hoy: myths of managing resistance 385 introduction to...

Hoy: Myths of Managing Resistance

385

INTRODUCTION TO THE SYMPOSIUM,THE MYTHS OF MANAGING RESISTANCE

M

ARJORIE

A. H

OY

Entomology & Nematology Department, University of Florida,Gainesville, FL 32611-0620

Resistance to pesticides in arthropod pests is a serious and increasingly perplexingproblem in Florida, the USA, and the world. Resistance to pesticides already has cre-ated significant economic, ecological, and public health problems in agricultural,household and garden, and medical/veterinary pest management programs. Exten-sive research has been conducted on diverse aspects of pesticide resistance, and wehave learned much during the past 40 years. However, to some degree, much of thediscussion about ‘resistance management’ has been based on ‘myths’. As an organizerof this symposium, one of my goals was to stress that managing resistance is a formi-dable task that will remain a perpetual pest management dilemma, because resis-tance is a fundamental survival response to stress by arthropods.

Five papers were presented in this symposium at the 1994 annual meeting of theFlorida Entomological Society, but one manuscript regarding the response by indus-try to resistance could not be published in this series.

In the first paper, Gary Leibee and John Capinera assess the impact of resistanceto pesticides in Florida and cite examples of resistance that limit pest managementoptions.

Julie Scott describes what we currently know about the molecular genetics of ar-thropod resistance to pesticides. The number of genes identified, and the diversity oftheir effects on the physiology of arthropods, verify that resistance is a normal re-sponse to diverse environmental stresses. ‘Pesticide resistance’ is part of a generalstress response with a long evolutionary history.

Leah Bauer describes what we know about resistance to various toxins of

Bacillusthuringiensis

(B.t.)

strains.

B.t.

provides microbial control of an increasingly diversegroup of arthropod species and is an increasingly important tool for integrated pestmanagement programs. The deployment of transgenic crop plants containing

B.t.

toxin genes is likely to be an effective method for inducing resistance in agriculturalpests. Despite the diversity of

B.t.

toxin genes isolated and cloned, cross resistancesare common. Thus,

B.t.

toxin genes are limited resources.Finally, I discuss a variety of resistance management methods and point out that

we cannot really

avoid

resistance—we can only

delay

its onset. I argue that resistancemanagement needs a paradigm shift that can best be accomplished if we recognizethat pest management must be changed from a single-tactic strategy to a multi-tacticmode. Delaying resistance, whether to traditional pesticides or to transgenic plantswith toxin genes, will require that we develop truly integrated pest management pro-grams, incorporating all appropriate tactics, including host plant resistance, culturalcontrols, biological controls, genetic controls, and biorational controls. Pesticidesshould be reserved for situations in which they perform best—as tools to resolve anunexpected pest population outbreak. Effective, fully-integrated IPM programs willdelay resistance because the number and rates of pesticide applications can be re-duced.

386

Florida Entomologist

78(3) September, 1995

PESTICIDE RESISTANCE IN FLORIDA INSECTS LIMITS MANAGEMENT OPTIONS

G

ARY

L. L

EIBEE

1

AND

J

OHN

L. C

APINERA

2

1

Department of Entomology and Nematology,University of Florida,

Central Florida Research and Education Center,Sanford, FL 32771

2

Department of Entomology and Nematology,University of Florida,Gainesville FL 32611

A

BSTRACT

Pesticide resistance in Florida was characterized through a survey and literaturereview. The survey was conducted in 1994 among public-sector entomologists to de-termine the current and future status, extent, context, pattern, and instances of pes-ticide (insecticide and acaricide) resistance in Florida. Results attested to the impactof pesticide resistance on the management of numerous arthropods in Florida.Twenty-five examples of insecticide and acaricide resistance were cited by survey re-spondents in agricultural, ornamental and landscape, medical and veterinary, orhousehold and structural pests. It remains possible to manage most arthropods by us-ing chemical pesticides, but the current and anticipated lack of efficacious materialsthreatens current practices in some areas. Trends in extent, context, or patterns of re-sistance were noted as follows: high value crops, frequently treated arthropods,smaller arthropods, and pyrethroids were all considered factors associated with resis-tance. Insecticide resistance and its management were reviewed in depth for the leaf-miner

Liriomyza trifolii

and the diamondback moth,

Plutella xylostella

, two majorinsect pests in Florida for which management options have become severely limitedbecause of insecticide resistance. Both cultural practices (continuous cropping, isola-tion, transport of infested seedlings) and pesticide use patterns (frequent applicationof broad spectrum pesticides) contributed to

L. trifolii

and

P. xylostella

resistance de-velopment. The history of pesticide resistance in these two insects is probably typicalof pest resistance in Florida and may portend similar future problems unless depen-dency on pesticides for pest suppression is reduced through adoption of IPM philoso-phy and practices.

Key Words: Insecticide resistance,

Liriomyza trifolii

,

Plutella xylostella

.

R

ESUMEN

La resistencia a los pesticidas en la Florida fue caracterizada a través de una en-cuesta y una revisión de la literatura. La encuesta fue conducida en 1994 entre los en-tomólogos del sector público para determinar el estado presente y futuro, extensión,contexto, patrón e instancias de la resistencia a pesticidas (insecticidas y acaricidas)en la Florida. Veinte y cinco ejemplos de resistencia a insecticidas y acaricidas fueroncitados por los que respondieron la encuesta sobre plagas agrícolas, de ornamentalesy de jardines, de importancia médica y veterinaria, o domésticas y de otras estructu-ras. Parece posible manejar la mayoría de los artrópodos usando pesticidas químicos,pero la falta actual y anticipada de materiales amenaza las prácticas presentes en al-gunas áreas. La tendencia en la extensión, contexto, o patrones de resistencia fuecomo sigue: cultivos de alto valor, artrópodos frecuentemente tratados, pequeños ar-trópodos, y piretroides fueron todos considerados como factores asociados con la resis-tencia.

Leibee and Capinera: Symposium on Pesticide Resistance

387

La resistenca a los insecticidas y su manejo fueron revisados en profundidad parael minador de las hojas

Liriomyza trifolii

y para la polilla de la col,

Plutella xylostella

,los insectos plagas principales en la Florida para los cuales las opciones de manejo sehan tornado severamente limitadas debido a la resisitencia a los insecticidas. Tantolas prácticas culturales (cosecha continua, aislamiento, transporte de plántulas infes-tadas) como los patrones de uso de pesticidas (aplicación frecuente de insecticidas deamplio espectro) contribuyeron al desarrollo de la resistencia de

L. trifolii

y

P. xylos-tella.

La historia de la resistencia a los pesticidas en estos dos insectos es probable-mente típica para la resistencia de las plagas en la Florida, y podría significarproblemas futuros similares a menos que la dependencia de los pesticidas para la su-presión de las plagas sea reducida a través de la adopción de filosofía y prácticas de

MIP.

Insecticide resistance has had an impact on the management of insect pests inFlorida since the mid-1940s following the widespread adoption of synthetic insecti-cides, especially the organochlorines, organophosphates, and pyrethroids. Numerousanecdotal reports exist, wherein consistently effective insecticides have become inef-fective and remained so for several seasons. Such reports have been considered ampleevidence of resistance development (Hoskins & Gordon 1956). In fact, Genung (1957)provided strong evidence based on anecdotal reports and data from field efficacy trialsfor resistance development in the cabbage looper,

Trichoplusia ni

Hubner, importedcabbageworm,

Artogeia rapae

(L.), a

Liriomyza

sp., and leafhoppers,

Empoasca

sp., ata session of the Florida State Horticultural Society Meeting in 1957 entitled “Sympo-sium-Vegetable Insect Resistance to Insecticides in Florida” (Brogdon 1957). Resis-tance episodes in Florida have also been documented in a number of species withlaboratory studies in which concentration-mortality response has been used to com-pare resistant and susceptible strains. Much of this work has been conducted withinthe last 10 years and involves species such as cabbage looper (Shelton & Soderlund1983), diamondback moth,

Plutella xylostella

(L.), (Leibee & Savage 1992a, Shelton etal. 1993, Yu & Nguyen 1992), silverleaf whitefly,

Bemisia argentifolii

Bellows & Per-ring, (G. L. L. unpublished data), house fly,

Musca domestica

L., (Bailey et al. 1970,Bloomcamp et al. 1987), German cockroach,

Blatella germanica

(L.), (Milio et al.1987, Koehler 1991, Hostetler & Brenner 1994),

Liriomyza trifolii

Burgess (Keil &Parella 1990, G. L. L. unpublished data), fall armyworm,

Spodoptera frugiperda

(J. E.Smith), (Pitre 1988, Yu 1992), cat flea,

Ctenocephalides felis

(Bouche), (El-Gazzar etal. 1986), and citrus rust mite,

Phyllocoptruta oleivora

(Ashmead), (Omoto et al.1994).

In hopes of providing a better understanding of current pesticide resistance and itsconsequences in Florida, we report here the results of a recent survey of public-sectorentomologists conducted to assess the extent of pesticide resistance in Florida, and itscurrent and potential impacts. In addition, we provide an in-depth account of two im-portant insect pests of vegetables in Florida, the dipterous leafminer

L. trifolii

andthe diamondback moth, for which management options have become extremely lim-ited because of insecticide resistance.

R

ESISTANCE

S

URVEY

During the spring of 1994, 16 public-sector entomologists were sent survey formsto measure their opinion about the extent of pesticide (defined as insecticide and ac-aricide) resistance in Florida, and its current and potential impact. We polled Univer-

388



sity and USDA entomologists from various backgrounds, representing the fields ofagricultural, ornamental and landscape, medical and veterinary, or household andstructural pest management. Entomologists with considerable field experience, and aclose relationship with producers or pest control professionals, were favored. We re-ceived responses from 14 of those surveyed, and 12 respondents provided useful infor-mation. Additional information was sought from other knowledgeable individuals toround out the survey. The questions and responses were as follows:

The Current and Future Status of Pesticide Resistance

Respondents were asked to indicate if resistance was: not a problem, a minor prob-lem, a significant problem, or a critical problem. Only a single response was re-quested. The time frame for future problems was specified as 10 years in the future.

The respondents differed in their assessment of the severity of the resistance prob-lem depending on the crop or environment being considered. Resistance was viewedto be a critical problem in greenhouses (foliage plants, flowering plants, and somewoody ornamentals), floriculture (both greenhouse and field-grown flowers), and ani-mal production (penned and free-ranging). Ornamental plants have long been consid-ered to be extremely sensitive to damage, hence they are treated frequently and proneto insecticide resistance problems. Resistance in animal production is a more recentphenomenon, however, apparently resulting from widespread use of insecticide im-pregnated ear tags.

Resistance was considered to be a significant, but not critical, problem in vegetablecrops, some field crops, and households. This might be viewed as surprising, becausemany vegetable crops, some field crops, and households in Florida receive insecticidetreatments at frequencies similar to the aforementioned situations where pesticideresistance was judged to be critical. It is likely that the severity of the problem is dueas much to corporate marketing strategies as to pesticide use patterns. Specifically,the pesticide market is smaller for greenhouse, floriculture and animal uses, so pes-ticide companies support fewer registrations. Therefore, when pesticide failures oc-cur, there are few options, or in some cases none. This, of course, results in a criticalsituation.

For medical pests, which in Florida is principally mosquitoes, the significance ofthe resistance problem apparently is related to location. Resistance was reported to bea significant problem in coastal locations, but only a minor problem in other areas.Coastal regions not only are extremely favorable for mosquito breeding, but a highproportion of the state’s population (79%) dwells along the coast, so there is frequentneed for chemical suppression.

Landscape plants seem to be relatively free of resistance problems. Woody orna-mentals are not usually planted in large single-species stands, which may help themto avoid development of high pest populations. Such landscape plants often tolerateconsiderable defoliation or pest density without obvious symptoms, so chemical treat-ment is not a regular feature of landscape maintenance. Also, in recent years therehas been a concerted effort to introduce native, hardy, pest-resistant plants into thelandscape, reducing the need for insecticide treatment. Among landscape plants, per-haps only turfgrass is treated regularly, and the southern chinch bug,

Blissus insu-laris

Barber, exhibits some degree of resistance, particularly in southern Florida.Nursery production of landscape ornamentals is also an exception, and mites canpresent resistance problems in this environment.

Although the number of pests displaying resistance to pesticides has increasedmarkedly in the last two decades, respondents generally did not see the resistanceproblem worsening greatly in the next 10 years. The only exception was the area of


389

household pest management, where the situation is anticipated to become critical.This generally optimistic attitude likely reflects faith in the agrichemical industry,which has continued to introduce novel pesticide chemistry or biorational materialsthat allow producers to continue with traditional agriculture and pest control prac-tices despite increasing numbers of pests that have become somewhat resistant to oneor more pesticides. The scientific community has also responded quickly and effec-tively to the onset of resistance by identifying alternative pest control chemicals andby helping to integrate other types of pest suppression into traditional production sys-tems.

The Extent of Resistance

Respondents were asked to indicate whether resistance applied to: a few com-pounds, numerous compounds, a few pests, or numerous pests. Up to two responseswere possible. Respondents were also asked to designate how many pests or com-pounds were affected and to indicate either a specific number or range.

The extent of the resistance problem was reported to be variable, depending onwhether the focus was the number of pests or pesticides. Resistance was generally re-ported to be limited to few pests in each commodity or environment. The number of re-sistant species was generally reported to be 3-5 per respondent, with a range given as1-10 per respondent. Although the number of species was small, the number of chem-ical compounds to which the pests were reported resistant was considerably larger.Respondents generally indicated that pests exhibiting resistance were resistant to5-10 compounds. The range in the number of compounds was given as from 1 productto all those on the market.

The Context of Pesticide Resistance

Respondents were asked to indicate if particular pests, crops, or environments ex-isted in which resistance occurred more frequently.

Respondents most frequently indicated that high value, damage-sensitive cropswere prone to have pesticide resistance problems. They cited greenhouse, floricul-tural, and vegetable crops as examples.

The environments next most frequently cited as having resistance problems werethose in which frequent or routine pesticide applications were made. Of course thiscorresponds to the aforementioned high value crops, but there are also situations inwhich value and damage sensitivity are not a major issue; examples are households,livestock, and certain field crops.

Only infrequently were the biological characteristics of the pests cited as favoringfrequent occurrence of resistance. Pests with short generation times and high intrin-sic rates of increase were suggested to be more prone to display resistance.

The Pattern of Pesticide Resistance

Respondents were asked if there were any patterns evident wherein entire classesof pesticide compounds or groups of arthropods displayed a tendency toward in-creased frequency of resistance, or whether resistance applied only to specific materi-als or pests.

