage results from virus diseases which are transmitted by A.craccivora, especially in groundnut. Among the virus vectored by this aphid invarious crops are: alfalfa mosaic, bean common mosaic, bean yellow mosaic,cowpea aphid-borne mosaic, cowpea banding mosaic, cowpea mild mottle, beanleaf roll and chickpea stunt virus. In mungbean, it transmits at least three viruses;green mosaic, leaf curl browning and little leaf (Benigno and Dolores 1978).

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Control Measures

Biological control

Most natural enemies of aphids are polyphagous attacking wide range ofaphid species in a particular habitat. Therefore, important natural enemiesattacking particular aphid species on crops tend to be different according to cropspecies and climate. This is especially true of aphid species such as A.craccivora, attacking a range of crop over large geographical areas. In addition,many natural enemies, especially parasitoids, are members of species complexes,morphologically very similar but with different host preferences and geographicaldistribution. Some of the important parasitoids of A. craccivora are: Thioxysindicus, Lysiphlebus fabarum and L. tesaceipes. Singh and Sinha (1983) found9.4% parasitism by T. indicus shortly after appearance of A. craccivora onpigeonpea, in India. The peak rate of 64.6% was observed in later stages ofinfestation which was sufficient to suppress aphid populations on pigeonpea.

Important predators include coccinellid beetles, e.g. Cheilomenessexmaculata and Coccinella septempunctata, neuropteran larvae, e.g. Micromustimidus and predatory diptera, e.g. Aphidoletes aphidimyza and a syrphidIschiodon scutellaris. Use of chemical insecticides however, suppresses activityof all these beneficial arthropods. To conserve these natural enemies insecticidesthat are least toxic to predators and parasites that too only cases of absolutenecessity.

Cultural control

Densely planted groundnut fields sown as soon as possible discouragescolonization by aphids. Early sowing allow plants to start flowering before aphidsappear, while dense sowing provide a barrier to aphids penetrating in from fieldedges. Sanitory measures are needed during the season and between seasons toprevent spread of viruses vectored by A. craccivora. Virus infected plants shouldbe removed and any volunteer plants or weeds that could harbor viruses should bedestroyed promptly. Insecticide applications were more effective in minimizingthe incidence of A. craccivora when chickpeas were intercropped with barley orlinseed (Prasad et al.. 1988). However, mungbean . cowpea or groundnut are notsuitable crops for intercropping due to the risk of spread of the insect betweenthese favorable host-plants.

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Entomological research at IITA emphasizes breeding of cowpea cultivarsresistant to aphids. Please read IITA's annual reports for the latest infolmation onaphid resistant cowpea. Potential exists for finding mungbean cultivars resistantto A. craccivora, however, due to uncertain incidence of A. craccivora in Taiwan,most of AVRDC's mungbean geimplasm has not being evaluated for resistance toaphids. Planting of aphid tolerant groundnut cultivars allows much retardedincreased in small aphid colonies, at the same time predators arriving from wildplants can buildup populations and reduce aphid infestation effectively (Heinze1977).:, ...

Chemical control

Most major groups of insecticides, especially organophosphorus andcarbamates, have been tested and some of them found effective against widevariety of aphid on economically important crops. Pirimicarb a selective aphicideis widely used to control various species of aphids. Other chemicals includeacephate, dimethoate, endosulfan, menazon, and thiometon which have beenrecommended for aphid control. Other sprays found promising on crops includeneem (Dimetry and El Hawary, 1995) and petroleum oil (El Sisi and El Hariry1991). Cost of some of these sprays could, however, be prohibitive to subsistencefarmers growing mungbean.

Integrated pest management

Potential exist for the integrated control of A. craccivora. Combinations ofselective insecticides, predators and parasites, cultural methods and resistantcultivars has potential of controlling the pest on a sustainable basis. In groundnut,monitoring pest populations to time insecticide spray application is combined withthe use of cultural methods and resistant cultivars (Mayeux 1984). In Bangladeshthe IPM involving using malathion along with natural predation of Menochilussexmaculatus was successful in controlling A. craccivora on beans (Ahmad andSardar, 1994).

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4. Pod Feeders

This group of pests include those that feed' externally on the pod at timesdevouring the pod, pierce their proboscis through the pod pericarp and such thecontents of developing seeds, and bore inside the pod and feed on developingseeds while remaining concealed inside the pod. All these pests cause direct yieldloss.

Some species of polyphagous hemipteran bugs belonging to familiesPentatomidae and Coreidae pierce green. mungbean pods and suck juice fromdeveloping seeds. The species belonging to genus Nezara of the former familyand genus Riptortus of the latter are especially destructive. Although these pestscan feed on stems, inflorescence and even leaves, they prefer green pods and theirdamage to pods is economically more important.

There are at least four serious lepidopterous pests that bore into pods andfeed on developing seeds. They include Maruca testulalis (Geyer) Ostriniafurnacalis (Guenee), Helicoverpa armigera (Huebner), and Etiella zinckenella(Treitschke). Some of these species are more serious at certain locations and insome cases their infestation levels vary from season to season. Not surprisingly, attimes, all species can be present simultaneously, since the ecological conditionsfor their growth and development are similar. The nature of their damage to podsis not too different. The larvae of these insects bore inside the pods and feed ondeveloping seeds, resulting in direct yield loss.

Nezara viridula

Among several species of hemipteran bugs belonging to families Coreidaeand Pentatomidae that damage mungbean pods, Nezara viridula (Figure 1), thecommonly known as the southern green stink bug, is the most destructive,geographically widespread, and has the widest host range. One other speciesRiptortus linearis is confined to South and Southeast Asia including Taiwan.Both species which attack mungbean are more serious on economically dominantsoybean crop which appears to be their primary host. Most of the infolmation onbiology, ecology and host-plant interaction is, therefore, generated with soybeanas a host-plant and very little on other crops. The nature damage and controlmeasures for both species are similar both on soybean and other legumes,including mungbean. Because of its greater economic importance, detailedinfolination only N. viridula will be included with brief account of R. linearis.

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Biology

Nezara viridula adult starts laying eggs 18-25 days after becoming a fulladult. The eggs are laid regularly in masses with oviposition lasting from 27-32days with an average of 29.2 days. The females oviposit 4-6 egg masses (Singh1973), with a normal egg mass consisting of 42-113 eggs.

I I l , 1

0 5 mm

Figure 1. Adults of Nezara viridula.

Egg incubation is 4-6 days, depending upon temperature. Singh (1973)found that the mean length of incubation for 103 masses containing 7019 eggs was4.9 days, with cool temperatures during the months of December throughFebruary prolonging incubation by about a day.

The first instar nymphs are yellowish orange and slightly bigger than theeggs, shortly after emergence, they darken, remaining compactly clustered andmotionless on the empty egg mass or adjacent to it. The first instar lasts 4-5 daysduring which the pest does not feed.

The second instar nymph has a bright red on the head, large orangethorax, black abdomen and black eyes. It moves slightly away from the clusterandfeeds on green pods, although it-can feed on any part of the host plant. Thisstage lasts 3-4 days (Singh 1973).

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The color, spots, and other markings remain essentially the same in thethird instar, which lasts for 3-4 days (Singh 1973). The insects disperseconsiderably for feeding and settle to form small feeding aggregations.

The color pattern varies markedly during the fourth instar, which lasts 3-4days. Individuals can be separated into light and dark, with the latter accountingfor 10-15% of the population (Singh 1973). The nymphs begin to disperse.

The general outline of the body of the fifth instar nymph is the same as thefourth instar, but the developing wing pods become conspicuous and cover thebasal portion of the abdomen. The aggregating behavior completely disappears bythe fifth instar when nymphs become widely dispersed for feeding as well asresting. The fifth instar stage 5-7 days, the total length of nymphal stages varyingfrom 18-28 days.

The adult is a large green bug. It has the shield-shaped form characteristicof a pentatomid, and it has the appearance typical of a stink bug. The female islarger than the male, and the males can be differentiated from females by a notchand two brown spots on the ventral surface. of the terminal end of the abdomen.