Patterns of pesticide resistance related to chemical or biological taxon were not es-pecially evident to our respondents. Many said that pesticide resistance wasspecies-specific, that biological taxon was not a very good predictor of resistance prob-lems. A few, however, suggested that whiteflies, thrips, and especially mites were re-

390



sistance prone. Similarly, although cross resistance within chemical classes wasacknowledged, respondents indicated that they generally considered each pesticide tohave unique chemical properties, so that development of resistance was difficult topredict based on chemical taxon. The exception to this generalization seems to be thepyrethroids, where there is general acknowledgment that resistance is likely to de-velop.

Instances of Pesticide Resistance

Respondents were asked to name specific instances of pesticide resistance, includ-ing the pesticide, pest, and approximate date, and also to indicate whether the infor-mation on resistance was “documented” or anecdotal.

Instances of pesticide resistance in Florida provided by respondents are shown inTable 1. Surely this is not a complete list, either of pests or problem pesticides, butserves to demonstrate adequately the diversity of arthropod taxa affected. Also, ar-thropods found in numerous environments or crop systems are affected, and some his-torical trends are evident. Respondents acknowledged that only about one-half of thepurported cases of resistance are “documented,” with the remainder based on anec-dotal information. However, we carefully selected experienced entomologists andasked them to respond only in their area of expertise. Thus, we are confident that in-stances of misapplication and other potential sources of erroneous reports of resis-tance are not included. Because some of the “documented” resistance is from industrysources and not accessible to us, we have not included this specific information. Notealso that this table does not include information on the leafminer

L. trifolii

and the di-amondback moth, two insects with well-documented histories of insecticide resistancein Florida. A review of insecticide resistance in these two troublesome insects follows.

I

NSECTICIDE

R

ESISTANCE

IN

L.

TRIFOLII

Past and Present Situation

Prior to 1945, leafminer problems on celery and other vegetables in Florida wereapparently almost nonexistent. Control consisted mainly of clean-up measures andapplication of nicotine sulphate (Wolfenbarger 1947). Wolfenbarger (1947) recom-mended chlordane for control of leafminer on potatoes in south Florida. Harris (1962)reported that dimethoate, which was not labeled for celery, and diazinon and naledwhich were labeled, could control leafminer on celery in 1962. Genung et al. (1979) re-ported that with the use of diazinon, naled, and azinphos-methyl, the mortality of veg-etable seedlings and yield reductions declined and leafminer populations remainedlow until 1974, when they began to heavily infest celery and tomato. Genung et al.(1979) reported that in 1974 growers could not control leafminers on celery with diaz-inon, naled, or azinphos-methyl and that dimethoate, which was approved for use oncelery the same year, also did not give the desired level of control. Poe & Strandberg(1979) reported that oxamyl, which was approved for use on celery in 1975, was effec-tive for about two years. They also reported that in 1976 and 1977 leafminer on celerywas uncontrollable in Florida by any insecticide labeled for use. Florida growers ac-quired the use of methamidophos in 1977 and permethrin in 1978 for the control ofleafminer on celery. Permethrin became ineffective for leafminer control on celery inless than two years. Methamidophos was then considered the only insecticide thatgave any amount of control in celery in Florida, and it was considered marginally ef-fective. The possibility of effective chemical control did not come until the spring of


391

T

ABLE

1. E

XAMPLES

OF

I

NSECTICIDE

AND

A

CARICIDE

R

ESISTANCE

IN

F

LORIDA

C

ITED

BY

R

ESPONDENTS

IN

A

1994 S

URVEY

OF

P

UBLIC

S

ECTOR

E

NTOMOLOGISTS

.

ArthropodDate

(decade) Pesticide

House fly 1940 DDT

Musca domestica

(L.) 1950 chlordane, dieldrin, lindane, malathion1970 dimethoate, ronnel, tetrachlorvinphos1980 cyromazine, methomyl, various pyrethroids

German cockroach 1950 chlordane, dieldrin, lindane

Blatella germanica

(L.) 1960 allethrin, diazinon, malathion1970 carbaryl, propoxur1980 cyfluthrin, cypermethrin

Cat flea 1950-70 diazinon, malathion

Ctenocephalides felis

(Bouché)1970-801980-90

bendiocarb, carbaryl, propoxurcyfluthrin, fenvalerate, permethrin

Horn fly

Haematobia irritans

(L.)

1980 fenthion, fenvalerate, flucythrinate, permethrin, stirophos

Salt marsh mosquito 1950 DDT

Culex nigripalpus

1960 malathionTheobald 1990 methoprene

Soybean looper

Pseudoplusia includens

(Walker)

1970 acephate, methomyl, various pyrethroids

Fall armyworm

Spodoptera frugiperda

(J. E. Smith)

1970-80 malathion, carbaryl, methyl parathion, diazinon, trichlorfon, fluvalinate, bifenthrin, tralomethrin

Southern green stinkbug 1970 carbaryl, methomyl

Nezara viridula

(L.) 1980 endosulfanTobacco budworm

Heliothis virescens

(Fabricius)

1970 ethyl parathion

Corn earworm 1950-60 malathion, diazinon,

Helicoverpa zea

(L.) 1960-70 ethyl parathion, carbarylPepper weevil

Anthonomus eugenii

Cano

1990 fenvalerate, permethrin, oxamyl

Beet armyworm

Spodoptera exigua

(Hübner)

1980 chlorpyrifos, methomyl

Tomato pinworm 1970 carbaryl

Keiferia

1980 methomyl, fenvalerate

lycopersicella

(Walsingham)1990 oxamyl

Western flower thrips

Frankliniellaoccidentalis

(Pergande)

1980 pyrethroids

Mole crickets

Scapteriscus

spp.1970 chlordane

392



1982 when the celery industry secured the use of cyromazine. With the use of cyro-mazine, leafminer problems were considered under control until late 1989 when anunusual lack of efficacy occurred in the Everglades area.

Laboratory studies confirmed the presence of a high level of cyromazine resistancein a suspect strain of

L. trifolii

(G. L. L., unpublished data). Larval mortality in theresistant strain at 300 ppm of cyromazine, the highest label concentration used in thefield, was low enough to explain the loss of efficacy. The cyromazine resistance was ex-pressed as an incompletely recessive trait and not sex-linked. Backcrossing suggestedthat the resistance was conferred by a major gene. The resistance was considered un-stable since sensitivity returned in the resistant strain (from an LC

50

of about 440ppm to an LC

50

of about 85 ppm) within 5 generations of laboratory rearing withoutselection. This was consistent with a survey of leafminer populations that indicatedsusceptibility to cyromazine had returned during the summer of 1990 (J. S. Ferguson,unpublished data). This reversion was probably due to the immigration of susceptibleindividuals during what is traditionally a period of little or no celery production andvery little use of cyromazine.

The cyromazine-resistant strain was not resistant to abamectin (G. L. L., unpub-lished data), the only logical alternative insecticide available for control of leafminerin celery. This information contributed to the granting of a crisis exemption (Section18, FIFRA) in early 1990 for the use of abamectin in celery to control leafminer. Fur-ther efforts of the Florida Fruit and Vegetable Association, celery growers, CIBA,Merck Research Laboratories, and the University of Florida resulted in the subse-quent granting by the EPA (Section 18, FIFRA) in October 1990 of a specific exemp-

Green peach aphid

Myzus persicae

(Sulzer)1970-80 malathion, diazinon, oxydemeton-methyl,

dimethoateCabbage looper 1950-60 DDT, toxaphene, parathion

Trichoplusia ni

1960-70 endrin, mevinphos, naled(Hübner) 1970-80 methomyl

Cowpea curculio

Chalcodermus aeneus

Boheman

1980 endosulfan

Citrus rust mite

Phyllocoptruta oleivora

(Ashmead)

1990 dicofol

Yellow pecan aphids

Monellia caryella

(Fitch),

Monelliopispecanis

Bissell

1980 various pyrethroids

Silverleaf whitefly

Bemisia argentifolii

Bellows & Perring

1990 bifenthrin, fenvalerate, permethrin, endosulfan

Two-spotted spider mite 1980 fenbutatin-oxide

Tetranychus urticae

Koch1990 avermectin

Melon aphid

Aphis gossypii

Glover1990 acephate

TABLE 1. (CONT.) EXAMPLES OF INSECTICIDE AND ACARICIDE RESISTANCE IN FLORIDACITED BY RESPONDENTS IN A 1994 SURVEY OF PUBLIC SECTOR ENTOMOLOGISTS.

ArthropodDate

(decade) Pesticide

Leibee and Capinera: Symposium on Pesticide Resistance 393

tion for the use of abamectin in celery. Since then, abamectin has been used in celeryunder specific exemptions. These specific exemptions are unique in that, in order todiscourage the onset of resistance to abamectin, only two consecutive applications areallowed, forcing rotation with another insecticide. However, no other effective insec-ticide was available for rotation except for cyromazine which, due to reversion, hadbecome efficacious again. Since cyromazine and abamectin have different modes of ac-tion and no cross resistance was indicated, cyromazine was included in the leafminercontrol program under well-defined resistance management guidelines.

Management of Cyromazine Resistance in L. trifolii

A program for managing cyromazine resistance in L. trifolii was presented to cel-ery growers. The goal of this program was to control L. trifolii while increasing andpreserving susceptibility to cyromazine and minimizing the possibility of selecting forresistance to abamectin. Cyromazine use patterns and celery culture were suggestedthat would reduce selection of resistant phenotypes and encourage the immigration offeral, hopefully susceptible, leafminers into resistant populations.

Recommendations included: using noninfested transplants; initiating the sprayprogram based on a threshold to reduce the number of insecticide applications; start-ing with abamectin to maximize early control; rotating two sprays of abamectin withtwo sprays of cyromazine to avoid excessive use of one insecticide; finishing a plantingwith two applications of abamectin to reduce the number of adults emerging from thesoil and the trash after harvest; disking in trash as soon as possible to remove thissource of leafminers; and not using pyrethroids, such as permethrin and esfenvaler-ate, to minimize adverse effects on parasites and predators.

In addition, since acreage is very low in the production fields as harvesting ends(June) and the transplanting begins (September), it was recommended to not use cy-romazine during the summer (June through September) to prevent the continued se-lection of isolated populations and to encourage the immigration of susceptibleindividuals when celery acreage is at its lowest. Except for seedling production, Julyand August are otherwise free of celery. Not using cyromazine at all in seedling bedswas recommended, since transplanting from infested seedling beds is considered animportant mechanism for transferring resistant leafminers to the production fields.Lastly, seedlings were recommended to be grown distant from the field production ar-eas to reduce the chances of infestation by resistant leafminers.

INSECTICIDE RESISTANCE IN THE DIAMONDBACK MOTH

Past and Present Situation

Historically, the diamondback moth was considered a minor pest, usually includedin a complex of cabbage caterpillars along with the cabbage looper and the importedcabbageworm, Artogeia rapae (L.), but of much less importance. Control recommen-dations for the diamondback moth generally have been the same as for the other cab-bage caterpillars (Sanderson 1921, Metcalf & Flint 1939, Watson & Tissot 1942,Metcalf et al. 1951, 1962). Prior to the mid-1940s, insecticides used for cabbage cater-pillar control included nicotine, arsenicals, pyrethrum, rotenone, kerosene, and hotwater (150°F), and from the mid-1940s through the 1970s included DDT, toxaphene,parathion, methoxychlor, mevinphos, endosulfan, naled, methomyl, and methami-dophos. Bacillus thuringiensis was also available, but was not used extensively due toexpense and the perception of less than desirable control. In the early 1980s growers

394 Florida Entomologist 78(3) September, 1995

switched to the newly available and extremely effective pyrethroids, permethrin andfenvalerate for control of the cabbage looper and diamondback moth, both of whichhad become difficult to control with the other insecticides.

Insecticide resistance had long been suspected as the cause of the poor cabbagelooper control (Genung 1957, Workman & Greene 1970), and Shelton & Soderlund(1983) showed that a population from Florida was one of the most resistant to meth-omyl in the eastern U.S. The poor control of diamondback moth has been attributedto the destruction of parasites by excessive use of insecticides, such as methomyl,which were applied for cabbage looper suppression, but which were relatively ineffec-tive on diamondback moth. However, the poor control of diamondback moth may haveactually been the earliest indications of resistance problems.

Permethrin and fenvalerate proved to be very effective for control of all cabbage in-sects until the mid-1980s when growers observed that these insecticides were nolonger providing effective control of the diamondback moth. University trials reflectedthe same lack of control with fenvalerate (Leibee 1986) and from the winter of 1986-87 to the present, pyrethroid insecticides provided poor control of diamondback mothat Sanford, FL (G. L. L., unpublished data). Magaro & Edelson (1990) noted that fail-ures to control diamondback moth in south Texas were first reported by cabbage pro-ducers in the spring of 1987. Leibee & Savage (1992b) reported a high level ofresistance to fenvalerate in a laboratory strain of diamondback moth collected in cen-tral Florida in 1987.

Loss of efficacy with pyrethroids for control of diamondback moth caused growersto switch to intensive use of several organophosphates, endosulfan, and B. thuring-iensis subspecies kurstaki (Btk), all of which did not provide the level of control pro-vided by pyrethroids prior to resistance. At present, many diamondback mothpopulations have become very difficult to control with any of the registered syntheticinsecticides and Btk.

The presence of Btk resistance in Florida was immediately suspected because Btkresistance in diamondback moth had been reported in Hawaii (Tabashnik et al. 1990),Japan (Tanaka & Kimura 1991), and Malaysia (Syed 1992); it was eventually con-firmed for Florida (Leibee & Savage 1992a, Shelton et al. 1993). With the presence ofBtk-resistance, there were essentially no effective insecticides available for control ofmany diamondback moth populations in Florida until the recent introduction of B.thuringiensis subspecies aizawai (Bta)-based insecticides. Bta-based insecticides(those possessing the Cry1C toxin) are being successfully used in areas where Btk-based insecticides have failed. Lack of resistance to Bta in diamondback moth resis-tant to Btk has been documented in Japan (Hama et al. 1992), Malaysia (Syed 1992),and Florida (Leibee & Savage 1992a, Shelton et al. 1993).

Diamondback moth abundance has been considered low for several seasons in cen-tral Florida (G. L. L., personal observation). This is due in part to the return of sub-stantial amounts of natural control from parasites, which in turn is attributed toreduced pyrethroid use. Growers are not spraying as frequently for diamondbackmoth and are able to use Btk-based insecticides, suggesting a return of susceptibilityto Btk.