Several different color types, varieties or forms of this insect have beenreported from tropical to subtropical Asia. At times they may have been confusedfor new species.

Riptortus linearis is widespread in tropical and subtropical Asia. Besideslegumes, this species also feed on members of Solanaceae and Convolvulaceae.Eggs are laid in clusters of 3-5 mainly on the underside of the leaves and on pods(Kalshoven 1981). When newly laid, the eggs are gray, turning dark brown beforehatching within 6-7 days. The nymphs undergo five instars in about 3 weeks inIndonesia and adult lives from 4 to 47 days. Both nymphs and adults feed ondeveloping seeds within pods.

Nature of Damage

Although all stink bug species can feed on leaves and tender sterns, ingeneral they prefer to feed on developing seeds within green pods. In soybean,while feeding stink bug injects histolytic agents into the seed, liquefying thecontent with the cells and causing cell wall to rupture. The insect then sucks thecontents.

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When stink bugs feed on developing pods, the seeds do not develop and, attimes, at least in soybean, pod drop occurs. The plant compensates for lost podsby setting new ones but these pods remain small. When harvested, the damagedseeds are shriveled and deteriorate rapidly in storage. In damaged seeds,germination is adversely affected. Stink bugs are also reported to transmit diseasecausing organisms in at least soybean seeds, which adversely affects germinationof the seeds. It is not known whether this phenomenon occurs in mungbean andsimilar other legumes.

Control. Measures

Several factors including weather and natural enemies regulate populationof stink bugs. Kiritani and Hokyo (1962) showed that more than 94% of N.

viridula die before becoming adult. Mortality from egg to third instar nymph was70-95%. Causal factors were stage-specific: parasites killed eggs, weather factorskilled the first-instar nymphs, and predators killed second instar nymphs.

Biological control

In Japan two scelionid parasites, Asolcus mitsukurii and Telenonusmakagawai, heavily infested eggs and were decisive in keeping the population ofN. viridula from causing major losses (Kiritani and Hokyo, 1962; Hokyo andKiritani, 1963). Kamal (1937) found that a proctotrapid egg parasite,Microphanurus megacephalus (Ashmead), was the only biotic agent preventingstink bug from becoming a pest in Egypt. In Hawaii, Trissolcus basalis

(Wollaston) and Trichopoda pennipes var. pilipes F. prevent outbreaks of N

viridula (Davis, 1961, 1967; Davis and Krauss, 1963), and in Java, Indonesia, twospecies, Ooencyrtus malayensis Ferr. and Telenomus sp. parasitize eggs of N

viridula. In Malaysia, a reduvid bug, Sycanus collaris (F.), keeps N viridulaunder control, and in the Philippines, Corpuz (1969) found one hymenopterousparasite, Ooencyrtus sp, preying on the eggs of N. viridula. At AVRDC inTaiwan, we found another hymenopterous parasite, Trissolcus sp., preying on theeggs of R. linearis.

Since insecticide use for the control of stink bugs and most other pestsinfesting mungbean and related field legumes such as cowpea and soybean is stillminimal compared to vegetables, use of these parasites may be the best means tocontrol stink bugs. In fact, N viridula was successfully controlled in Hawaii byintroduction of parasites from other countries (Davis, 1961, 1967, Davis andKrauss, 1963). Trissolcus basalis was imported from Australia and a tachinid,Trichopoda pennipes var. pilipes from the West Indies - the two species exertingenough pressure on N. viridula to eliminate serious outbreaks since 1963 (Singh,X973).

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Chemical control

Several chemicals have been screened and some recommended for thecontrol of stink bugs. When synthetic organic insecticides were first introduced,organochlorines such as DDT gave satisfactory control. However, in later tests,DDT proved ineffective and all stages of N. viridula were reported to havedeveloped resistance to DDT in Hawaii (Miyazaki and Sh6iman, 1966). Severalorganophosphorus insecticides and later carbamates such as carbaryl andmexacarbate also gave satisfactory control of stink bugs (Mitchell, 1965;Miyazaki and Sherman, 1966; Swaine, 1969). In a test in Taiwan, one syntheticpyrethroid, fenvalerate, gave good control of stink bugs and considerably reducedpod damage (AVRDC, 1982a).

Maruca testulalis

The Maruca podborer, Maruca testulalis (Geyer) (Figure 2) is a tropicalinsect attacking several species of food legumes in Asia, Africa, Central America,and South America. Within its wide host range, covering practically alleconomically important food legumes, Maruca feeds on practically all above-ground plant parts - young shoots, flower buds, stems, flowers, pods, anddeveloping seeds. There are no reports so far of this insect feeding on leavesexcept when oviposition take place on leaves when newly hatched larvae maybriefly feed on leaves before moving to feed inside flower or pod. Its damage tomajor plant parts severely affects the productivity of food legumes wherever theinsect has achieved pest status.

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Figure 2. Maraca testulalis adult.

Biology

The biology and the nature of damage by Maruca podborer is similar inmost Vigna species, especially between cowpea and mungbean, two of Maruca'seconomically important hosts. Since relatively more work is done on this pestwith cowpea, most of the infoliiiation on the biology of this pest conies fromstudies on this legumes than on mungbean.

Eggs are mostly deposited on. flower buds and flowers of the legume host-plant. Jackai (1980) found 18.9 eggs on flower buds, 31.0 on flowers, 28.2 onabscission scars, 18.9 on peduncles, 2.6 on terminal shoots and 0.4 on pods ofcowpea plants. Sporadic oviposition on leaves, leaf axils, terminal shoots, andpods has been . observed. Because of the erratic oviposition behavior duringrearing Maruca podborer moth in laboratory, the reports of fecundity of this pestvary considerably: 8-140 eggs/female (Taylor 1967), 6-189 (Akinfewa 1975), 6 to194 (Okeyo-Owuor and Ochieng 1981). Over 72% of the eggs in the latter studywere fertile. In studies in India on three legume hosts; pigeonpea, cowpea andhyacinth bean, the fecundity varied between 35 to 38 eggs per female and 83% to89% of the eggs were viable (Ramasubramanian and Sundara Babu 1989).

The roundish to slightly elongated oval eggs measure 0.65 x 0.45 mm.They are slightly yellow, translucent and have reticulate sculpturing on the thinand delicate chorion. The eggs are deposited in batches of 2 to 16 between thewhorls of flower buds. Ramasubrainanian and Sundara Babu (1989) found insectto prefer hyacinth bean over cowpea or pigeonpea for oviposition.

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The eggs hatch in about 2 to 3 days and the insect undergoes 5 larvalinstars during the total larvae period of 9 to 14 days depending upon host-plantspecies and temperature. The caterpillar is dull white with dark spots on eachbody segment, forming dorsal longitudinal rows (Figure 3). The mature

Figure3. Maruca testulalis larva.

caterpillar is 16 mm long.Larva is a voracious feederfeeding mainly on flowersand green developingseeds inside the maturingpods.

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The 2-week larvalperiod is followed by 2 days of prepupal period during which feeding ceasescompletely and larva becomes pupa. The pupa is greenish or pale yellow initiallydarkens to a grayish-brown. Pupation takes place in a silken cocoon in the pod ormore often in soil. Taylor (1978) describes the pupa, when pupation takes placein soil, as a double-walled pupal case consisting of an outer wall of silk, soilparticles and other debris and an inner wall of loose strand of white silk woven infishing net fashion and open at the anterior end. Pupation lasts 6 to 8 days and 68to 76% pupae emerge into adults. (Ramsubramanian and Sundara Babu 1989).

The adult that emerges from pupa has light brown forewings with 3 distinctwhite spots. The hindwings are pearly white with distal brown markings. Thewingspan measures 16 to 27 mm. In study in India with pigeonpea, cowpea andhyacinth beans, the adult longevity was 5.9 to 6.1 days for male and 8.5 to 10.0days for female (Ramsubramanian and Sundara Babu 1989). Mating period lasted1.6 to 2.7 days and total oviposition period 3.6 to 3.9 days. Sex ratio was 1:0.50to 1:0.84 in favor of females. The total life-cycle varies from 18 to 35 daysdepending upon temperature.