Management of Resistance in the Diamondback Moth

Crop culture and control recommendations were made that would reduce the se-lection of resistant phenotypes of diamondback moth and encourage the immigrationof feral, hopefully susceptible, individuals into resistant populations. These recom-mendations were based on the following knowledge. Susceptibility to Btk had beengreatly reduced in some populations. Bacillus thuringiensis resistance in diamond-


back moth in Hawaii was shown to be inherited as a recessive trait (Tabashnik et al.1992) and observations from field and laboratory studies in Florida suggested thesame (G. L. L., unpublished data). Bacillus thuringiensis subspecies aizawai-basedproducts (those possessing the Cry1C toxin) appeared to be effective in populationswhere Btk susceptibility was reduced. Tank-mixing Bt with mevinphos was shown tobe quite effective at reducing infestations in early season (G. L. L. unpublished data);however, the use of mevinphos was to be discontinued in 1995, eliminating the mosteffective insecticide other than Bt for diamondback moth control on cabbage in Flor-ida. Use of pyrethroids and carbamates can select for resistance that might further re-duce the efficacy of organophosphate insecticides and endosulfan, and also destroy theparasites and predators providing natural control of the diamondback moth.

Crop culture recommendations included: not growing cabbage in the warmestmonths (May through September in central Florida) when insect pressure is the high-est and Bt-based insecticides are the least efficacious; immediately disposing of cropresidues to prevent migration from heavily selected populations into new plantingsand seedling production areas; and using noninfested transplants, which not onlycontributes to control but also reduces spread of diamondback moths to new locations.Diamondback moths that infest purchased transplants may be highly resistant due toheavy usage of insecticides on the transplants or in fields near transplant productionareas. Producers growing their own transplants are at an advantage because theyhave more control over infestation levels and have specific knowledge about the resis-tance problems in their production areas.

Control recommendations included: inspecting crops frequently (about twice perwk) to determine the presence of the pest of concern or unexpected pests; beginninginspections at the seedling stage because reducing infestations in early season ap-pears to be critical to managing diamondback moth; minimizing insecticide applica-tions whenever possible by using action thresholds developed through research or byintuition; and using pheromone traps to monitor the presence or absence of diamond-back moth before and during the growing season, and also for monitoring peaks ofadult activity (Baker et al. 1982) for timing insecticide applications.

Specific insecticide recommendations included: using Btk and Bta as the principleinsecticides for control of diamondback moth; if the population was known to be sus-ceptible to Btk, alternating a Bta-based product with a Btk-based product to avoid re-petitive applications of the same insecticide to reduce the selection of resistance toany one product; using only Bta to insure maximum control if Btk resistance wasknown to be present or the status of Btk susceptibility was unknown; applying Bttwice weekly and tank-mixing with mevinphos weekly, starting with the tank-mix tomaximize the control that is critical early in crop; including endosulfan, chlorpyrifos,and methamidophos as alternatives or substitutes for mevinphos in the tank-mixeswith Bt, especially endosulfan, since it belongs to a different chemical class thanmevinphos; avoiding the use of carbamates; and, not using pyrethroids.

OBSERVATIONS ON INSECTICIDE RESISTANCE IN L. TRIFOLII AND DIAMONDBACK MOTH

Probably the greatest factor contributing to the development of insecticide resis-tance in L. trifolii and diamondback moth was long term and frequent use of single in-secticides or classes of insecticides. However, celery and cabbage cultural practices inFlorida probably contribute to the rapidity and degree with which insecticide resis-tance develops in these two insects. Both crops are grown in relatively small and iso-lated areas, or pockets, of agricultural activity. In addition, both crops are grown insome form year round. This isolation and the lack of a substantial crop-free period re-sult in the containment and “cycling” of resistant populations. This results in the


same population being exposed to insecticides continually. In addition, the constantuse of insecticides removes susceptible individuals that may be immigrating into thepopulation, thus preventing the opportunity for reversion.

These factors are believed to be especially evident with the development of highlevels of resistance in the diamondback moth in the 1980s. Several changes in the pro-duction of cabbage in the 1980s contributed greatly to the “cycling” of resistant popu-lations in the production areas and the movement of resistant populations betweenproduction areas within and outside Florida. Among these changes was the lengthen-ing of the crucifer production period by harvesting later in the spring and transplant-ing earlier in the summer. Prior to this situation, much of the summer(June-September) in central Florida was basically a crucifer production-free periodand diamondback moth populations were very low. Crucifer production was thrustinto the warmer, drier parts of the year when diamondback moth became more of aproblem, resulting in increased use of insecticides and subsequent exposure of addi-tional generations to more selection pressure. With development of the containergrown transplant industry and the ability to grow transplants in the summer, a situ-ation was eventually created in which crucifers were continually produced through-out the year in the same localities, either as transplants or field crops, or both.Insecticide resistant diamondback moth could move from the fields to transplants inthe summer, and be redistributed back to the fields in late summer and early fall.Therefore, heavy populations of resistant diamondback moth were being perpetuatedlocally throughout the year and continually exposed to insecticides. Opportunities forthe return of susceptibility by the immigration of individuals with susceptible pheno-types was essentially eliminated.

Transplants probably were a major factor in the development of resistance prob-lems in diamondback moth on a national level during the 1980s; container-growntransplants were popular and a healthy containerized transplant industry developedin the south. Much of the transplant production in the south in the winter and springsupplied more northern growers with transplants to establish stands in the springwhen environmental conditions were not conducive to direct seeding. Transplantswere also produced during summer months in the north to facilitate establishment ofstands in the south when soil temperatures are too high for germination.

Transplant growers likely were oblivious to the fact that they were shipping resis-tant diamondback moth. They were probably controlling the later larval stages thatcause obvious damage and shipping what appeared to be uninfested plants. Adultsflying into the open-sided greenhouses from the field could maintain a supply of eggsand early mining instars before shipment. In addition, larvae have been observed tobe deep inside the bud of the transplant, out of reach of the insecticide and the eye ofthe grower. It is possible that avoidance of the insecticide deposits drives diamond-back moth larvae emerging from the leafmine into the bud.

Another aspect of the transplant industry that could have contributed to the de-velopment of resistance was that only a few large growers in areas of high levels of re-sistance produced most of the transplants used in Florida and the rest of the U.S. Thisgreatly increased the probability of many growers receiving transplants from areaswith resistant diamondback moth populations.

CONCLUSION

The results of the survey attest to the impact of insecticide resistance on past andpresent pest management in Florida. The review of resistance problems with L. tri-folii and diamondback moth illustrates how insecticide resistance can complicate themanagement of pestiferous arthropods. Insecticide resistance seems to be pervasive


in Florida, suggesting that we are not adequately considering the consequences of theway we use pesticides. Resistance management strategies should be integrated withnonchemical control whenever possible. In the case of the aforementioned L. trifoliiand diamondback moth problems, crop management (isolation and continuous crop-ping) were key factors in development of resistance.

In most cases, pesticides are the most efficient, easiest, and cheapest methods ofcontrolling pestiferous insects and mites. The number of pesticides available is dwin-dling rapidly, however, due to cancellation of registrations and lack of re-registration.As a result, the conservation of arthropod susceptibility to the remaining pesticides,and to newly developed pesticides, is becoming extremely important. Unfortunately,we have not made adequate effort to conserve susceptibility of arthropods to pesti-cides. The minimum effort should include development of baseline data that would al-low the investigation of potential resistance episodes in a timely manner. Conservingpesticide susceptibility takes the development of knowledge and a commitment fromthose responsible for producing, using, and conducting research on pesticides. Thus,educational efforts should be enhanced.

Something everyone can do to help manage resistance to any pesticide is to refineits use. Application of the correct amount and type of insecticide in a timely and effi-cient manner would help forestall the onset of resistance, particularly if nonchemicaltechniques could be used to reduce the numbers and frequency of application. This isbest accomplished following research on, and implementation of, IPM strategies. In-creasingly, pesticide resistance management must be considered an important com-ponent of IPM.

ACKNOWLEDGMENTS

We gratefully acknowledge the assistance of the following individuals who gavegenerously of their time to make the survey component of this paper possible: JoeFunderburk, Jerry Hogsette, Freddie Johnson, Joe Knapp, Phil Koehler, Russ Mizell,Charlie Morris, Lance Osborne, Jim Price, Dakshina Seal, Dave Schuster, PhilStansly, and Simon Yu. We are also grateful to Brett Highland for his critical reviewof the manuscript. Florida Agricultural Experiment Stations Journal Series No. R-04551.

REFERENCES CITED

BAILEY, D. L., G. C. LABRECQUE, AND T. A. WHITFIELD. 1970. Resistance of house flies(Diptera: Muscidae) to dimethoate and Ronnell in Florida. Florida Entomol.53:1-5.

BAKER, P. B., A. M. SHELTON, AND J. T. ANDALORO. 1982. Monitoring of diamondbackmoth (Lepidoptera: Yponomeutidae) in cabbage with pheromones. J. Econ. En-tomol. 75:1025-1028.

BLOOMCAMP, C. L., R. S. PATTERSON, AND P. G. KOEHLER. 1987. Cyromazine resis-tance in the house fly (Diptera: Muscidae). J. Econ. Entomol. 80:352-357.

BROGDON, J. E. 1957. Symposium—vegetable insect resistance to insecticides in Flor-ida. Proc. Florida State Hort. Soc. 70:143-144.

EL-GAZZAR, L. M., J. MILIO, P. G. KOEHLER, AND R. S. PATTERSON. 1986. Insecticideresistance in the cat flea (Siphonaptera: Pulicidae). J. Econ. Entomol. 79:132-134.

GENUNG, W. G. 1957. Some possible cases of insect resistance to insecticides in Flor-ida. Proc. Florida State Hort. Soc. 70:148-152.

GENUNG, W. G., S. L. POE, AND C. A. MUSGRAVE. 1979. Insect and mite pests of celery,pp. 29-52 in S. L. Poe and J. O. Strandberg [eds.], Opportunities for integrated


pest management in celery production. University of Florida, Institute of Food& Agricultural Sciences. UF-IFAS-IPM 2. 104 pp.

HAMA, H., K. SUZUKI, AND H. TANAKA. 1992. Inheritance and stability of resistance toBacillus thuringiensis formulations of the diamondback moth, Plutella xylos-tella (Linnaeus) (Lepidoptera: Yponomeutidae). Appl. Entomol. Zool.27:355-362.

HARRIS, E. D., JR. 1962. Insecticides and intervals between applications for leafminercontrol on celery. Proc. Florida State Hort. Soc. 75:184-189.

HOSKINS, W. M., AND H. T. GORDON. 1956. Arthropod resistance to chemicals. Annu.Rev. Entomol. 1:89-122.

HOSTETLER, M. E., AND R. J. BRENNER. 1994. Behavioral and physiological resistanceto insecticides in the German cockroach (Dictyoptera: Blatteridae): an experi-mental reevaluation. J. Econ. Entomol. 87:885-893.

KEIL, C. B., AND M. P. PARELLA. 1990. Characterization of insecticide resistance intwo colonies of Liriomyza trifolii (Diptera: Agromyzidae). J. Econ. Entomol.83:18-26.

KOEHLER, P. G. 1991. Toxicity of hydramethylnon to laboratory and field strains ofGerman cockroach (Orthoptera: Blattellidae). Florida Entomol. 74:345-349.

LEIBEE, G. L. 1986. Caterpillar control on cabbage, spring 1985. CFREC-Sanford Re-search Report SAN 86-10. Univ. Florida IFAS. 2 p.

LEIBEE, G. L., AND K. E. SAVAGE. 1992a. Observations on insecticide resistance in di-amondback moth, pp. 41-46 in Seminar Proceedings: Global Management ofInsecticide Resistance In The 90s. Abbott Laboratories. Sept. 15-17, 1992. LakeBluff, IL.

LEIBEE, G. L., AND K. E. SAVAGE. 1992b. Toxicity of selected insecticides to two labo-ratory strains of insecticide-resistant diamondback moth (Lepidoptera: Plutel-lidae) from central Florida. J. Econ. Entomol. 85:2073-2076.

MAGARO, J. J., AND J. V. EDELSON. 1990. Diamondback moth (Lepidoptera: Plutel-lidae) in south Texas: a technique for resistance monitoring in the field. J. Econ.Entomol. 83:1201-1206.

METCALF, C. L., AND W. P. FLINT. 1939. Destructive and useful insects. McGraw-HillBook Co., New York.

METCALF, C. L., W. P. FLINT, AND R. L. METCALF. 1951. Destructive and useful in-sects. McGraw-Hill Book Co., New York.

METCALF, C. L., W. P. FLINT, AND R. L. METCALF. 1962. Destructive and useful in-sects. McGraw-Hill Book Co., New York.

MILIO, J. F., P. G. KOEHLER, AND R. S. PATTERSON. 1987. Evaluation of three methodsfor detecting chlorpyrifos resistance in german cockroach (Orthoptera: Blattel-lidae) populations. J. Econ. Entomol. 80:44-46.

OMOTO, C., T. J. DENNEHY, C. W. MCCOY, S. E. CRANE, AND J. W. LONG. 1994. Detec-tion and characterization of the interpopulation variation of citrus rust mite(Acari: Eriophyidae) resistance to dicofol in Florida Citrus. J. Econ. Entomol.87:566-572.

PITRE, H. N. 1988. Relationship of fall armyworm (Lepidoptera: Noctuidae) from Flor-ida, Honduras, Jamaica, and Mississippi: susceptibility to insecticides with ref-erence to migration. Florida Entomol. 71:56-61.

POE, S. L., AND J. O. STRANDBERG. 1979. Crop protection through prevention andmanagement, pp 1-4 in S. L. Poe and J. O. Strandberg [eds.], Plant protectionthrough integrated pest management. Opportunities for integrated pest man-agement in celery production. University of Florida, Institute of Food & Agri-cultural Sciences. UF-IFAS-IPM 2. 104 pp.

SANDERSON, E. D. 1921. Insect pests of farm, garden, and orchard. John Wiley andSons, Inc., London.

SHELTON, A. M., AND D. M. SODERLUND. 1983. Varying susceptibility to methomyl andpermethrin in widely separated cabbage looper (Lepidoptera: Noctuidae) pop-ulations within eastern North America. J. Econ. Entomol. 76:987-989.


SHELTON, A. M., J. L. ROBERTSON, J. D. TANG, C. PEREZ, S. D. EIGENBRODE, H. K.PREISLER, W. T. WILSEY, AND R. J. COOLEY. 1993. Resistance of diamondbackmoth (Lepidoptera: Plutellidae) to Bacillus thuringiensis subspecies in thefield. J. Econ. Entomol. 86:697-705.