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Nature of damage

Flower buds, flowers, and pods are major plant parts attacked by Marucapod-borer. Young larvae usually attack buds and flowers and older ones bore inmaturing pods. The larva soon after emergence from egg starts feeding initiallyeven on the leaves if oviposition has occurred on the foliage. In flower budswhich usually harbors 1st or 2nd instar larvae, larva makes holes through the budand feeds hidden inside. In open flowers all flower parts are damaged and excretais accumulated in the flower. In most cases these flowers drop and do not bearany pod. The larval feeding in pods starts with small hole usually in proximalparts of the pod if the oviposition has occurred in the inflorescence. It feeds onthe developing seeds moving from seed to seed concealed inside until it is readyfor pupation.

In mungbean, invariably adjacent 2-3 pods are stuck together around areawhere insect has made entry hole in a pod. Sometimes, usually when the eggs arelaid on leaves and if the leaf is in the proximity of a pod, larva sticks together partof the pod touching the leaf and makes hole in the pods be it distal or central partof the pod (Figure 4). We rarely find a larva boring inside a pod which is nottouching to other pod or leaves. This indicates that a mungbean cultivar with podsnot touching each other and radiates from well above the foliage so that the podswill not touch stem or foliage could be "resistant" to Maruca podborer.

Figure 4. Typical damage in mungbean by M. testulalis. Part of the podtouching a leaf is stuck together with the leaf and insectlarva bores inside the pod. In the picture above the pod isturned slightly to expose the larvae.

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In Sesbania indica in Taiwan at least, insect lays eggs on leaves and larvaefeed on foliage throughout the season. Newly emerged larva sticks several leafletsof compound leaves together and feeds concealed inside the leaf whorl. As aresult of such feeding, the leaves are destroyed. Sometimes most of the foliage iscovered with larval feeding leaf whorls and eventually the foliage is destroyed.Most mature larvae descend to ground and pupate in the top soil layer.

Maraca pod borer is a major biological limiting factor for cultivation ofcowpea in Africa, especially West Africa and India (Taylor 1978, Saxena 1978).It is widespread in the Pacific and causes major damage to commonly grownlegumes (Waterhouse and Norris 1987). No reliable data exists on the yieldreduction in mungbean and other legumes because of the simultaneous presence ofother pests that attack the crop. In cowpeas Karel (1985) reported losses of 31%due to Maruca podborer in Tanzania. In India, Patel and Singh (1977) attributed10% yield loss and Lalasangi (1988) report over 37% yield loss due to Maruca incowpea.

Control Measures

Because of the hidden nature of larval stage, the damaging stage, and thatof the pupal stage, it is difficult to control Maruca podborer by chemicals or otherconventional means. Insecticides have been widely used in Asia, especially onyardlong bean where fresh pods are marketed as vegetables in Southeast Asia.However, due to very brief period soon after hatching before larvae enter buds orflower or pods, that the insect is exposed on the plant surface when insecticidescan come easily in contact with the pest and kill it, chemicals have to be appliedfrequently. This is not always economical. However, due to the lack of suitablealternative control measures, vegetable farmers keep on spraying yard-long bean.This kind of pesticide use has to decrease.

In field crops like mungbean and cowpea, insecticide use is much less dueto relatively shorted post-flowering period, compared to yardlong bean, when theinsect is especially active. However, the damage by the pest in these crops isunabated.

Biological control

Large number of parasites have been reported to feed on Maruca larvae andsome on pupae (Waterhouse and Norris 1987). However, in variably in all casesthe extent of parasitism is low. Don Pedro (1983) found Phanerotonia sp. andBraunsia sp. to be the most important parasitoids in Nigeria. However, theirparasitism did not exceed 7%. Okoye-Owuor et al. (1991) reported pest mortalityof 40.7 and 35.6% due to parasitoids and pathogens at two sites in Kenya. They

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reported presence of seven parasitoids, two predators, one nematode, and severalpathogens attacking Maraca. Antrocephalus sp. was the predominant parasitoids,however, observed parasitism contributed 3.25% and 3.8% at two sites. In India,Lalasangi (1988) recorded Bracon greeni and Apanteles taragamae on Marucalarvae. The latter parasite was recently found attacking Maruca larvae onSesbania indica at AVRDC in Taiwan. A parasitism of up to 92% of larvae wasobserved in summer. This is the highest parasitism of Maruca by any parasitoidanywhere reported so far. AVRDC is now exploring utility of this braconid incontrolling Maruca on mungbean, cowpea and yardlong bean.

Cultural control

At the International Center for Insect Physiology and Ecology in Kenya,has over the years found intercropping of sorghum and cowpea reduced borerdamage to both crops, including Maruca damage to cowpea (ICIPE). SinceMaruca attacks wide range of legumes, voluntary legume plants should bepromptly destroyed to reduce the carry over of the pest into the next season.


The entomological research at International Institute of TropicalAgriculture (IITA) in Ibadan, Nigeria, emphasizes breeding of cowpea cultivarsresistant to Maruca. They have screened over 7000 cowpea accessions and entrieswith varying levels of resistance have been identified and used in their host-plantresistance breeding. New breeding lines carrying varying levels of resistance toMaruca have been developed and made available to interested scientists. Pleasecheck 'IITA's annual reports for the latest information on this topic.

At the Asian Vegetable Research and Development Center (AVRDC) inTaiwan, mungbean germplasm consisting of over 5000 accessions have beenscreened for resistance to Maruca during the past two years. Several leastdamaged accessions have been selected for further screenings to confirm theresistance. This is a continuing research at AVRDC. Please check AVRDC'sAnnual Reports for the latest information.

Chemical control

Insecticides have been widely used for the control of Maruca on yard-longbean which is an important vegetable all over Southeast Asia especially during hotwet season when other vegetables are in short supply. In most cases chemicals areapplied weekly or more often prophylactically. A wide variety of chemicals, oftenin mixtures of two or more, are used irrespective of whether such use leads to anybetter control of the pest. This poses various health hazards to farmers andconsumers in addition to contamination soil and eventually water due to run-off.This misuse of chemicals is unlikely to stop soon unless alternative safe control

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measures are developed. Availability of various formulation of neem which givesequal or better control of Maruca on cowpea (Ramasubramanian and SundaraBabu 1991, Jackai and Oyediran 1991) could .replace the more toxic presentlyused chemicals.

For Maruca on cowpea, several commonly used insecticides such asendosulfan, carbaryl, methomyl, monocrotophos have been found effective (Singh1977, Lalasangi 1988). The first application should be made at least one weekbefore flowering and continued at a weekly interval until three weeks after peakflowering. No such information exists on Maraca control in mungbean.

Etiella zinckenella

Morphology

The white oval eggs (0.6 mm long) are laid singly or in batches of 2-12 onyoung pods, calyx or leaf stalk. Towards the end of incubation period of 3-16days, eggs turn pink. The first instar larvae are 1 mm long with yellowish bodiesand black heads. They wriggle violently if their pod is opened and they aredisturbed. There are five larval instars. Just before pupation larvae become greenwith dark pink stripes. Full-grown larvae are 15 mm long. Freshly formed pupaeare light brown but progressively turn dark brown to black as the time for adults toemerge approaches. Male pupae are generally larger, 8.5 mm long, than female,8.0 mm. Pupae can be found in soil 2-4 cm below the surface. Adult forewingsare brownish gray with a white strip along the leading edge of narrow fore-wings(Figure 5). Hindwings are transparent to opaque with darker outer edges. Wingspan is 24-27 mm.

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e 5. Adult of E. zinckenella.

Similarity with other pests

An additional morphologically similar Etiella species, E. hobsoni (Butler)infests soybean in Indonesia. The nature of damage by both species is practicallyidentical. Naito et al. (1986) gives details of distribution of both species inIndonesia. The minute morphological differences in egg, larvae, pupae and adultsare described by Naito et al (1986). The only substantial difference that can beused in distinguishing both species is found in adults. The ground color of theforewing of E. hobsoni is dark brown or dark reddish brown, without a whitecoastal streak found in E. zinckenella. The antemedial transverse fasca in E.hobsoni is orange edged with metallic scale. In contrast, the forewing of E.zinckenella is variably colored from reddish brown to purplish gray, but not dark,and has white coastal streak. The antemedia transverse fasca is orange brown toorange red, frequently with gold iridescence.

When the adults fold their wings at rest, the antemedial bands of the forewingsof the forewings of E. hobsoni seen as a straight transverse band across the wings,while those of E. zinckenella are not straight. Etiella hobsoni_ is generally smallerthan E. zinckenella; the length of the forewing of the foliner is 7.5 ? 0.6 mm, andthat of the latter 8.7 ? 0.7 mm.

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Biology

The white oval eggs (0.6 mm long) are laid singly or in batches of 2-12 onyoung pods, calyx or leaf stalks. A single female lays 60-200 eggs during herlifetime (Kobayashi, 1976). Incubation lasts 3-16 days, depending upontemperature. The first instar larvae are 1 mm long with yellowish body and blackheads. These larvae spend about half an hour moving about on the pod; they thenspin a small web, bore through pod pericarp covered by the web and begin feedingon the developing seeds. There are five larval instars. A number of larvae mayenter pod, but cannibalism reduces them to one or two. If the food supply in onepod is inadequate, they migrate to another. The larvae wriggle violently if theirpod is opened and they are disturbed. Just before pupation they become greenwith dark pink stripes. Larval development lasts 2G days.

Full-grown larvae are 15 mm long (Kobayashi, 1976) when they leave thepod to pupate in a cocoon in the soil, 2-4 cm below the surface (Figure 6).Pupation lasts for 1-9 weeks, depending on the temperature. After emergence, themoths live up to 20 days. They are brownish gray with white stripe along theleading edge of the narrow forewings. The wingspan is 24-27 mm (Hill 1975).

Figure 6. Eliella zinckenella pupa exposed fromthe cocoon in soil.

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Nature of damage

Pod injury in soybean by E. zinckenella is recognizable even when the larva isabsent. Large pods are marked with brown spot where the larva has entered; asthe larva within the pod develops, the buildup of faces causes soft rotten patcheson the pod. Seeds are partially or entirely eaten, and considerablefrass and silk are present. A large hole is evident where the larva has escaped topupate in the soil. In cowpea, lentil and pigeon pea, blossom drop and some poddrop occurs due to very small larvae feeding on the blossom and young pods.Usually one or two larvae can be found in each pod.

Economic importance

Etiella zinckenella is a cosmopolitan pest with worldwide distribution (Qu andKogan. 1984). It attacks cultivated legumes including cowpea, garden pea,limabean, mungbean, pigeon pea, common bean and soybean. Amongst all hostplants, soybean is by far the most preferred. This insect seems to have biotypes indifferent parts of the world. For example, it is a serious pest of common bean(Phaseolus vulgaris) in the USA but does not attack soybean there despite hugearea under .cultivation. In most of Southeast Asia, it is a threat to soybean butdoes not readily attack common bean. A sex pheromone blend which attractsEuropean strain in Hungary (Toth et al., 1989) is not effective against SoutheastAsian strain of the pest.

Damage to soybean in Southeast Asia is widespread. It damages about 10-15% pods in Taiwan, however, in Indonesia, the damage can reach 80% of pods(Talekar 1987). In the Philippine's Iloilo Province where soybean is a new crop,E. zinckenella damaged . 57% of the pods even in insecticide-protected plots(Litsinger et al. 1978b). In Iran E. zinckenella causes yield loss in. soybean ofabout 40% in the Province Lorestan and adjacent areas (Parvin 1981). In Indiathis pyralid infested 11.43 and 50.9% pods of lentils and peas, respectively. Thisresulted in yield loss of 10.6 and 23.9% respectively (Singh and Dhooria 1971).In Egypt, this pest is reported to cause 40% loss of yield in cowpea (COPR 1981).

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Control measures

Biological control

Etiella zinckenella has wide host range and occurs in the tropics practicallythroughout year. It also has large number of parasites despite cryptic nature ofthe pest. This presents an opportunity for biological control of this pyralid.However, no effort has been made either to introduce the natural enemies wherethe pest problem is serious or inundative use of any parasite. Sustained efforts inpursuing this field of research would lead to considerable reduction in pestdamage in countries like Indonesia where this pest is especially serious.

Sex pheromone

A four component sex pheromone has been identified for E. zinckenella inHungary (Toth et al., 1989). However, this chemical is not effective in SoutheastAsia, particularly in Taiwan (Toth et al., 1995). Development of a sex pheromonewould aid in detecting onset of infestation of the pest especially in soybean wherethe damaged plant parts-pods-are hidden in plant canopy. However, much workneeds to be done to develop the pheromone for southeast Asian strain of this pest.


To date, little effort has been made to breed mungbean or soybean cultivarsresistant to E. zinckenella although sources of resistance to this pest in soybeanare available (Talekar and Lin, 1994). This is because of the lack of adequateresearch in host-plant resistance breeding in southeast Asia.

Chemical control

Being an internal feeder it is difficult to control E. zinckenella byconventional means such as insecticide application unless chemicals are sprayedfrequently to kill the first instar larvae while they are still outside pod. The spraysshould be directed towards pods, and this requirement presents a mechanicalproblem in soybeans, as the leaf canopy completely covers the pods. In greenpods of other crops such as lima bean, snap bean which are sold as fresh podssuch pesticide use is prohibitive for health hazards reason. Among 54 insecticidetested by Stone (1965) carbaryl and azinphos-methyl and mexacarbate gavesatisfactory control of E. zinckenella_on lima bean. In Taiwan monocrotophos,triazophos, fenvalerate and quinalphos gave satisfactory control of E. zinckenellaon soybean (AVRDC, 1982). Triazophos and carbaryl, however, can causephytotoxicity in certain soybean cultivars.

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Integrated control

Since very little success has been achieved so far in controlling this pest byanything other than chemical control, no efforts have been made to develop IPMfor this pest in Southeast Asia or elsewhere. Unless additional information isgenerated through sustained research in host-plant resistance, biological controland sex pheromone, chances of development of usable IPM to reduce pest damageare meager.

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5. Storage Pests

Introduction

Among scores of species of bruchids — insects belonging to thecoleopterous family Bruchidae — that infest food legumes in the tropics, threespecies. Callosobruchus chinensis (L.), C. nmaculatus (F.) and C. analis (F.), infestmungbean (Vigna radiata [L.] Wilczek) in the field and during storage. Thefoinier two species are native of Asia and Africa. Mungbean, cowpea andpigeonpea serve as their principal hosts. Callosobruchus analis, an Asian native,is now found to be a pest of cowpea in Africa (Southgate 1978). Due to themovement of grains via trade, these pests. especially the former two, are nowfound in all six continents where they attack a wide range of pulses. Although thebruchids, commonly called pulse beetles or cowpea weevils, attack mungbean inthe field and storage. it is the infestation of grains during storage that results in thegreatest loss. In this section, therefore, after a brief discussion of theiridentification and biology, a detailed account of the nature of their damage andmeasures to minimize storage losses incurred by them will be discussed.

Identification

Due to the lack of adequate published reports on the systemics ofBruchidae, there have been numerous misidentifications of the bruchid species inthe past. However, two publications by Southgate (1958) and his colleagues(Southgate et al. 1957) have removed much of the confusion. Since certain oldnames are still being used in the literature, the synonymy of each of the threespecies that attack mungbean is summarized in Table 1 (Vazirani 1976).

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Table 1. Synonymy of three bruchid species that infest mungbean.

Callosobruchuschinensis (L )

Callosobruchusmaculatus (F )

Callosobruchusanalis (F )

Curculio chinensis L. 1758 Bruchus maculatus F 1775 Bruchus analis F. 1781

Bruchus pectinicorus L. 1767 Bruchus quadrimaculatus F. 1792 Bruchus jekeli All. 1847

Bruchus rufus De Geer 1775 Bruchus ornatus Boh. 1829 Bruchus glaber All. 1847

Bruchus scutellaris F. 1792 Bruchus vicinus Gyllen. 1833 Callosobruchus analis

Southgate, Howe et

Brett.1957

Bruchus bistriatus F 1801 Bruchus ambiguus Gyllen. 1833

Bruchus barbicornis F 1801 Callosobruchus maculatus Pic 1913

Bruchus elegans Sturm. 1826

Bruchus chinensis Sch. 1833

Bruchus adustus Mots. 1874

Callosobruchus chinensis

Mukerji & Chatterji 1951

Source: Vazirani 1976.