SYED, A. R. 1992. Insecticide resistance in diamondback moth in Malaysia, pp437-442 in N. S. Talekar [ed.], Proceedings of the second international work-shop on the management of diamondback moth and other crucifer pests. AsianVegetable Research and Development Center, Tainan, Taiwan.

TABASHNIK, B. E., N. L. CUSHING, N. FINSON, AND M. W. JOHNSON. 1990. Field devel-opment of resistance to Bacillus thuringiensis in diamondback moth (Lepi-doptera: Plutellidae). J. Econ. Entomol. 83:1671-1676.

TABASHNIK, B. E., J. W. SCHWARTZ, N. FINSON, AND M. W. JOHNSON. 1992. Inherit-ance of resistance to Bacillus thuringiensis in diamondback moth (Lepidoptera:Plutellidae). J. Econ. Entomol. 85:1046-1055.

TANAKA, H., AND Y. KIMURA. 1991. Resistance to Bt formulation in diamondbackmoth, Plutella xylostella L., on watercress. Japan J. Appl. Entomol. Zool.35:253-255.

WATSON, J. R., AND A. N. TISSOT. 1942. Insects and other pests of Florida vegetables.Univ. of Florida Agric. Exp. Stn. Bull. 370.

WOLFENBARGER, D. O. 1947. The serpentine leaf miner and its control. Univ. of Flor-ida Agric. Exp. Sta. Press Bul. 639.

WORKMAN, R. B., AND G. L. GREENE. 1970. Cabbage looper control at Hastings andSanford Florida during 1970. Proc. Florida State Hort. Soc. 83:166-167.

YU, S. J. 1992. Detection and biochemical characterization of insecticide resistance infall armyworm (Lepidoptera: Noctuidae). J. Econ. Entomol. 85:675-682.

YU, S. J., AND S. N. NGUYEN. 1992. Detection and biochemical characterization of in-secticide resistance in the diamondback moth. Pestic. Biochem. Physiol. 44:74-

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81.

Scott: Symposium on Pesticide Resistance

399

THE MOLECULAR GENETICS OF RESISTANCE:RESISTANCE AS A RESPONSE TO STRESS

J

ULIE

A. S

COTT

Knipling-Bushland USDA Livestock Insect Research LaboratoryAgricultural Research Service

Kerrville, TX 78028-9184

A

BSTRACT

In this overview of the molecular genetics of resistance, pesticides are regarded asone of the many environmental stresses against which insects must defend them-selves to survive. Examined at the genetic level, pesticide resistance appears to be apreadapted response to stress and not due to novel mutations caused by pesticide ex-posure. The genetic mutations—gene amplification, altered gene regulation, struc-tural alteration of a gene—which result in resistance are described and explained anda possible distribution mechanism of resistance genes is considered. Resistance mech-anisms, their associated biological processes and the types of genetic mutations asso-ciated with each are detailed. Finally, the potential of molecular technology for thedevelopment of novel methods to detect and monitor for resistance is examined andcompared to more traditional technology.

400



Key Words: Pesticide resistance, molecular, mutation, genetic, adaptation, stress, in-sect.

R

ESUMEN

En esta revisión de la genética molecular de la resistencia, los pesticidas son con-siderados como uno de los muchos estréses ambientales de los cuales los insectos de-ben defenderse para sobrevivir. Examinada a nivel genético, la resistencia a lospesticidas parece ser una respuesta preadaptada al estrés y no debida a nuevas mu-taciones causadas por la exposición a los pesticidas. Son descritas y explicadas las mu-taciones genéticas-amplificación genética, regulación génica alterada, alteraciónestructural de un gen-que producen la resistencia, y es considerado un posible meca-nismo de distribución de los genes de resistencia. Son detallados los mecanismos deresistencia, los procesos biológicos y los tipos de mutaciones asociados con cada uno deellos. Finalmente, es examinado y comparado a la tecnología más tradicional el poten-cial de la tecnología molecular para el desarrollo de nuevos métodos de deteccción y

monitoreo de la resistencia.

When we discuss pesticide resistance, we generally refer to our ability to control apest, not the pest’s ability to defend itself. Yet resistance is really a form of self-de-fense. To an insect, exposure to a pesticide is just one of the myriad of dangers whichmust be avoided in order to survive. In this sense, pesticide exposure may be de-scribed as an environmental stress and resistance as the overt expression of an in-sect’s natural response to that stress. Insects have been confronted with lethal andnonlethal stresses for as long as they have existed. And, they have evolved effectivedefense mechanisms to deal with these stresses. Their potential to adapt and developresistance to stress becomes most apparent when examined at the molecular geneticlevel. It is the objective of this paper to examine how and why insects adapt to stress,particularly pesticide exposure at the molecular genetic level and to emphasize thatresistance is a part of the normal response of insects to stress. In addition, the advan-tages and limitations of traditional and molecular technologies for monitoring for re-sistance are briefly compared. This paper is a general overview to introduce readersto concepts of molecular genetics and pesticide resistance. It is not a technical reviewof the molecular biology of specific resistance mechanisms.

S

TRESS

, R

ESISTANCE

AND

T

OLERANCE

: D

EFINITIONS

AND

I

NTERACTIONS

Before discussing the molecular genetics of resistance, some definition and discus-sion of the relationships of stress, resistance, and tolerance is required. Stress hasbeen broadly defined as “any environmental change that acts to reduce the fitness ofan organism” (Koehn & Bayne 1989). Stresses may have physical, biotic, and/or toxiccomponents (Fig. 1) which, in turn, may be acute, chronic, and/or seasonal and affectinsects at the community, population, and/or individual level. The deleterious effectsof excesses in temperature or humidity are self-evident. Exposure to ultraviolet radi-ation can influence trophic-level interactions of entire communities to the advantageof one population at the expense of another (Bothwell et al. 1994). Predation, parasit-ism, disease, inter- and intraspecies competition all act to determine the success orfailure of a population to occupy a particular ecological niche. Toxic components of theenvironment are also stresses that can affect populations and can be divided intothree groups: pollutants, which may be natural or artificial, pesticides, and plant al-


401

lelochemicals. Each of these stresses can affect insects differently, but all must be ad-equately dealt with for continued survival.

Resistance has been defined as the development of the ability in a strain of an or-ganism to tolerate doses of a toxicant which would prove lethal to the majority of in-dividuals in a normal (i.e. susceptible) population of the same species (Anonymous1957). This definition is somewhat imprecise, because it infers that resistance can de-velop in an individual either before or after exposure to a toxin, two very differentevents. Resistance is the phenotypic expression throughout a population of a herita-ble trait that was already expressed in at least some of the individuals in the popula-tion

prior

to exposure to a toxicant. The development of a measurable shift in apopulation’s susceptibility to a toxin is due to the specific selection of these pre-adapted individuals in the population, often over several generations, by exposure toamounts of toxicant which are sublethal to the pre-adapted individuals but may ormay not be sublethal to others in the population. For many years this adaptive eventwas difficult to understand; however, we now know that the toxic components of somepesticides are similar to ones present in plants (e.g. pyrethrum) and that the detoxi-fication systems that deal with these plant allelochemicals are the same systems thatdetoxify pesticides. Therefore insects which can detoxify certain plant allelochemicalswell are pre-adapted to detoxify and develop resistance to pesticides which have thesame mode of action as the allelochemicals even before the insects are ever exposedto the pesticides.

Resistance and tolerance are often used interchangeably in the literature and todefine one another; however, tolerance is also used to describe shifts in susceptibilitythat occur within a single generation

after

exposure to stress which is not expressedby succeeding generations until

after

exposure to a similar stress. This phenomenonis different from the one previously described. For example 6-day-old bollworms,

He-

Figure 1. Examples of environmental stresses which can act on living organismsand force individuals, populations, and entire communities to continuously adapt tonew and ever changing conditions in order to survive.

402



liothis zea

(Boddie), only detoxify relatively small amounts of methyl parathion, butat 12 days, they can detoxify 30-fold more methyl parathion. If gossypol, a toxic alle-lochemical present in cotton, is added to the diet, then 12-day-old larvae can detoxifynot 30-fold but 75-fold more methyl parathion. The presence of gossypol in the diet in-duced the larvae to produce more detoxifying enzymes (Muehleisen et al. 1989). Yet,the 6-day-old progeny of these individuals do not retain either the 30- or 75-fold in-crease in tolerance of their parents but must develop it over time and after exposureto gossypol. The ability to metabolize methyl parathion is inherent; the increase intolerance to it is not. The difference between resistance and tolerance can becomeblurred when a population is subjected to a strong selection pressure, such as chronicpesticide exposure, and the mechanism that is induced by the pesticide exposure toyield tolerance is also the mechanism that is specifically selected to yield resistance.Resistance, as it will be discussed here, refers to a decrease in susceptibility which isheritable and does not need to be induced before it can be expressed; however, expo-sure to a stress, such as an insecticide, may result in an increase in the expression ofthe resistance gene(s) which may or may not be heritable.

Two other terms which are often used and may be confused with one another arecross-resistance and multi-resistance. Cross-resistance is resistance to two or moreclasses of pesticides which occurs because the pesticides have the same, or very sim-ilar, modes of action. Organophosphate and carbamate pesticides intoxicate by simi-lar modes of action, and resistance to one usually results in resistance to the other.Multi-resistance refers to resistance to two or more classes of pesticides because of thecoexistence of two or more different resistance mechanisms. For example, a resistantinsect may have both metabolic resistance to organophosphates and target-site resis-tance to pyrethroids.

T

HE

W

AYS

AND

M

EANS

OF

A

LTERING

G

ENETIC

M

ATERIAL

It should be apparent that pre-exposure to plant allelochemicals can only help ex-plain those cases of resistance where the pesticide is rendered inactive by the samedetoxification mechanism. It does not explain resistance mechanisms which do notappear to be selected for by plant allelochemicals or seem to arise spontaneously or in-crease in amplitude after exposure to a pesticide. How can exposure of the parentalgeneration to stress result in their progeny being more resistant to that stress? Theanswer is that exposure to a stress causes the genetic material (i.e. the DNA) to be al-tered.

There appear to be three general types of alterations, i.e. mutations, that can occurand result in resistance (Fig. 2). A gene may be

amplified

so that instead of only hav-ing one copy of the gene, there are now many copies present in the DNA. If an insecthas ten copies of a gene, then it can make ten times as much product as an insect withonly one copy of the gene. If the amplified gene encodes for a detoxifying enzyme, thenthat insect can detoxify 10-fold as much toxicant as the insect with only one gene.

The expression of a gene may also be altered to yield resistance. In this case, thereis only one copy of the gene present in the mutated insect but that gene’s regulationis altered so that it produces more (or less) product compared to a susceptible individ-ual. For example, in a susceptible insect the gene to gene-product ratio may be 1:1 butin a resistant insect that ratio may be changed. The gene may be

up-regulated

to pro-duce more product, that is, the gene to gene-product ratio is now 1:10 or

down-regu-lated

to make less product (the ratio is now 1:0.1). When a pesticide is applied in itstoxic form, up-regulation of a detoxifying enzyme will increase resistance. When apro-insecticide, i.e. the material must be metabolized first in order to become toxic, isapplied, down-regulation of the metabolizing enzyme will increase resistance.


403

The third type of mutation that can result in resistance is a structural change ina gene which yields a corresponding structural change in its product. A single

pointmutation

, i.e. one nucleotide in the gene’s coding region is substituted with a differentnucleotide so that a different amino acid is encoded for at a specific position and thischange causes the gene product to have a different three-dimensional structure, canresult in resistance in several ways. It may decrease the ability of the insecticide tophysically bind to its site of action, or increase or decrease the gene product’s abilityto metabolize the insecticide. A structural change does not alter the quantity of theproduct made but alters the quality of the product made.

It is important to recognize that these alterations to an insect’s DNA do not createnew genes. They only affect pre-existing genes. The idea that exposure to a pesticidecauses resistance genes to be “created” has been debated periodically; however, nosubstantiating data have ever been proffered. It is much more likely that resistancegenes already exist in the pesticide-naive population at low frequency prior to selec-tion by a pesticide.

How resistance genes are spread throughout a population is not completely cer-tain. It is clear that pesticide exposure plays a key role. However, other factors mayalso be important in the spread of resistance genes. One hypothesis which has re-ceived considerable attention is that

transposable

or

mobile elements

play a signifi-cant role in some cases of gene amplification. Transposable or mobile elements (TEs)are discrete sections of DNA that can move to new chromosomal locations and prolif-erate at a higher frequency relative to other genomic sequences (i.e. more and morecopies of the TE are inserted into the genome) after they have moved. TEs can alsomove genes that were previously not mobile and whose functions are not related tothe transposition. The genes that are moved with the TEs are also replicated at ahigher frequency (Berg 1989). Gene amplifications may be initially distributedthroughout a population by TEs or a gene may be transposed to a new location whereits expression is altered to yield resistance. For example, it has been indirectly dem-onstrated in the laboratory that the transposition of

alleles

, alternative forms, of thegene

Met

results in insecticide resistance in

Drosophila

(Wilson & Turner 1992). But

Figure 2. Graphic representation of the types of genetic mutations which occurand cause resistance. (a) A gene is amplified to increase its number of copies in the ge-nome and consequently increase the amount of gene product made (b) the regulatoryexpression of a gene is altered to increase the amount of gene product made (note thatgene expression may also be altered to decrease the amount of product made) (c) thegenetic code is rewritten to produce a structurally different product.

404



so far there is absolutely no evidence that such an event has occurred in a field popu-lation.

R

ESISTANCE

M

ECHANISMS

AND

A

SSOCIATED

T

YPES

OF

M

UTATIONS

Interestingly, only specific types of genetic mutations appear to be associated withspecific resistance mechanisms (Table 1). The two types of mechanisms that causehigh levels of resistance are generally referred to as metabolic resistance and target-site insensitivity, respectively. Each of these consists of several biological mecha-nisms. Metabolic resistance can be divided into three principle enzyme systems: cyto-chrome P450 monooxygenases (P450s), nonspecific esterases, and glutathione S-transferases (GSTs). Components of each of these enzyme systems may be mutated toalter the detoxification of a pesticide.