Certain striking morphological characters which differ in these species. andwhich are useful in bruchid identification, are described below.

Observation of the morphological characters of the adults provides theeasiest form of identification (Figure 1). The shapes of antennae and the hindfemur are two common characters that are used to easily distinguish the threebruchid species. In C. chinensis males, the fourth through apical segments arepectinate to highly pectinate whereas in females these segments are serrate. In C.maculatus the antennae are slightly serrate from the fourth through apical segmentand in C. analis antennae are wholly testaceous and not serrate. The hind femur inC. chinensis is ventrally bicarinate with a denticle situated on each carina near theapex. The outer tooth is blunt and the inner tooth is long and straight, and roundedat the tip. In C. maculatus the hind femur is ventrally bicarinate, with a large blunttooth on the outer carina and a sharp tooth of similar size on the inner carina. Both

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1

teeth are situated near the apex. In C. analis, the hind femur is usually ventrallybicarinate, with a large pointed tooth on the outer carina. The tooth on the innercarina is very minute or absent. Mukerji and Chatterjee (1951) give details ofdifferences in the genital structures of various bruchid species.

Since larvae and pupae are always hidden, their identity require greaterefforts. Vats (1974) and Begum et al. (1982) give details of distinguishingcharacters of bruchid larvae (Figure 2) and these characters can be used inconjunction with those of the adult's to confirm the identity of each species.Wightman and Southgate (1982) provide very useful information on thedistinguishing characters of eggs of nine bruchid species based on scanningelectron microscopic (SEM) studies. Southgate et al. (1957) and Southgate (1958)should be referred to in order to confirm the identity of the species discussed inthis paper.

Biology

Several studies, mainly in the Indian subcontinent, report on the biology ofCallosobruchus on various pulses (Rahman et al. 1943, Arora and Pajni 1957,1959, Rajak and Pandey 1965, Raina 1970). In general, the life cycle history of allthree species follows a typical coleopterous insect. There is very little differenceamong the three species. Raina (1970) made a detailed comparative study of thebiology of the three species reared on mungbean at 30°C and 70% relativehumidity (RH), a condition considered ideal for the development of the threebruchid species. The following information is extracted largely from his results.

Mating and oviposition

Adults mate within an hour after emergence from the seed. Mating lasts 5to 8 minutes in C. chinensis, 3 to 8 minutes in C. maculatus and 3 to 6 minutes inC. analis. Although the insects mated several times, only one mating is sufficientto ensure egg laying. Eggs are covered with a sticky substance which fastens theeggs to the seed surface (Southgate 1979). At the time of oviposition, C. chinensis

and C. niaculatus deposit a chemical

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OWNSINIM4

Figure 1. Adults of three species of Callosobruchus.A and B; C. chinensis, C and D: C. maculatus,E and F: C. analis. (Source: Raina, 1970)

45

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AnteclypeusLabrum

1t1 1

0

Antenna

Figure 2. Clvpeus, labrum, and antennae of larvae of U C. rnaculatus,® C. chinensis, and©C. analis (Source: Vats, 1974).

The bar length is 0.1 mm.

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oviposition marker' on the seed surface which has an ovicidal and arrestant action(Oshima et al. 1973, Yamamoto and Honda 1977, Honda et al. 1978). Thischemical, a mixture of fatty acids, triglycerides and hydrocarbons, prevents thehatching of more than one or two eggs per seed and helps regulate the pestpopulation and maximize use of the food. Yamamoto (1976) suggests that thischemical can be used as a possible oviposition inhibitor to control the bruchids.Certain edible oils (see discussion under Control Measures) give a similar ovicidaleffect.

Callosobruchus chinensis laid an average of 78 eggs over a period of eightdays; C. maculatus laid 128 eggs and C. analis 96 eggs over a nine-day period.Howe and Currie (1964) reported a slightly different fecundity data of the threespecies but this could be due to the selection of different host, cowpea, in theirstudy.

Usually one to three eggs are laid over an individual seed although as manyas five eggs in a study at AVRDC (unpublished) and seven in Raina's (1970) studywere found on a single mungbean seed, when some seeds were still without eggs.The number of eggs laid was significantly correlated to the seed size (r = +0.95) inone study at AVRDC (unpublished). The average incubation period was 3.5, 4 and5 days, respectively, for the eggs of C. chinensis, C. maculatus and C. analis. Egghatching for all three species ranged between 94% and 99%.

Larval stage

Soon after hatching the larva makes a hole in the seed coat, just underneaththe spot where the egg is laid, and enters the kernel where it feeds concealedinside the seed. When the eggs are laid on the pods, as in the case of insectinfestation in the field, the newly hatched larva makes a hole through the podcover, enters the developing seed and feeds and pupates inside the developingseed. Before pupation bruchid larva gnaws a circular hole until-only a thin layer or'window' of seed coat is left intact. The combined larval and pupal period was18.8, 20 and 23.5 days for C. chinensis, C. maculatus, and C. analis, respectively(Raina 1970).

Adult Stage

Adults of all three bruchid species emerge by cutting open the 'window' inthe seed testa. The entire development from egg to the adult stage takes an averageof 22.3, 24 and 28.5, days respectively, for C. chinensis, C. maculatus and C.analis at 30°C and 70% RH (Raina 1970). A similar developmental time wasobserved by Atwal et al. (1968) in C. analis on mungbean under similarenvironmental conditions. There was no difference in developmental time and lifespan between male and female in all three species and the sex ratio was 6:5, 7:6

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and 1:1 males to females, respectively, for C. chinensis, C. maculatus and C.analis. Callosobruchus chinensis males and females lived an average of 7.6 and7.4 days, C. maculatus, 8.2 and 7.6 days, , and C. analis, 6.8 and 8.0 days,respectively. Developmental mortality from egg to the adult stage was 23% in C.chinensis, and only 9% for each of the remaining two species. Most of themortality observed was in the egg and early larval instars.

Nature and Extent of Damage

Damage in the field

Although bruchids attack mungbean in the field, damage to seeds per se isonly minor. However, when infested seeds are stored, the adults emerge and layeggs on the neighboring seeds. This secondary infestation is much moredamaging. Banto and Sanchez (1972) report from 7.8% to 9.9% seed infestationby C. chinensis at the time of harvest. Infested seeds harboredbruchid larvae of varying stages of development.

Damage during storage

Three aspects of bruchid damage are of particular importance: (i) theoverall weight loss; (ii) changes in nutritional quality and presence of off-smellingby-products of insect infestation; and (iii) loss in seed viability.

Seed weight loss. Weight loss can be a direct consequence of bruchids feeding onthe seed. It may also occur as a result of accelerated loss of moisture due toperforation by bruchids of the mungbean seed. Vimala and Pushpamma (1983a)found that the level of insect infestation, as assessed by insect count, kerneldamage, frass content and weight loss, increased with the period of storage up toone year. The percentage of kernels damaged in mungbean increased from 0.53%at the beginning of storage to over 16% after one year of storage and weight lossfrom 0.32 % to 7.22 % o during the corresponding period. Gujar and Yadav (1978)report a weight loss of 55.6% to 73% in individual seeds damaged by a single C.maculatus and 30.2% to 55.7% by C. chinensis in one generation. Banto andSanchez (1972) reported total destruction of seeds when newly harvested, infestedmungbean seeds (9.9% seeds damaged) were stored for three months. Infestedseeds were unfit for human consumption.

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Seed quality changes. Loss or denaturation of proteins and vitamins reduce thenutritional quality of pulses. In addition, the presence of insect excrement, castlarval skins, pieces of insect chitin or dead insects can have an abrasive effect onthe human alimentary canal.

Vimala and Pushpamma (1983a) found up to 45 dead/alive insects per 100g mungbean seeds after one year storage when practically no insects were presentat the initiation of storage. Mungbean seeds contained from 0.39% to 0.41% frassafter one year storage when practically none existed at the initiation of storage. Nosignificant changes occur r ed in the grain moisture content. Uric acid, a metabolicby-product of insects which is present in insect excrement, increased from barelydetectable levels at the initiation of storage to up to 31.5 g/100 g mungbean seeds.The authors reported a significant positive cor relation between the number ofinsects and the uric acid contents of rnungbean seeds. The uric acid contentreached above safety level after eight months of storage and still remained abovethat level after one year of storage. Similarly, Singh et at (1982) observed thatfree fatty acids, reducing sugars and uric acid contents increased with the increasein infestation of C. chinensis during a five-month period in three mungbeancultivars. The uric acid content was considerably greater in mungbean than inblack gram as was the bruchid infestation. Doharey et al. (1983) also observed anincrease in free fatty acids and alcoholic acidity in mungbean during 120 days ofstorage due to C. chinensis and C. maculatus infestation. They also reported thatC. chinensis infestation increased protein content from 22.15% to 47.14%, and C.maculatus infestation increased from 22.15% to 57.55%, whereas in the check itonly increased to 32.34%. No explanation for this change in protein content isoffered by the authors. However, this anomaly appears to be due to the use of aninappropriate analytical method to determine the protein concentration. Dohareyet al. (1983) estimated protein by analyzing total nitrogen rather than proteinnitrogen (see Gujar and Yadav 1978). Some of nitrogen they analyzed could havecome from uric acid rather than protein.

Vimala and Pushpamma (1983b) found that the starch content of mungbeanwas decreased by 6.19% after one year of storage and certain changes wereobserved in the reducing and nonreducing sugars and digestibility of storedmungbean seeds. But whether this is due to bruchid infestation or a normal changeduring storage is unknown. Pingale et al. (1956) found a reduced concentration ofthiamin in stored mungbean seed infested with C. chinensis. The reduction wasroughly in proportion to the amount of insect damage to the seed. In addition,insect damage increased fat acidity and caused slight denaturation of protein.

I

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Seed viability loss. Even slight feeding damage by bruchids to the embryo impairsgermination. Such feeding on the cotyledon will not affect germination but thevigor of the young seedling will be reduced, as in cowpea, due to similar damageby Acanthoscelides obtectus (Say) (Chin 1980). In a study with three mungbeanand two blackgram cultivars, Singh and Sharma (1982) observed a progressiveincrease in seed damage and a proportional decrease in seed geimination during afive-month storage period. In mungbean, the seeds damaged by C. maculatusvaried from 42.53% to 57.77% and loss in seed germination from 47.53% to70.60%. There was much less seed damage and less reduction in viability in blackgram seeds. Significant differences were observed in seed damage and viabilityamong both mungbean and blackgram cultivars. As the level of Callosobruchusinfestation to mungbean seed kernels increased from 4.33% to 16.67%, the loss ofviability increased from 16.23% to 28.90% (Vimala and Pushpa mna 1983c).

Control Measures

The nature and extent of bruchid damage described above .entails soundcontrol practices in order to protect the harvest from the ravages of bruchids,especially during storage. Because of the primitive nature of the storage facilitiesin many of the villages where most of the mungbean crop is grown on smallfarms, the small volume of produce, and the fact that the grains are frequentlyused for consumption, the use of fumigants or other insecticides is impractical.For bruchid control, therefore, sound farming practices, good storage facilities,coupled with nonchemical control measures are necessary. These measuresinclude, the drying of seeds before storage, use of bruchid resistant cultivars,nontoxic chemicals such as vegetable oils, sex pheromones, biological control,and as a last resort, the use of selective and safe chemicals.

Although several hymenopterous parasites have been reported to attackbruchids in the egg, larval and pupal stages, their impact on populations isinsignificant. Their use in biological control of bruchids during storage is notpractical.

Drying of Seeds

A seed moisture content of below 10% impairs normal activity anddevelopment of storage insects and at moisture levels below 9.5% certain of thesepests do not even oviposit (Girish 1983). Besides, most insects die within 10 to20 minutes at temperatures of from 55° to. 60°C. The tropical sun is helpful inheating and drying the grains, thus ridding the seeds of bruchid infestation.Yoshida and Gichuki (1983) found that when adzuki bean seeds are spread in thesun in a layer of 3 cm deep, 550C was reached in 1.5 hours and maintained for 4hours and 40 minutes. With a layer of 1.5 cm deep, 55°C was reached within 30

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minutes and maintained for 4 hours. Sun drying of the grains before storage willthus not only reduce the risk of insects from primary infestation in the field beingcarried into storage, the reduction in seed moisture will minimize reinfestationfrom secondary sources. Low seed moisture also prolongs the effectiveness ofvegetable oil and insecticide treatments which might be used to protect the seedsfrom storage insects (Doharey et al. 1984, Talekar and Mookherjee 1969).

Resistant Cultivars

Cultivars resistant to bruchids are yet to be developed althoughconsiderable progress in this field has been made. Due to its low cost and ease inuse, coupled with the limited utility of other methods, especially chemical control,the use of bruchid resistant mungbean cultivars has a considerable potential. Sincebruchids can infest mungbean pods in the field, as well as seeds in storage,resistance either in the pod, seed or both is desirable.

Resistance in the field. Doria and Raros (1973) screened 66 mungbean cultivarsfor resistance to C. chinensis damage to the pods. None of the entries was resistantto oviposition but resistance to larval survival was evident in EG Glabrous, EG-MG-4, and EG-MG-7. Mungbean accessions UPCA 23, 25 and 325 had the leastnumber of eggs and lowest larval survival.

At AVRDC, field, greenhouse and laboratory experiments on 525 Vignaaccessions at AVRDC led to the identification of the two accessions with differentmodes of resistance to C. chinensis (Talekar and Lin, 1981). Accession VM 2011was least damaged when insect infestation occur r ed via pods, whereas VM 2164was highly resistant in seeds. Both accessions are V. mungo. Hairiness on thepod in the case of the former, and antibiosis in the latter, are believed to be theresistance mechanisms involved (Talekar and Lin, 1981, 1992). Interspecificcrosses to incorporate bruchid resistance of VM 2164 into mungbean proved to bedifficult (AVRDC 1988, Fernandez and Talekar 1990). Hence a breeding programutilizing these resistance sources was discontinued. Renewed screening of onlymungbean germplasm resulted in identification of two mungbean accessions, V2709 and V 2802, which have moderate to high levels of resistance to C. chinensis(AVRDC 1990). Accession V 2709 (PI25393),. also designated as LM501, is aland race from India and V 2802 (PI 25461) comes from the Philippines.

In a series of laboratory and greenhouse tests, characteristics of resistancein V 2709, V 2802 and VM 2164 were investigated (Talekar and Lin 1992).Oviposition and emergence of first generation adults of C. chinensis weresignificantly reduced in pods of V 2709 and V 2802, compared with thesusceptible check VC 1973A (Table 2). Artificial seeds prepared from mixturesof varying quantities of powdered seeds of resistant and susceptible accessions

"(Shade et al 1986) were then exposed to infestation by C. chinensis. As the

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concentration of resistant accessions in the artificial seeds increased, the numberof C. chinensis adults emerging after feeding as larvae in such seeds decreased(Figure 1). Adults that emerged from the artificial seeds made from mixtures ofresistant and susceptible accessions were significantly lighter than from artificialseeds made from the susceptible accession alone. These results suggest thatantibiotic factors may be present in these resistant accessions.

Table 2. Bruchid infestation on pods of resistant and susceptiblemungbean accessions.