Cytochrome P450s catalyze a variety of detoxification reactions in insects, includ-ing the hydroxylation of DDT, the epoxidation of cyclodienes, the aromatic hydroxyla-tion of the carbamates carbaryl and propoxur, and oxidation of phosphorothioates(Feyereisen et al. 1990). Given the variety of reactions stimulated by these enzymes,it is likely that several different P450 enzymes are present in any one insect and thatseveral alleles of each gene may exist. Such is the case for the mosquito

Anopheles al-bimanus

in which seventeen P450 genes were discovered (Scott et al. 1994) and forthe termites

Mastotermes darwiniensis

and

Coptotermes acinaciformis

in which mul-tiple isoenzymes of cytochrome P450s were detected biochemically (Haritos et al.1994). There is no evidence to suggest that P450 genes are amplified or structurallyaltered to yield insecticide resistance. But there are numerous examples of their ex-pression being altered by various substances (Rose et al. 1991, Jeong et al. 1992, Wax-man & Azaroff 1992, Snyder et al. 1993). By definition, if the expression of a P450 is

T

ABLE

1. T

HE

G

ENETIC

M

UTATIONS

A

SSOCIATED

WITH

E

NZYMES

AND

R

ECEPTORS

T

HAT

R

ESULT

IN

D

IFFERENT

T

YPES

OF

R

ESISTANCE

.

Associated Genetic Mutations

Types ofResistance

Gene Amplification

Altered Expression

Structural Change

Metabolic

P450 oxidases ND

1

+ NDEsterases + ND +GSTs ND + ?

Target site insensitivity

Acetylcholinesterase ND ND +GABA receptor ND ND +Sodium channel ND ? ?JH receptor ND ? ?

Other

Reduced penetration — — —Behavioral change — — —

1

ND = not detected; + = confirmed or strongly indicated; ? = implied but not confirmed; — = no data available.


405

altered to yield resistance, then susceptible and resistant strains will have quantita-tively different amounts of that P450. Three P450s have been demonstrated to beover-expressed by resistant insect strains: P450Lpr (Wheelock & Scott 1992),CYP6A1 (Cariño et al. 1994), and CYP6A2 (Waters et al. 1992), respectively. P450Lprhas been directly implicated as the major enzyme causing pyrethroid resistance inone strain of house fly (Wheelock & Scott 1992, Hatano & Scott 1993). CYP6A1 ap-pears to be a major cyclodiene-metabolizing enzyme in the house fly (Andersen et al.1994). The primary function of CYP6A2 has not been reported. A major handicap todetermining which P450s are involved in resistance is our inability to distinguish theactivities of individual uncharacterized P450s from each other with a high degree ofaccuracy. The activity of a specific P450 towards an insecticide must be determinedbecause its over-expression does not necessarily prove that it is the enzyme responsi-ble for resistance. The over-expression of CYP6A1 by the house fly is a case in point.CYP6A1 is over-expressed in a house fly strain that is resistant to DDT, organophos-phates, and carbamates but does not have significant resistance to cyclodienes. To de-termine which P450s are actually responsible for resistance, more specific substratesare needed.

Esterases are a large group of enzymes which metabolize a wide variety of sub-strates. All esterases are able to hydrolyze ester bonds in the presence of water. Sincemany insecticides, especially organophosphates and carbamates, contain ester bonds,it is not surprising that the mechanism of resistance in many cases is caused by ele-vated levels of esterases (Fournier et al. 1987, Field et al. 1988, Carlini et al. 1991,Kettermen et al. 1992, Chen & Sun 1994). Esterase levels can be elevated by eithergene amplification or altered gene expression. So far, no data have indicated that theexpression of esterase genes is altered to yield resistance, but the molecular charac-terization of esterases is limited to a very few insect species and this type of mutationcannot be discounted.

Considerable data show that certain non-specific esterase genes are amplified toyield resistance. The esterases which cause resistance in

Myzus persicae

Sulz. and

Culex

mosquitoes have been particularly well-studied. In these insects the resistantesterase genes are highly amplified and up to 250 copies of the same gene may befound in a single individual (Mouchès et al. 1986, Poirié et al. 1992). The more the es-terase genes are amplified, the greater the level of resistance that they provide (Fieldet al. 1988, Poirié et al.1992). This increase in resistance appears to be because the es-terases interact with the insecticides more readily than the insecticides’ own target.When the esterases are present in approximately an equal molar ratio to the insecti-cides, they are able to effectively sequester the insecticides and then slowly hydrolyzethe insecticides (Devonshire & Moore 1982, Ketterman et al. 1992, Karunaratne et al.1993).

How recently these esterases have been amplified in

Culex

populations has been amatter of some debate. One group has suggested that two esterases associated withresistance, A2 and B2, were amplified in a single event within the past forty years, i.e.since the use of organophosphate insecticides became widespread, and that A2 and B2have been distributed across three continents by migration since that event (Ray-mond et al. 1991). If this hypothesis is correct then all A2 and B2 genes should beidentical. But data show that they are not. Kinetic studies of the insecticidal interac-tion of purified A2 and B2 esterases found that different forms of each enzyme werepresent in a number of resistant strains (Ketterman at al. 1993), and three amino aciddifferences have been found between the two B2 genes that have been sequenced(Vaughan et al. 1995). Therefore, not all of the A2 and B2 genes are identical and ei-ther these genes were amplified in at least two separate events or they were amplified

406



once long ago and have since diverged. Examination of other amplified esterase genesfrom different

Culex

strains clearly reveals numerous differences at the molecularlevel and strongly suggests that multiple amplification events have occurred withthem (Vaughan et al. 1995).

In addition to amplification, esterases may be mutated to produce structurally dif-ferent enzymes which are able to metabolize insecticides more efficiently. In the Aus-tralian sheep blow fly

Lucilia cuprina

and the mosquito

C. tarsalis,

acarboxylesterase appears to be structurally altered in resistant populations to pro-duce high levels of resistance to malathion (Whyard et al. 1994a, 1994b). In neitherspecies is more of the enzyme produced. The difference between susceptible and resis-tant populations is strictly a qualitative difference in the enzyme produced. Whetheror not the carboxylesterases from the blow fly and mosquito are homologous to eachother will only be known through further molecular and biochemical analysis.

The final group of enzymes which may provide metabolic resistance are the GSTs.Both organophosphate and cyclodiene pesticides can be detoxified by GST pathways.These enzymes have been somewhat less studied at the biochemical and molecularlevel in insects than the P450s and esterases. Elevated GST levels are found in manyresistant insect strains (Motoyama & Dauterman 1975, Ottea & Plapp 1984, Aham-mad-Sahib et al. 1994, Hoffman & Fisher 1994) and increased GST activity is clearlythe underlying resistance mechanism in some cases (Kao & Sun 1991, Wang et al.1991, Prapanthadara et al. 1993). But in other resistant populations the increasedGST activity does not cause resistance (Bush et al. 1993, Hemingway et al. 1993, Ar-gentine et al. 1994). Both insecticides and plant allelochemicals induce increased GSTproduction (Yu 1992a, Lagadic et al. 1993, Leszczynski et al. 1994), and generalistplant feeders seem to rely on GST pathways more heavily than do specialist feedersto metabolize plant allelochemicals (Yu 1992b). Because GSTs can metabolize a widevariety of substances, increased GST activity may be part of a generalized compensa-tory change due to exposure to an environmental stress. How GST activity is in-creased has only been examined in a few Diptera. In dipterans, increased GSTactivity does not appear to be the result of gene amplification. In both the house flyand

Drosophila

, several GSTs contribute to resistance and their expression appears tohave been increased through a regulatory change (Wang et al. 1991, Cochrane et al.1992, Fournier et al. 1992). In addition, at least one resistance gene is structurally al-tered in

Drosophila

(Cochrane et al. 1992). Whether or not this structural change af-fects the resistance level in the flies remains to be determined. As numerous GSTshave been found in several insects, each of which appears to be encoded by a differentgene (Cochrane et al. 1992, Fournier et al. 1992, Baker et al. 1994), it is likely that re-sistance caused by GSTs is due to altered gene expression and/or structural changesand is not due to gene amplification in most, if not all, cases.

The second major resistance mechanism is target-site insensitivity, which refers toan alteration of the molecule(s) that directly interacts with the pesticide to reduce tox-icity. Both acetylcholinesterase (AChE) and the gamma-aminobutyric acid (GABA) re-ceptor are known targets of insecticides, and resistant alleles of each have been found.Voltage-gated sodium channels and the juvenile hormone (JH) receptor are putativetargets of insecticides. Their direct interaction with insecticides has not been con-firmed, but it is clearly evident that they play a key role in the intoxication process.

Acetylcholinesterase is the target site of both organophosphates and carbamates.These pesticides bind to AChE and prevent the enzyme from stopping the action of theneurotransmitter acetylcholine. Multiple forms of AChE that confer varying degreesof resistance have been found in a variety of arthropods (Nolan et al. 1972, Devonshire& Moore 1984, Pralavorio & Fournier 1992). In each case examined so far, the affinity


407

of AChE for the pesticide has been reduced. Neither gene amplification nor altered ex-pression of the gene encoding AChE has been detected. Instead, point mutations haveoccurred to structurally change the enzyme. Recently, Mutero et al. (1994) identifiedfive point mutations in

D. melanogaster

which are associated with reduced sensitivity.Several strains of resistant flies were found to have a combination of mutations. Indi-vidually, the mutations gave only low levels of resistance, but when several of themwere combined, high levels of resistance resulted. Mutero et al. (1994) hypothesizethat decreased sensitivity by AChE is the result of a combination of several muta-tions, each of which provides a little resistance instead of the appearance of a singlemutation which yields strong resistance.

GABA receptors are the primary target of cyclodiene insecticides. In vertebrates,these receptors group together to form a complete chloride ion channel. It is inferredthat invertebrates have similar ion channels. Most cases of cyclodiene resistance ap-pear to be due to decreased sensitivity of the GABA subtype A receptor (ffrench-Con-stant et al. 1991), an integral part of the chloride ion channel. Like AChE, decreasedsensitivity by GABA receptors is due to a structural change of the protein. Neitheramplification nor altered expression of the GABA receptor gene has been detected.Unlike AChE, only a single point mutation which causes one specific amino acid to besubstituted with another results in high levels of resistance to cyclodienes. Otherpoint mutations have been detected, but no others appear to cause resistance or areconsistently associated with resistance (ffrench-Constant et al. 1993, Thompson et al1993).

Voltage-gated sodium channels play an integral role in the transmission of neuralimpulses. Pyrethroids disrupt neural transmissions by interrupting the normal func-tioning of voltage-gated sodium channels. Target-site insensitivity to pyrethroids, aphenotypic response commonly referred to as knockdown resistance (kdr), results inthese sodium channels becoming less sensitive to intoxication. Although it is clearthat sodium channels are adversely affected by pyrethroids, there is disagreement asto whether or not kdr is the result of a mutation to the sodium channels or to someother molecule which is integral to the functioning of the sodium channels. A pointmutation to a sodium channel, which could structurally alter it, was detected in a re-sistant insect strain (Amichot et al. 1992) but so far this mutation has not been shownto cause resistance. Other reports indicate that the mutation which causes kdr isclosely linked to (physically close to or a part of) the gene encoding one type of sodiumchannel (Knipple et al. 1994, Dong & Scott 1994) and this linkage has been inter-preted as strong evidence that a mutation(s) to the sodium channel gene is associatedwith kdr. On the other hand, a mutation to a regulatory protein or receptor could alsoresult in kdr. There is some indirect evidence to support this alternative hypothesis(Rossignol 1991, Osborne & Pepper 1992). A third hypothesis that has been proposedand for which there is limited electrophysiological evidence is that kdr is caused bychanges to two closely linked genes. One involves an altered sodium channel and theother may be associated with calcium-activated phosphorylation of a protein(s) in-volved with neurotransmitter release (Pepper & Osborne 1993). At this time, it canonly be stated that there is no indication that a gene amplification event is associatedwith kdr. It is most likely that either the expression of a gene associated with sodiumchannel function has been altered and/or a structural mutation to sodium channelsresults in kdr.

Juvenile hormone analogs such as methoprene compete with the natural hormonefor the JH receptor. Most resistance to JH analogs is either metabolic and/or a reduc-tion in penetration of the insecticide through the cuticle. The only reports of targetsite insensitivity to JH analogs are in mutants isolated from laboratory colonies of

408



Drosophila

(Shemshedini & Wilson 1990, Wilson & Turner 1992). The resistance gene

Met

that has been isolated from these colonies is associated with a less sensitive cys-tolic JH binding protein. Transposition of alleles of

Met

by a TE can induce resistance(Turner 1993); however, transposition is not required for the resistance gene to be ex-pressed. Therefore, it seems most likely that the insensitive JH binding protein is ei-ther the result of a mutation that structurally changes the protein or the expressionof the insect growth cycle has been altered. Gene amplification of JH receptors has notbeen implicated.

Two other types of resistance that have been described are the reduced penetra-tion of a pesticide and altered behavior to avoid a pesticide. It is presumed that the cu-ticular structure is somehow altered to reduce the rate of penetration of a pesticide.Avoidance behavior appears to be stimulated by brief contact, either through tactileor olfactory receptors, with a pesticide. Alone, neither of these mechanisms cause highlevels of resistance, but they are often found in combination with other types of resis-tance and can make a significant contribution to the overall resistance displayed byan insect. For example, Raymond et al. (1989) calculated that reduced penetration in-teracts with any other resistance mechanism multiplicatively. Experimental datasupport this conclusion (Hoyer & Plapp 1968, Plapp & Hoyer 1968). The underlyingphysiological, genetic and molecular mechanisms that cause these types of resistancecan only be speculated and the genetic mutations which cause them cannot be in-ferred at this time.

D

ETECTING

AND

M

ONITORING

FOR

R

ESISTANCE

Resistance is a widespread phenomenon and resistant populations of nearly alleconomically important pests can now be found (Georghiou 1994, Leibee & Capinerathis issue). Where control failures have occurred, the history of pesticide applicationusually indicates that chronic pesticide exposure resulted in high levels of resistancewhich caused the failures. Does the development of a resistant population then meanthat a control failure is inevitable? Certainly the continuous selection of the same re-sistance mechanism(s) over and over will result in resistance levels that are highenough to cause a control failure. But if the selection pressure (i.e. the pesticide) foreach resistance mechanism is removed prior to significant resistance developing, thena control failure may be avoided (for an alternative view on this subject, see Hoy thisissue). Keys to the successful avoidance of a control failure are the detection of resis-tance at low level(s), the routine monitoring for changes in the level(s) and type(s) ofresistance present in the pest population and the implementation of, and strict adher-ence to, a multi-tactic pest management program. In many cases, the two former ele-ments have not been fully utilized to allow for the design of an effective pestmanagement program. This lack of implementation is due in part to the unavailabil-ity of easy to use resistance detection methods.