Accession No. of No. of No. of adultseggs/pod adults/pod /10 seed locules

First flushV2709' 5.95+3.68 0 0

V2802 14.75+12.85 1.41+_2.73 1.48+2.65VC1973A 44.30+26.47 25.79±6.41 29.12±6.84

LSD 5% 12.64 2.64 2.72

LSD 1% 16.93 3.55

Second flush

3.64

V2709 14.69_+2.47 0 0

V2802 24.45+7.64 0.87+0.45 1.03+0.55

VC1973A 74.68+24.37 25.31+9.14 30.09±9.59

LSD 5% 9.47 3.35 3.54

LSD 1% 12.72 4.51

Third flush

4.75

V2709 17.85_+5.52 0 0V2802 18.15+6.52 0.49_+0.41 0.66±0.58VC1973A 25.80+4.58 14.32±4.18 19.36+5.81

LSD 5% 4.08 1.52 2.14LSD 1% 5.47 2.04 2.87

Data are means (+SEM) of 15 replicates, one plant per replicate.

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VM 2164

V 2802

V 2709

0 25 - 50 75 100% OF RESISTANT SEED IN SUSCEPTIBLE

Figure 3. Emergence of C. chinensis adults from artificialseeds of susceptible accession VC 1973A containingvarying levels of seeds of resistant accessions V 2709,V 2802, VM 2164.

A V. sublobata accession TC1966 was found to have high levels ofresistance to C. chinensis (AVRDC 1991). This species is cross-compatible withmungbean. Japanese researchers report vignatic acid, a cyclopeptide alkaloid, asone of the factors responsible for resistance of TC1966 to C. chinensis(Sugawara et al. 1996). This chemical did not show any cytotoxic effect onChinese hamster ovarian tumor cells and some humor tumor cells, indicatingthereby that a resistance cultivar bred from using TC 1966 as the resistant parentcould be safe for human consumption. This accession along with V 2709 and V2802 are now being used at AVRDC to breed bruchid resistant mungbean.

The agromyzid resistant V. glabrescens accession, V 1160, also showsantibiosis type resistance to C. chinensis (AVRDC 1990).

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Rajapakse et al. (1983) screened 11 mungbean cultivars for resistance to C.chinensis. Cultivars Uthong 1, H101 and CES 87 were relatively resistant as thenumber of insects emerging from the seeds of these cultivars were the least and C.chinensis required a longer period to develop from egg to adult in these cultivars.Epino and Morallo-Rejesus (1982) screened 60 mungbean accessions forresistance to C. chinensis in seeds. Based on the first generation bruchids thatemerged, UPCA accessions 11 and 30 showed moderate to high levels ofresistance. The seeds of resistant accessions adversely affected the survival,increased developmental time and reduced growth index and body weight.

Use of vegetable oils and plant products

The use of edible oils to protect stored grains, especially pulses, againstinsect pest damage is an ancient method of pest control in India. In addition tcedible oils, extracts and plant parts of certain readily available plants, such asneem (Azadirachta indica Adr. Juss.) have also been utilized in villages wherestorage facilities are poor. In light of the adverse effects of insecticides on theenvironment, these methods of pest control are now attracting greater attentionand research input.

In studies at AVRDC, mixing of soybean or groundnut oil at the rate of 2 to3 ml/100 g seeds gave mungbean considerable bruchid protection for up to twomonths. Neither treatment affected seed germination but prolonging groundnut oiltreatment to five months reduced seed germination considerably (AVRDC 1976).In a similar study Varrna and Pandey (1978) utilized oils of coconut, mustard,groundnut, sesame and sunflower mixed with mungbean seeds at the rate of 0.3 gper 100 g (w/w) of seeds. Oviposition of C. maculatus was completely inhibitedwhen coconut and mustard - oils were used; very few eggs were found onmungbean seeds when other oils were used. Development of the adult populationwas prevented for at least five months, and the viability of the treated seeds was

. unaffected. Coconut oil was the most effective followed by mustard, groundnutand sesame. Pandey et al. (1981) used oils of cotton seed, rice bran and sal(Shorea robusta Gaertn. f.) at the rate of 0.3 to 0.5 g per 100 g of mungbeanseeds. All three oils protected the seeds for five to six months. Neither ranciditynor free fat acidity increased significantly in the treated seeds, and there was noadverse effect on seed viability. Doharey et al. (1984) studied the utility of oils ofcoconut, groundnut, mustard, rice bran, safflower, sesame and taramira (Erucasaliva Mill.) in protecting mungbean seeds, adjusted to the moisture content of9.6%, 10.8% and 12.8%, against C. chinensis and C. maculatus. A concentrationof 1% oil (w/w) protected the seed from both species. A higher grain moisture of12.8% significantly reduced the efficacy of safflower oil against both bruchid

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. species. In a separate study Sujatha and Punnaiaii (1985) found that mungbeanseeds can be effectively protected from C. chinensis by treatment with the oils ofsesame, cotton seed, oil palm or neem at concentrations of 0.25% and those ofgroundnut or coconut at 0.5%. Treatment with all the oils at 0.125% resulted inlower infestation than the control.

In a study on the mode of action of groundnut oil, Sharma and Srivastava(1984) found a 90% reduction in oviposition by the oil treatment. Bruchid eggsaffixed on the oil-treated seeds had 90% of the eggs died within 48 h as a result ofcoagulation of protoplasmic contents in the embryo. h the remaining eggs, theinhibition of embryonic development could be observed up to a certain extent butthe embryo died soon thereafter. - Singh et al. (1978), however, found that thegroundnut oil treatment of cowpea seeds prevented the emergence of bruchidprogeny from the seeds rather than affecting the oviposition or mortality of thebruchid adults. The oil entered the eggs of C. maculatus through the micropile andin 1- to 2-day-old eggs, protoplasmic movement stopped and the protoplasmcoagulated. In 3- to 5-day-old eggs where the larvae are partially or fully folmed,larval death occurred within minutes of the entry of the oil.

Jotwani et al. (1968) mixed 0.5, I or 2 g of crushed neem seeds with 100 gof mungbean seeds and observed the damage by C. maculatus in storage overseveral months. At the end of eight months, only 9.8% of the seeds were damagedin 2.0 g, 11.9% in 1.0 g and 18.8% in 0.5 g neem treatments, whereas in thecontrol 59 % seeds were infested by bruchids . In a second experiment, theauthors found that treatment with up to 2.5 g crushed neem seed per 100 gmungbean seeds reduced the bruchid oviposition considerably over a period offour months. Neem apparently repelled the adults from laying eggs on the treatedseeds. In similar experiments, Yadav (1985) applied 2 to 50 mg neem seed oil per10 g mungbean seeds and confined adults of C. analis, C. chinensis or C.maculatus over the treated seeds. Treatment of 50 mg neem oil preventedoviposition of C. maculatus as against 40 mg in the remaining two species.Dosages of 30, 10 and 20 mg neem seed oil suppressed adult emergence in thethree species, respectively. This was due to the toxic action of neem oil against thebruchid eggs (Yadav 1985). Rajasekaran and Kumaraswami (1985) obtainedcomplete control of C. chinensis when extracts of karanj (Pongamia glabra Vent.)and neem were coated on mungbean seed at the rate of 0.6% v/v and 0.8% w/w,respectively. In a series of laboratory tests in India, Qadri (1985) showed thatneem extract synergized the toxicity of custard apple (Anona sp.) extract, andgarlic extract synergized the toxicity of oleoresin obtained from chrysanthemum,to C. chinensis.

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In a series of experiments, scientists at USDA's Stored Products InsectsResearch and Development Laboratory found that oils of several citrus fruits,extracts of black pepper (Piper nigrum L.), dill (Anethum graveolens L.), Chinesecinnamon (Cinnamomum cassia Nees ex Blume) are toxic to several insect pestsincluding C. maculatus (Su et al. 1972, Su 1978, 1985a,1985b). Similarly,Chander and Ahmed (1983) obtained good protection of mungbean against C.maculatus by mixing the powders of rhizomes of Acorus calamus L. (1%) andCucuma zedoaria (Christm.) Roscoe (5% ) and seeds of Carum roxburghianunrBenth. Pranata (1984) found turmeric (Curcuma longa L.) powder extract to betoxic to the adults of C. maculatus when the extract was mixed with mungbeanseeds. In one case vapors of Acorus calamus extract showed chemosterilant effecton C. chinensis. Exposure of adult females to the vapors reduced fecundity andcaused regression in the terminal follicle at the vitellarium (Bhaskar et al. 1976).