The ideal detection method is fast, inexpensive, easy to use, diagnostic for all typesof resistance and able to detect resistance at frequencies as low as 1%. Numerousmethods to detect resistance are currently available but all fall short of being the idealtechnique. In fact, it is very unlikely that such an ideal technique will ever be devel-oped. Instead, we must rely upon traditional methods, principally bioassays and alimited number of biochemical assays, and novel molecular techniques to detect andmonitor for resistance. Certain general features are shared by the detection assayswithin each of these two technological groupings, i.e. traditional and molecular; andconsequently, they have similar advantages and disadvantages which are summa-rized in Table 2.


409

In general, detection methods which utilize molecular technology are better able todistinguish between the different resistance genotypes, i.e. heterozygotes (SR), ho-mozygous susceptible (SS) and homozygous resistant (RR), than traditional detectionassays. Because they detect only genetic differences, molecular assays can eliminatethe environmental components which often increase the variability in bioassay andbiochemical results. Direct comparison of a molecular assay and a bioassay (Aron-stein et al. 1994) indicates that molecular assays better approximate the perfectly di-agnostic assay described by Roush & Miller (1986) than traditional bioassays and,therefore, can require up to 5-fold fewer insects to yield the same information. And un-like traditional assays, molecular techniques can use material from a single insect toperform several different assays so that the resistance levels to a wide variety of pes-ticides can be determined from the same individuals. However, molecular assays arelimited to the detection of known resistance genes and a separate assay must be donefor each gene. Traditional bioassays are better able to detect the overall level of resis-tance present in a population in a single test. In fact, molecular assays cannot detectmany types of resistance at this time since with few exceptions we do not know whichspecific genes cause resistance. In time, molecular assays will be developed to detectmore resistance genes; however, it is unlikely that molecular assays will completelyreplace the traditional ones. In addition, molecular assays are more costly in both ma-terial and equipment and require greater technical training than simple bioassays.They cannot be used in the field and usually take significantly longer to complete.

Although many resistance genes have not been isolated, it is clear from the molec-ular and genetic data that it is highly unlikely that resistance genes are the result ofnovel mutations which create new genes. Instead resistance, a phenotypic response,is the result of the selection by pesticides of alleles of pre-existing genes, i.e. specificgenotypes, that regulate or enhance particular defensive mechanisms. How many

T

ABLE

2. C

OMPARISON

OF THE ADVANTAGES AND DISADVANTAGES OF DETECTIONMETHODS THAT ARE BASED ON EITHER MOLECULAR OR TRADITIONAL TECH-NOLOGY.

Molecular Detection Assay vs. Traditional Detection Assay

can distinguish resistant genotypes even at low levels

less sensitive; only detects resistant phenotypes, not genotypes

distinguishes SS, SR, and PR genotypes reliably even when

phenotypes are hard to distinguish

cannot distinguish SR and RR from each other unless the phenotypes are

distinct from one anothergenerally greater accuracy and less variability because environmental

components are eliminated

less accurate and more variable because environmental components

cannot be separated from genetic onesfewer insects can yield more

information because material from one insect can be used for several assays

requires more insects to get the same data because the same individuals cannot be used in several bioassays

currently not adapted for field use many are easily used in the fieldgenerally less rapid, it may take days

to complete a testvery rapid, often only requiring a

couple of hours to domaterial and equipment often costly; more technical expertise often needed

generally inexpensive and simple to prepare and execute

limited to detection of known resistance genes

can detect any type of resistance, even if the resistance gene is not known


genes can cause resistance is not known. Data on various esterases and cytochromeP450s suggests that multiple genes may yield the same type of resistance. Only fur-ther analyses will determine if this is true. How resistant genes are distributedwithin and between populations is also not clear and requires further study. Under-standing the genetic flow of resistance genes and the biological costs of resistance areof particular importance because without this information it will be difficult, if not im-possible, to design successful control programs for many pests. Hopefully, the elucida-tion of the molecular genetics of resistance genes will help in the design of effectivecontrol programs and suggest novel control methods.

ACKNOWLEDGMENTS

The author gratefully thanks M. A. Hoy and D. C. Herzog for reviewing this manu-script and J. Hemingway for providing useful insights and a preprint of a relevantmanuscript. This paper was presented as part of the symposium “The Myths of Man-aging Resistance” at the Florida Entomological Society Annual Meeting, August 8-11,1994.

REFERENCES CITED

AHAMMAD-SAHIB, K. I., R. M. HOLLINGWORTH, M. E. WHALON, P. M. IOANNIDIS AND E.J. GRAFIUS. 1994. Polysubstrate monooxygenases and other xenobiotic-metab-olizing enzymes in susceptible and resistant Colorado potato beetle. Pestic.Biochem. Physiol. 49:1-12.

AMICHOT, M., C. CASTELLA, A. CUANNY, J. B. BERGÉ AND D. PAURON. 1992. Targetmodification as a molecular mechanism of pyrethroid resistance in Drosophilamelanogaster. Pestic. Biochem. Physiol. 44:183-190.

ANDERSEN, J. F., J. G. UTERMOHLEN AND R. FEYEREISEN. 1994. Expression of housefly CYP6A1 and NADPH-cytochrome P450 reductase in Escherichia coli andreconstitution of an insecticide-metabolizing P450 system. Biochemistry33:2171-2177.

ANONYMOUS. 1957. World Health Expert Committee on Insecticides 7th Report. WHOTechnical Report Series No. 125.

ARGENTINE, J. A., K. Y. ZHU, S. H. LEE AND J. M. CLARK. 1994. Biochemical mecha-nisms of azinophosmethyl resistance in isogenic strains of Colorado potato bee-tle. Pestic. Biochem. Physiol. 48:63-68.

ARONSTEIN, K., P. ODE AND R. J. FFRENCH-CONSTANT. 1994. Direct comparison ofPCR-based monitoring for cyclodiene resistance in Drosophila populationswith insecticide bioassay. Pestic. Biochem. Physiol. 48:229-233.

BAKER, W. L., A. G. CLARK, G. FAULDS AND J. S. NIELSEN. 1994. Multiple glutathioneS-transferases in Galleria mellonella; their detection with fluorigenic sub-strates. Insect Biochem. Molec. Biol. 24:301-307.

BERG, D. E. 1989. Transposable elements in prokaryotes, pp. 99-137 in S. B. Levy andR. V. Miller [eds.] Gene Transfer in the Environment. McGraw-Hill, New York,New York.

BOTHWELL, M. L., D. M. J. SHERBOT AND C. M. POLLOCK. 1994. Ecosystem responseto solar ultraviolet-B radiation: influence of trophic-level interations. Science265-97-100.

BUSH, M. R., Y. A. I. ABDEL-AAL, K. SAITO AND G. C. ROCK. 1993. Azinophosmethyl re-sistance in the tufted apple bud moth (Lepidoptera: Torticidae): reversion, di-agnostic concentrations, associated esterases, and glutathione transferases. J.Econ. Entomol. 86:213-225.

CARIÑO, F. A., J. F. KOENER, F. W. PLAPP AND R. FEYEREISEN. 1994. Constitutiveover-expression of the cytochrome P450 gene CYP6A1 in a house fly strain withmetabolic resistance to insecticides. Insect Biochem. Molec. Biol. 24:411-418.

Scott: Symposium on Pesticide Resistance 411

CARLINI, E. J., B. A. MCPHERON, C. M. FELLAND AND L. A. HULL. 1991. Elevated es-terase activity in resistant tufted apple bud moth, Platynota idaeusalis(Walker) (Lepidoptera: Tortricidae). Comp. Biochem. Physiol. 99C:375-377.

CHEN, W-L. AND C. N. SUN. 1994. Purification and characterization of carboxy-lesterases of a rice brown planthopper, Nilaparvata lugens Ståal. Insect. Bio-chem. Molec. Biol. 24:347-355.

COCHRANE, B. J., M. HARGIS, P. CROCQUET-DE-BELLIGNY, F. HOLTSBERG, AND J. COR-ONELLA. 1992. Evolution of glutathione S-transferases associated with insecti-cide resistance in Drosophila. ACS Symp. Ser. American Chem. Soc.Washington, D.C.: The Society 505:53-70.

DEVONSHIRE, A. L. AND G. D. MOORE. 1982. A carboxylesterase with broad substratespecificity causes organophosphorus, carbamate and pyrethroid resistance inpeach-potato aphids (Myzus persicae). Pestic. Biochem. Physiol. 18:235-246.

DEVONSHIRE, A. L. AND G. D. MOORE. 1984. Different forms of insensitive acetylcho-linesterase in insecticide-resistant house flies (Musca domestica). Pestic. Bio-chem. Physiol. 21:336-340.

DONG, K. AND J. G. SCOTT. 1994. Linkage of kdr-type resistance and the para-homol-ogous sodium channel gene in German cockroaches (Blatella germanica). In-sect Biochem. Molec. Biol. 24:647-654.

FEYEREISEN, R., J. F. KOENER, F. A. CARIÑO AND A. S. DAGGETT. 1990. Biochemistryand molecular biology of insect cytochrome P450., pp. 263-272 in H. H. Hage-dorn, J. G. Hildebrand, M. G. Kidwell, and J. H. Law [eds.] Molecular InsectScience, Plenum, New York.

FFRENCH-CONSTANT, R. H., D. P. MORTLOCK, C. D. SHAFFER, R. J. MACINTYRE AND R.T. ROUSH. 1991. Molecular cloning and transformation of cyclodiene resistancein Drosophila: an invertebrate γ-aminobutyric acid subtype A receptor locus.Proc. Natl. Acad. Sci. USA 88:7209-7213.

FFRENCH-CONSTANT, R. H., J. C. STEICHEN, T. A. ROCHELEAU, K. ARONSTEIN AND R.T. ROUSH. 1993. A single-amino acid substitution in a γ-aminobutyric acid sub-type A receptor locus is associated with cyclodiene insecticide resistance inDrosophila populations. Proc. Natl. Acad. Sci. USA 90:1957-1961.

FIELD, L. M., A. L. DEVONSHIRE AND B. G. FORDE. 1988. Molecular evidence that in-secticide resistance in peach-potato aphids (Myzus persicae Sulz.) results fromamplification of an esterase gene. Biochem. J. 251:309-312.

FOURNIER, D., J. M. BRIDE, C. MOUCHÈS, M. RAYMOND, M. MAGNIN, J-B. BERGÉ, N.PASTEUR AND G. P. GEORGHIOU. 1987. Biochemical characterization of the es-terases A1 and B1 associated with organophosphate resistance in Culex pipiensL. complex. Pestic. Biochem. Physiol. 27:211-217.

FOURNIER, D., J. M. BRIDE, M. POIRIÉ, J-B. BERGÉ AND F. W. PLAPP. 1992. Insect glu-tathione S-transferases: biochemical characteristics of the major forms fromhouseflies susceptible and resistant to insecticides. J. Biol. Chem. 267:1840-1845.

GEORGHIOU, G. P. 1994. Principles of insecticide resistance management. Phytopro-tection 75 (suppl.): 51-59.

HARITOS, V. S., J. R. J. FRENCH AND J. T. AHOKAS. 1994. Cytochrome P450 monooxy-genase and glutathione S-transferase activity of two Australian termites: Mas-totermes darwiniensis and Coptotermes acinaciformis. Insect Biochem. Molec.Biol. 24:929-935.

HATANO, R. AND J. G. SCOTT. 1993. Anti-P450Lpr antiserum inhibits the activation ofchlorpyrifos to chlorpyrifos oxon in house fly microsomes. Pestic. Biochem.Physiol. 45:228-233.

HEMINGWAY, J., G. J. SMALL AND A. G. MONRO. 1993. Possible mechanisms of organ-ophosphorus and carbamate insecticide resistance in german cockroaches (Dic-tyoptera: Blattelidae) from different geographical areas. J. Econ. Entomol.86:1623-1630.

HOFFMAN, E. R. AND S. W. FISHER. 1994. Comparison of a field and laboratory-derivedpopulation of Chironomus riparius (Diptera: Chironomidae): biochemical andfitness evidence for population divergence. J. Econ. Entomol. 87:318-325.


HOY, M. A. 1995. Multitactic resistance management: An approach that is long over-due? Florida Entomol. 78:443-451.

HOYER, R. F. AND F. W. PLAPP. 1968. Insecticide resistance in the house fly: identifi-cation of a gene that confers resistance to organotin insecticides and acts as anintensifier of parathion resistance. J. Econ. Entomol. 61:1269-1276.

JEONG, T. C., H. G. JEONG AND K-H. YANG. 1992. Induction of cytochrome P-450 bydimethyl sulfoxide in primary cultures of adult rate hepatocytes. Toxicol. Let-ters 61:275-281.

KAO, C. H. AND C. N. SUN. 1991. In vitro degradation of some organophosphorus in-secticides by susceptible and resistant diamondback moth. Pestic. Biochem.Physiol. 41:132-141.

KARUNARATNE, S. H. P. P., K. G. I. JAYAWARDENA, J. HEMINGWAY AND A. J. KETTER-MAN. 1993. Characterization of a B-type esterase involved in insecticide resis-tance from the mosquito Culex quinquefasciatus. Biochem. J. 294:575-579.

KETTERMAN, A. J., K. G. I. JAYAWARDENA AND J. HEMINGWAY. 1992. Purification andcharacterization of a carboxylesterase involved in insecticide resistance fromthe mosquito Culex quinquefasciatus. Biochem. J. 287:355-360.

KETTERMAN, A. J., S. H. P. P. KARUNARATNE, K. G. JAYAWARDENA AND J. HEMING-WAY. 1993. Qualitative differences between populations of Culex quinquefas-ciatus in both the esterases A2 and B2 which are involved in insecticideresistance. Pestic. Biochem. Physiol. 47:142-148.

KNIPPLE, D. C., K. E. DOYLE, P. A. MARSELLA-HERRICK AND D. M. SODERLUND. 1994.Tight genetic linkage between the kdr insecticide resistance trait and a volt-age-sensitive sodium channel gene in the house fly. Proc. Natl. Acad. Sci. USA91:2483-2487.

KOEHN, R. K. AND B. L. BAYNE. 1989. Towards a physiological and genetical under-standing of the energetics of the stress response. Biological Journal of the Lin-nean Society 37:157-171.

LAGADIC, L., A. CUANY, J. B. BERGÉ AND M. ECHAUBARD. 1993. Purification and par-tial characterization of glutathione S-tranferases from insecticide-resistantand lindane-induced susceptible Spodoptera littoralis (Boisd.) larvae. InsectBiochem. Mol. Biol. 23:467-474.

LEIBEE, G. L. AND CAPINERA, J. L. 1995. Pesticide resistance in Florida insects limitsmanagement options. Florida Entomol. 78:386-399.

LESZCZYNSKI, B., M. MATOK AND A. F. G. DIXON. 1994. Detoxification of cereal plantallelochemicals by aphids: activity and molecular weights of glutathione S-transferase in three species of cereal aphids. J. Chem. Ecol. 20:387-394.

MOTOYAMA, N. AND W. C. DAUTERMAN. 1975. Interstrain comparison of glutathione-dependent reactions in susceptible and resistant houseflies. Pestic. Biochem.Physiol. 5:489-495.

MOUCHÈS, C., N. PASTEUR, J. B. BERGÉ, O. HYRIEN, M. RAYMOND, B. R. DE SAINT VIN-CENT, M. DE SILVESTRI G. P. GEORGHIOU. 1990. Amplification of an esterasegene is responsible for insecticide resistance in a California Culex mosquito.Science 233:778-780.

MUEHLEISEN, D. P., J. H. BENEDICT, F. W. PLAPP, JR. AND F. A. CARIÑO. 1989. Effectsof cotton allelochemicals on toxicity of insecticides and induction of detoxifyingenzymes in bollworm (Lepidoptera: Noctuidae). J. Econ. Entomol. 82:1554-1558.

MUTERO, A., M. PRALAVORIO, J-M. BRIDE AND D. FOURNIER. 1994. Resistance-associ-ated point mutations in insecticide-insensitive acetylcholinesterase. Proc.Natl. Acad. Sci. USA 91:5922-5926.

NOLAN, J., H. J. SCHNITZERLING AND C. A. SCHUNTNER. 1972. Multiple forms of ace-tylcholinesterase from resistant and susceptible strains of the cattle tick, Boo-philus microplus (Can.). Pestic. Biochem. Physiol. 2:85-94.

OSBORNE, M. P. AND D. R. PEPPER. 1992. Mechanisms of kdr and super-kdr resis-tance. ACS Symp. Ser. American Chem. Soc. 505:71-89.

Scott: Symposium on Pesticide Resistance 413

OTTEA, J. A. AND F. W. PLAPP. 1984. Glutathione-S-transferase in the house fly: bio-chemical and genetic changes associated with induction and insecticide resis-tance. Pestic. Biochem. Physiol. 22:203-208.

PEPPER, D. R. AND M. P. OSBORNE. 1993. Electrophysiological identification of site-in-sensitive mechanisms in knockdown-resistant strains (kdr, super-kdr) of thehousefly larva (Musca domestica). Pestic. Sci. 39:279-286.

PLAPP, F. W. AND R. F. HOYER. 1968. Insecticide resistance in the house fly: decreasedrate of absorption as the mechanism of action of a gene that acts as an intensi-fier of resistance. J. Econ. Entomol. 61:1298-1303.

POIRIÉ, M., M. RAYMOND AND N. PASTEUR. 1992. Identification of two distinct ampli-fications of the esterase B locus in Culex pipiens (L.) mosquitoes from Mediter-ranean countries. Biochem. Genet. 30:13-26.

PRALAVORIO, M. AND D. FOURNIER. 1992. Drosophila acetylcholinesterase: character-ization of different mutants resistant to insecticides. Biochem. Genet. 30:77-83.

PRAPANTHADARA, L. A., J. HEMINGWAY AND A. J. KETTERMAN. 1993. Partial purifica-tion and characterization of glutathione S-transferases involved in DDT resis-tance from the mosquito Anopheles gambiae. Pestic. Biochem. Physiol. 47:119-133.

RAYMOND, M., D. G. HECKEL AND J. G. SCOTT. 1989. Interactions between pesticidegenes: model and experiment. Genetics 123:543-551.

RAYMOND, M., A. CALLAGHAN, P. FORT AND N. PASTEUR. 1991. Worldwide migrationof amplified insecticide resistance genes in mosquitoes. Nature 350:151-153.

ROSE, R. L., F. GOULD, P. E. LEVI AND E. HODGSON. 1991. Differences in cytochromeP450 activities in tobacco budworm larvae as influenced by resistance to hostplant allelochemicals and induction. Comp. Biochem. Physiol. 99B:535-540.

ROSSIGNOL, D. P. 1991. Analysis of pyrethroid binding by use of a photoreactive ana-logue: possible role for GTP-binding proteins in pyrethroid activity. Pestic. Bio-chem. Physiol. 41:103-120.

ROUSH, R. T. AND G. L. MILLER. 1986. Considerations for design of insecticide resis-tance monitoring programs. J. Econ. Entomol. 79:293-298.

SCOTT, J. A., F. H. COLLINS AND R. FEYEREISEN. 1994. Diversity of cytochrome P450genes in the mosquito, Anopheles albimanus. Biochem. Biophys. Res. Comm.205:1452-1459.

SHEMSHEDINI, L. AND T. G. WILSON. 1990. Resistance to juvenile hormone and an in-sect growth regulator in Drosophila is associated with an altered cytosolic ju-venile hormone-binding protein. Proc. Natl. Acad. Sci. USA 87:2072-2076.

SNYDER, M. J., E-L. HSU AND R. FEYEREISEN. 1993. Induction of cytochrome P-450 ac-tivities by nicotine in the tobacco hornworm, Manduca sexta. J. Chem. Ecol.19:2903-2916.

THOMPSON, M., J. C. STEICHEN AND R. H. FFRENCH-CONSTANT. 1993. Conservation ofcyclodiene insecticide resistance-associated mutations in insects. Insect Molec.Biol. 2:149-154.

VAUGHAN, A., M. RODRIGUEZ AND J. HEMINGWAY. 1995. The independent gene ampli-fication of indistinguishable esterase B electromorphs in insecticide-resistantCulex quinquefasciatus. Biochem. J. 305:651-658.

WANG, J-Y., S. MCCOMMAS AND M. SYVANEN. 1991. Molecular cloning of a glutathioneS-transferase overproduced in an insecticide-resistant strain of the housefly(Musca domestica). Mol. Gen. Genet. 227:260-266.

WATERS, L. C., A. C. ZELHOF, B. J. SHAW AND L-Y. CHANG. 1992. Possible involvementof the long terminal repeat of transposable element 17.6 in regulating expres-sion of an insecticide resistance-associated P450 gene in Drosophila. Proc.Natl. Acad. Sci. USA 89:4855-4859.

WAXMAN, D. J. AND L. AZAROFF. 1992. Phenobarbital induction of cytochrome P-450gene expression. Biochem. J. 281:577-592.

WHEELOCK, G. D. AND J. G. SCOTT. 1992. The role of cytochrome P450 Lpr in delta-methrin metabolism by pyrethroid-resistant and susceptible strains of houseflies. Pestic. Biochem. Physiol. 43:67-77.


WHYARD, S., R. J. RUSSELL AND V. K. WALKER. 1994a. Insecticide resistance andmalathion carboxylesterase in the sheep blowfly, Lucilia cuprina. Biochem.Genet. 32:9-22.

WHYARD, S., A. E. DOWNE AND V. K. WALKER. 1994b. Isolation of an esterase confer-ring insecticide resistance in the mosquito Culex tarsalis. Insect Biochem.Molec. Biol. 24:819-827.

WILSON, T. G. AND C. TURNER. 1992. Molecular analysis of Methoprene-tolerant, agene in Drosophila involved in resistance to juvenile hormone analog insectgrowth regulators, in C. Mullin and J. G. Scott [eds.], Molecular mechanisms ofinsecticide resistance. American Chemical Society Symp. Series 505:99-112.

YU, S. J. 1992a. Plant-allelochemical-adapted glutathione transferases in Lepi-doptera. ACS Symp. Ser. American Chem. Soc. 505:174-190.

YU, S. J. 1992b. Detection and biochemical characterization of insecticide resistance

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

in fall armyworm (Lepidoptera: Noctuidae). J. Econ. Entomol. 85:675-682.

414



RESISTANCE: A THREAT TO THE INSECTICIDAL CRYSTAL PROTEINS OF

BACILLUS THURINGIENSIS

L

EAH

S. B

AUER

USDA Forest Service, North Central Forest Experiment StationPesticide Research Center & Department of Entomology

Michigan State UniversityEast Lansing, MI 48823-5290

A

BSTRACT

Insecticidal crystal proteins (also known as

δ

-endotoxins) synthesized by the bac-terium

Bacillus thuringiensis

Berliner (

Bt

) are the active ingredient of various envi-ronmentally friendly insecticides that are 1) highly compatible with natural enemiesand other nontarget organisms due to narrow host specificity, 2) harmless to verte-brates, 3) biodegradable in the environment, and 4) highly amenable to genetic engi-neering. The use of transgenic plants expressing

Bt

δ

-endotoxins has the potential togreatly reduce the environmental and health costs associated with the use of conven-tional insecticides. The complex mode of action of

Bt

is the subject of intensive re-search. When eaten by a susceptible insect

δ

-endotoxin crystals are solubilized in themidgut; proteases then cleave protoxin molecules into activated toxin which binds toreceptors on the midgut brush border membrane. Part of the toxin molecule insertsinto the membrane causing the midgut cells to leak, swell, and lyse; death resultsfrom bacterial septicemia. Insecticides formulated with

Bt

account for less than 1% ofthe total insecticides used each year worldwide because of high cost, narrow hostrange, and comparatively low efficacy. Environmental contamination, food safety con-cerns, and pest resistance to conventional insecticides have caused a steady increasein demand for

Bt

-based insecticides. The recent escalation of commercial interest in

Bt

has resulted in more persistent and efficacious formulations. For example, im-proved

Bt

-based insecticides have allowed management of the diamondback moth,

Plutella xylostella

(L.). Unfortunately this has resulted in the evolution of resistanceto

δ

-endotoxins in

P. xylostella

populations worldwide. The recent appearance of

Bt

re-sistance in the field, corroborated by the results of laboratory selection experiments,demonstrates genetically-based resistance in several species of Lepidoptera, Diptera,and Coleoptera. The genetic capacity to evolve resistance to these toxins is probably

Bauer: Symposium on Pesticide Resistance

415

present in all insects, and the heritability, fitness costs, and stability of the resistancetrait are documented in several insect populations. In two strains of

Bt

-resistant lep-idopteran species, mechanisms of resistance involve reductions in the binding of toxinto midgut receptors. Research on other resistant strains suggests that other mecha-nisms are also involved. Unfortunately, the high stability of the resistance trait, aswell as broad spectrum cross-resistance to other

δ

-endotoxins, undermines many po-tential options for resistance management. Genetically engineered plants, expressing

δ

-endotoxin continuously and at ultrahigh doses, ensure intense and rapid selectionof the target insect population. The efficacy of transgenic plants can be preserved onlyby developing an integrated pest management program that is designed specifically toreduce selection pressure by minimizing exposure to

Bt

and increasing other mortal-ity factors, thereby slowing the rate of pest adaptation to

Bt

.

Key Words:

δ

-endotoxin, cross-resistance, transgenic plants, resistance management

R

ESUMEN

Las proteínas de cristales insecticidas (también conocidas como

δ

-endotoxinas)sintetizadas por la bacteria


Berliner (

Bt

) son el ingrediente ac-tivo de varios insecticidas ambientalmente amistosos que son 1) altamente compati-bles con los enemigos naturales y otros organismos no objetos de control debido a suestrecha especificidad de hospedante, 2) inocuos a los vertebrados, 3) biodegradablesen el ambiente, y 4) altamente dóciles para la ingeniería genética. El uso de plantastransgénicas expesando

δ

-endotoxinas de

Bt

tiene la posibilidad

de reducir grande-mente los costos ambientales y de salud asociados con el uso de insecticidas conven-cionales. El modo complejo de acción de

Bt

es sujeto de investigación intensiva.Cuando son ingeridos por un insecto susceptible, los cristales de

δ

-endotoxina son di-sueltos en el intestino medio; las proteasas abren las moléculas de proteínas transfor-mándolas en toxinas activadas que se unen a receptores en el cepillo de la membranadel intestino medio. Parte de la molécula de la toxina se inserta en la membrana cau-sando que las células del intesitno medio pierdan el control de la permeabilidad, sehinchen y rompan, y la muerte ocurre luego por septicemia bacteriana. Los insectici-das formulados con

Bt

son menos del 1% del total de los insecticidas usados cada añoen todo el mundo debido a su alto costo, estrecho rango de hospedantes, y eficacia com-parativamente baja. La contaminación abiental y la resistencia de las plagas a los in-secticidas convencionales han causado un incremento constante en la demanda deinsecticidas de

Bt

. La reciente escalada de interés comercial en

Bt

ha provocado laaparición de formulaciones más persistentes y eficaces. Sin embargo, el uso intensivode insecticidas mejorados a base de

Bt

autorizados para el manejo de la polilla de lacol,

Plutella xyllostella

(L.), ha traído como reusltado la evolución de resistencia a las

δ

-endotoxinas en las poblaciones de

P. xyllostella

en todo el mundo. La reciente apar-ción de reistencia a

Bt

en el campo, corroborada por los resultados de experimentos deselección de laboratorio, demuestra que existe resistencia genética en varias especiesde Lepidoptera, Diptera y Coleoptera. La capacidad genética de evolucionar la resis-tencia hacia esas toxinas está probablemente presente en todos los insectos, y la he-redabilidad, costo de ajuste, y estabilidad de la resistencia son documentados envarias poblaciones de insectos. En dos especies de lepidópteros resistentes a

Bt

, losmecanismos de resistencia incluyen reducciones en la unión de la toxina a los recep-tores del intestino medio. La investigación sobre otras especies de insectos resistentessugiere que otros mecanismos están también relacionados. Desafortunadamente, laalta estabilidad de la resistencia, así como la resistencia cruzada de amplio espectroa otras

δ

-endotoxinas, determina pocas opciones potenciales para el manejo de la re-sistencia. Las plantas transgénicas expesando

δ

-endotoxinas continuamente, y las do-sis muy altas, aseguran la selección intensa y rápida de la población de insectos acontrolar. La eficacia de las plantas transgénicas puede ser preservada sólo desarro-llando un programa de manejo integrado de plagas diseñado específcamente para re-

416



ducir la presión de seleccción, minimizando la exposición a

Bt

e incrementando otros

factores de mortalidad, para disminuir la velocidad de adaptación de la plaga a

Bt

.