Attempts were made to contiol Callosobruchus chinensis (L.), withfishbean, Tephrosia vogelii (Hook, F.), foliage powder. Mungbean seeds werecoated' with 1, 2 or 4 g fishbean leaf powder/kg seeds and stored in nylon netsbags at room temperature for one year. Seed samples withdrawn from the bagsonce every 2 weeks were exposed for 24 hours to bruchid adults and insectmortality, number of first generation adults emerged after 1 month, and percentageof seeds damaged were recorded throughout the year. All dosages of fishbeanpowder remained highly effective for up to 1 year when the test was discontinued(AVRDC 1997). At every observation mortality of insects confined over the seedswas practically 100%. Very few, if at all, first generation adults emerged from theseeds and damaged seeds rarely exceeded 2% when damage to untreated seedsometimes passed 70% seeds. Dried fishbean which is used by certainsubsistence farmers in Africa to control insect pests and the foliage of whichcontains rotenone, therefore, is very useful in protecting mungbean seed frombruchid damage for up to three cropping seasons. The leaf powder coatedmungbean seeds, however, cannot be used for human consumption until they arethoroughly washed to remove all residues of the foliage powder.

Sex pheromones

Utilization of sex pheromone chemicals represents the safest form of pestcontrol. Although this approach has not been used in the control of any of thethree bruchid species which attack mungbean, recent studies have pointed out theexistence of sex pheromone in bruchids and their potential, especially for pestmonitoring purposes. In a laboratory study at AVRDC (1976) which utilizedvirgin females and unmated males, blowing of air over virgin females placed in anolfactometer attracted large number of males towards the virgin females. One- totwo-day-old females attracted a greater number of males than the older ones. Themales showed characteristic excitatory behavioral response including rapidantennal movement and extension of wings. Similar observations were made for

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C. maculatus by Rup and Sharma (1978) and Qi and Burkholder (1982). Thechemical properties of the sex pheromone isolated from C. chinensis have beendescribed (Honda and Yamamoto 1977, Tanaka et al. 1982). The pheromoneconsists of a mixture of callosobruchusic acid [(E)-3, 7-dimethyl-2-octenedioicacid] and several hydrocarbons. Neither the acid nor the hydrocarbons are activealone, their effect is synergistic.

Burkholder and Ma (1985) give details of the use of pheromones formonitoring various storage pests. The presence of a single bruchid in a trapindicates initiation of infestation and at such time suitable treatments can beutilized to reduce further loss. Bruchid pheromones can be utilized in the field in asimilar fashion to monitor primary infestation of bruchids.. The presence ofbruchids in the pheromone-baited traps will indicate initiation of infestation ofpods. At this time the whole crop can be sprayed with a suitable chemical toprevent further spread of the pest or the seeds from the infested field can be driedthoroughly to reduce grain moisture to below 9%. The heat of the drying will alsokill the larvae, pupae and possibly adults inside the seeds and a reduced grainmoisture level will considerably reduce the secondary infestationduring storage.

Chemical control

The appropriate insecticide when used properly gives assured andimmediate control of insect pest during storage as it does in the field. However,the use of insecticides to protect mungbean in storage has serious limitations onsmall farms. Firstly, the grains are stored for short duration, in most cases fromseason to season. During this period the seeds are often used for familyconsumption. Under such conditions mixing grains with insecticides, even ofrelatively low persistence, is not advisable. Secondly, the use of fumigants is notpractical because of the small sized produce and special precaution and trainingrequired to handle fumigants. Also in most cases mungbeans, along with othergrains, are stored in living quarters where the use of furigants poses hazards.Thirdly, mungbean is still a low priced low input crop and use of insecticides maybe uneconomical. Under such circumstances alternative methods, such as use ofvegetable oils, clean cultivation, storing seeds after thorough drying in a cleanstorage space, will assure protection from bruchid attack.

In community storage and large-scale commercial storage facilities,insecticides can be applied by mixing with seeds, spraying the surface of bulkstorage or stacks and by fumigation to protect mungbeans from bruchids.

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Insecticide dusts or sprays. Initially the storage space can be disinfected byspraying the area with suitable chemicals such as fenitrothion or chlorpyrifosmethyl at the rate of 1 g a.i./m 2 (COPR 1981). Subsequent risk of reinfestation canbe reduced by spraying the surfaces of loosely stored grains or bags at intervals ofabout eight weeks with the same chemicals. In the past mixing seeds with lindaneor pyrethrum dusts was suggested to control bruchids. However, due to theavailability of chemicals such as malathion, DDVP, fenitrothion, pirimiphosmethyl, which are less toxic to mammals and less persistent than lindane but morepersistent and cheaper than pyrethrum, use of lindane and pyrethrum are no longerrecommended. These chemicals can be mixed at the rate of 5 ppm with grainsmeant for long storage. Among the synthetic pyrethroids Duguet and Wu (1986)

applied deltamethrin at the rate of 0.75 and 1.00 ppm to artificially infestedcowpea in storage. This treatment protected the grains against C. maculatus for upto six months, when pirimiphos methyl dust applied at the rate of 10 ppm waseffective for only three months. In China spraying of mungbean seeds withdeltamethrin plus piperonyl butoxide at the rate of 0.25, 0.50 and 1.00 ppmprotected the seeds against C. chinensis for up to 228 days (Duguet and Wu1986). Synthetic pyrethroids, which are as selective for their less toxic effect onmammals but higher toxicity to insects as natural pyrethrins, but which are morepersistent than the natural pyrethrins, have promise in protecting mungbean seedsin storage. However, their dosages should be carefully chosen to give protectionfor only the intended length of storage so that it will not leave excessive residues.

Recently, Davis et al. (1984) found that exposure of adults of C. chinensisand C. maculatus to the dust of tricalcium phosphate, a commonly used fertilizer,causes complete mortality in 6 to 8 hours. When insects were exposed to thecompound mixed with snapbean (Phaseolus vulgaris L.) seeds at 0.01% to 0.25%concentrations, the number of Fl adults of C. chinensis that emerged was greatlyreduced. -Similar results were obtained when C. maculatus adults were exposed tocowpea seeds treated with tricalcium phosphate and, in fact, at doses of 0.1% andabove no Fl adults emerged. Tricalcium phosphate is readily available andrelatively inexpensive. However, its mode of action and possible health hazardsassociated with mixing with grain need further study before this treatment can besuggested to small-scale producers.

Fumigation. Like the above-described insecticide trea(inents, fumigation ofmungbean is. practical only in large-scale storage facilities. The greatest advantageof fumigation is the property of the fumigant to penetrate through the layers ofgrain and reach the target insect. Fumigants also penetrate through the feedingholes and oviposition windows in the seed and kill the larvae and pupae which arenot reached by conventional insecticide application. In addition fumigants arecapable of penetrating cracks and crevices which might harbor insects from thepreviously infested grains. Fumigation treatment does not leave persistent toxicresidues and treated grains can be utilized after one to two days of aeration.

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Several fumigants have been tested for their effectiveness against bruchidsinfesting various legumes and a few have proved to be more effective than theothers (Singh and Srivastava, 1980, 1983, Mundhe and Pandey 1980, El Sayedand Kamel 1978, Tsuruta and Tadauchi 1983, Abu and Muthu 1985, Sadomov1984). The adult stage is the most susceptible and the pupal the most resistant tofumigant action. The species of the host food legume does affect the susceptibilitybut such influence is of minor significance. Among the fumigants tested,phosphine is the most effective and convenient to use, especially for small-scalestorage. The chemical is available in convenient pellet or tablet forms. Quantitiesof mungbean can be packed into jute bags with polyethylene liners into whichpellets or tablets are placed. The bags are then sealed and left for four to five daysduring which the phosphine gas penetrates through the layers of grains and killsthe bruchids. Usually Ito 1.5 g tablet per cubic meter space is a suitable dose toachieve complete disinfestation. If kept sealed, the phosphine treatment will alsoprevent insect reinfestation.

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