The bacterium


Berliner (

Bt

) is a complex of subspecies char-acterized by their ability to synthesize crystalline inclusions during sporulation. Thesecrystalline inclusions are comprised of relatively high quantities of one or more glyco-proteins known as

δ

-endotoxins or Cry toxins (Table 1). The toxins produced by

Bt

playa vital role in the pathogenicity of this bacterium to insects and other invertebrates. TheCry toxins have enormous commercial value as safe, biodegradable pesticides. Thespecificity of

Bt

toxicity is highly desirable in integrated pest management (IPM) pro-grams, particularly in sensitive aquatic and forest ecosystems where other life forms,including many beneficial and nontarget insects, must be conserved (May 1993). The se-lective toxicity, rapid environmental degradation, and vertebrate safety of

Bt

-based in-secticides provide growers and the public with environmentally friendly and effectivealternatives to conventional insecticides (Meadows 1993). Advances in biotechnologyand genetic engineering, as well as the proteinaceous nature of the Cry toxins, led to theselection of the

cry

genes as the primary insect-resistance genes transferred into, andexpressed in, plants and microbes (Gasser & Fraley 1989, Adang 1991, Peferoen 1992,Ely 1993, Gelernter & Schwab 1993).

Subspecies of

Bt

are distributed in a variety of diverse habitats worldwide (Martin& Travers 1989) and are typically isolated from soil, leaf surfaces, and environmentsrich with insects, such as grain bins and insectaries (Smith & Couche 1991, Burges &Hurst 1977). In fact, the first reports of

Bt

were from colonies of the silkworm

Bombyxmori

(L.) (Ishiwata 1901), and from the Mediterranean flour moth,

Ephestia kueh-

T

ABLE

1.

B

T

S

TRAINS

, T

HEIR

R

ESPECTIVE

δ

-

ENDOTOXINS

AND

H

OST

R

ANGES

,

IN

I

NSECT

R

ESISTANCE

S

TUDIES

.

Bt

Strain

1

δ

-Endotoxin

2

Spectrum

kurstaki

HD-1 (

Btk

) CryIA(a), CryIA(b), CryIA(c), CryIIA, CryIIB

LepidopteraDiptera

3

kurstaki

HD-73 CryIA(c) Lepidoptera

aizawai

HD-112 (

Bta

) CryIA(a), CryIA(b), CryIC, CryID, CryIG, CryII

4

Lepidoptera

aizawai

HD-133 (

Bta

) CryIA(a), CryIA(b), CryIC, CryID Lepidoptera

thuringiensis

HD-2 CryIA, CryIB LepidopteraColeoptera

5

entomocidus

HD-198 (

Bte

) CryIA(a), CryIA(b), CryIC, CryID Lepidoptera

sotto

(

Bts

) CryIA(a) Lepidoptera

israeliensis

(

Bti

) CryIVA, CryIVB, CryIVC, CryIVD, CytA Diptera

tenebrionis

(

Btt

) CryIIIA Coleoptera

1

Bt

strains may contain multiple toxins, and composition may differ slightly from those reported here.

2

Organized by amino acid sequence by Höfte & Whiteley (1989).3CryIIA accounts for the dipteran activity of this strain.4Presence of CryIG uncertain, and CryII type unknown (McGaughey & Johnson 1994).5CryIB is toxic to some coleopterans (Bradley et al. 1995).

Bauer: Symposium on Pesticide Resistance 417

niella (Zeller) (Berliner 1911). Ecological considerations of Bt as an insect pathogenand in the environment are discussed by Meadows (1993).

In the United States, commercial development of Bt into a formulated insecticidedid not begin until the late 1950s. Most Bt-based insecticides are formulated mixturesof δ-endotoxin crystals and Bt spores, which are known to synergize the toxicity of thecrystals. Although the effectiveness of these early Bt-based insecticides was often er-ratic, progress was slow in research and development of improved Bt formulation, de-livery, and application technologies, as well as in the discovery of more active strains.Until the mid-1970s, it was generally accepted that lepidopterans were the only tar-get of Bt.

The discovery of Bt subsp. israelensis, which is toxic to larval mosquitoes and blackflies (Goldberg & Margalit 1977), and the discovery of Bt subsp. tenebrionis (Krieg etal. 1983), which is toxic to several beetle species, stimulated sudden and dramaticcommercial interest in Bt. During the 1980s, new biotech companies and large agro-chemical and pharmaceutical corporations initiated research programs to isolate Btfrom various environmental samples and to screen for toxicity in agriculturally andmedically-important target organisms (Van Frankenhuyzen 1993). Lambert & Pefer-oen (1992) estimated that 40,000 strains of Bt are now stored, mainly in private col-lections, worldwide. The spectrum of activity of Bt toxins has expanded from speciesin three insect orders (Lepidoptera, Diptera, and Coleoptera) to species in eight insectorders (Homoptera, Orthoptera, Mallophaga, Hymenoptera, Siphanoptera) (Bauer,unpublished) and various mites, nematodes, flukes, mollusks, and protozoans (Feitel-son et al. 1992). Commercial products formulated with Bt are now registered for con-trol of lepidopteran (Navon 1993), dipteran (Becker & Margalit 1993), andcoleopteran pests (Keller & Langenbruch 1993). The short half-life of Bt, due to ultra-violet inactivation when topically applied, has stimulated considerable research intoalternative delivery strategies. By far the most controversial strategy is the use of in-sect-resistant crops expressing Bt δ-endotoxin genes, which are already in the field inAsia and the United States where potato, cotton and corn are registered.

As concerns over environmental quality and food safety increase, Bt-based insec-ticides will become increasingly important in the development of IPM strategies. Cur-rently, the use of Bt in insect control programs accounts for less than 1% ofinsecticides used worldwide each year. The comparatively high production cost of Bt-based insecticides is a primary impediment to more widespread usage. However, themajor impetus for greater use of Bt in agriculture is the development of resistance toconventional insecticides (Georghiou 1994, Watkinson 1994). In fact, many growerstypically add Bt to conventional sprays because of concerns about chemical controlfailure (Marrone & MacIntosh 1993). Because of their environmental safety, microbialinsecticides are one of the few pesticides that can be developed and registered quicklyand cheaply. In addition, resistance to conventional insecticides does not confer cross-resistance to Bt toxins due to the unique mode of action of δ-endotoxin (Stone et al.1991, Tabashnik 1994a).

Resistance is a major problem associated with the intensive use of pesticides in ag-riculture and human health protection, and hundreds of insect and mite species canno longer be controlled by one or more pesticides (Georgiou & Lagunes 1988). Resis-tance is documented in diverse groups of insecticides, including neurotoxins, chitinsynthesis inhibitors, juvenile hormone analogues (National Research Council 1986),and, most recently, Bt (Tabashnik et al. 1990). Adaptation to individual insecticides isa consequence of their intensive and prophylactic use, and many conventional insec-ticides are being lost to resistance faster than industry can replace them. Many re-searchers now predict that the use of Bt cry genes in the genetic engineering of insect-


resistant plants will expedite selection for resistance in the target organisms morerapidly than has occurred through conventional application methods (Gould 1988a,1988b, Van Rie 1991, McGaughey & Whalon 1992, Marrone & MacIntosh 1993, May1993, Whalon & McGaughey 1993, McGaughey 1994, Tabashnik 1994a).

Our understanding of the nature of Bt insecticidal crystal proteins has advancedrapidly since the mid-1980s, thanks to highly collaborative research programs involv-ing entomologists, microbiologists, physiologists, geneticists, protein biochemists,and molecular biologists. Research on Bt is fast-paced, often involving worldwide liai-sons between researchers in industry, university, and government, leading to anabundance of new research results, methodologies, and discoveries. In this paper, myobjectives are to provide researchers, even those only peripherally involved or knowl-edgeable of Bt δ-endotoxins, with a greater understanding of 1) Bt mode of action, 2)resistance and cross-resistance to Cry toxins, and 3) resistance management, as theyrelate to the use of both improved Bt insecticides and genetically-engineered plantsexpressing δ-endotoxin genes. The research results presented here will complimentand update those reported in the comprehensive review by Tabashnik (1994a). In pre-senting this information, I hope to attract more researchers into the complex area ofIPM and management of Bt resistance. The design, validation, and implementation ofeffective and sound IPM practices are some of the greatest challenges facing research-ers in agriculture, agroforestry, and vector control today.

BT MODE OF ACTION

A generalized flow chart of the events leading to Bt intoxication by Cry toxins in asusceptible host reveals various levels at which resistance might evolve in an insectpopulation (Fig. 1). The high degree of host specificity, as well as the complexity of Btmode of action, results from the interaction of the toxin within the complex environ-ment of the insect’s midgut lumen and on the surface of the midgut epithelial cells(English & Slatin 1992). Although researchers discovered relatively early that themidgut was the primary site of δ-endotoxin activity (Heimpel & Angus 1959), the mo-lecular mechanisms of Bt intoxication continue to be the subject of intensive research(for reviews see Gill et al. 1992, English & Slatin 1992, Lambert & Peferoen 1992,Aronson 1993, Honée & Visser 1993, Knowles & Dow 1993, Yamamoto & Powell 1993,Federici 1993, Visser et al. 1993). A small family of Bt δ-endotoxins known as the cy-tolytic or Cyt toxins, an important component of Bt subsp. israelensis (Table 1), is notcovered in this review (see Chilcott et al. 1990, Koni & Ellar 1994, Wu et al. 1994).

Ingestion

Feeding stimulants are known to greatly enhance Bt performance since most sus-ceptible insects stop feeding after consuming food treated with δ-endotoxin. Detectionand behavioral avoidance of food treated with Cry toxins have also been reported formany target species (Gould & Anderson 1991, Gould et al. 1991, Ramachandran et al.1993). Formulation and application technology has improved the rate of toxin inges-tion, increasing the probability that the target insect will consume a lethal dose aftertreatment.

Crystal Solubilization

Following ingestion, solubilization of crystalline δ-endotoxin is a prerequisite to allsubsequent events in the intoxication pathway (Tojo & Aizawai 1983, Du et al. 1994).


High midgut pH (>9.5) was once thought to be essential to crystal solubility, but co-leopteran-specific toxins were found to function at much lower pH (Koller et al. 1992).Midgut detergency and redox potential also affect solubilization. The rate and extentof crystal solubilization greatly influence toxicity levels in different hosts (Aronson etal. 1991, Bradley et al. 1995), as well as the rate of intoxication (Bauer & Pankratz1992, Koller et al. 1992).

Enzymatic Processing

The Cry proteins are synthesized as protoxins that require processing by midgutenzymes to generate activated toxin (Ogiwara et al. 1992). The larger protoxins ofabout 130-140 kDa (e.g. CryI) are proteolytically cleaved, exposing the activated toxinwhich is a protease-resistant core of about 55-65 kDa (Höfte & Whiteley 1989). Manyother toxins (e.g. CryII, CryIII, CryIVD) are synthesized as 70 kDa proteins and aresimilar to the N-terminal half of the larger protoxins. Enzymatic processing of thesesmaller 70 kDa toxins also occurs, with amino acids cleaved from the N terminus(Carroll et al. 1989).

Receptor Binding

The action of Cry δ-endotoxins on the midgut epithelium begins with binding ofthe activated toxin to receptors (Hoffman et al. 1988a, 1988b). Much of the host spec-ificity of Bt toxins results from their ability to bind to specific receptors on the brushborder membrane, although most toxins bind to more than one receptor (Van Rie et al.1989, 1990a, Denolf et al. 1993, Estada & Ferré 1994). Amino acid sequence similarityin the receptor-binding domain of the toxin molecule is a useful predictor of overallhost specificity (Van Rie et al. 1990, Cummings & Ellar 1994). The binding domain isalso the most variable region of the toxin molecule (Li et al. 1991).

The function of these receptors in midgut physiology is an elusive research ques-tion. Recently, aminopeptidase N, a 120 kDa glycoprotein, was purified from the lep-idopteran Manduca sexta and identified as the receptor for CryIA(c) (Knight et al.1994, Sangadala et al. 1994). Aminopeptidase N is an abundant Zn+-dependent ec-toenzyme present in the brush border of membranes of the alimentary tracts of mostanimals (Ellar 1994, Garczinski & Adang in press).

Binding, while essential, is not sufficient to produce mortal damage, as shown byseveral studies that found specific binding of toxins to receptors on brush border prep-arations is not correlated with in vivo toxicity (Van Rie 1990a, Wolfersberger 1990,Ferré et al. 1991, Garczynski et al. 1991, Gould et al. 1992, Escriche et al. 1994, San-chis et al. 1994, Masson et al. 1995). Other researchers showed that in vivo toxicity is,however, strongly correlated with measures of membrane disruption (Wolfersberger1991) or membrane permeability (Carroll & Ellar 1993). They stressed that, althoughreceptors play an essential role, post-binding factors are required for successful intox-ication by Bt δ-endotoxins.

Intercalation, Pore Formation, and Cell Lysis

After binding to a receptor on the cell surface, the toxin then inserts or intercalatesinto the plasma membrane (English & Slatin 1992, Knowles & Dow 1993). Evidencefrom electrical conductance and ion leakage studies suggests that several toxin/recep-tor complexes aggregate to form lesions or leaky regions in the brush border mem-brane (Walters et al. 1993, 1994). Using a very different method, Masson et al. (1995)


Figure 1. Generalized flow chart of events leading to Bt intoxication in a suscepti-ble host.


also detected toxin-toxin aggregation, supporting evidence that the toxin acts as amultimer on the cell surface. Pores in the plasma membrane are estimated at 1-2 nmdiam, disrupting the actively maintained osmotic balance, causing the cells to swelland burst by a process known as colloid-osmotic lysis (Knowles & Ellar 1987). Carroll& Ellar (1993) demonstrated that δ-endotoxin-induced changes in cell permeability isnon-selective to cations, anions, neutral solutes, and water. A three-dimensionalmodel of the CryIIIA protein structure supports the hypothesis that the toxin causespores or channels to form in the lipid bilayer (Li et al. 1991). However, the role of mid-gut receptors in this toxin-induced leakage is still unclear (Knowles & Dow 1993,Parenti et al. 1995, Garczynski & Adang, in press).

Bacterial Septicemia and Death

The synergistic effect of Bt spores, in the presence of δ-endotoxin, on insect mor-tality leads to speculation that δ-endotoxins facilitate bacterial exploitation of the nu-trients present in the insect hemolymph (Ali et al. 1985, Wilson & Benoit 1990,Borgonie 1995). Death occurs when lysis of midgut cells causes irreparable break-down of the midgut integrity, allowing Bt and other bacteria present in the lumen togain access to the body cavity. The insect hemolymph provides an excellent mediumfor bacterial growth. Death caused by bacterial septicemia usually occurs within 2-3days post-ingestion. However, the immediate cessation of feeding observed in most in-sects after ingestion of Bt (Angus 1954), as well as the rapid regenerative capabilityof midgut epithelial cells, can allow damaged regions of the midgut to heal. The actualrecovery of treated insects is dependent on many intrinsic and extrinsic factors in-cluding host genetics, age, and vigor; dosage and potency of toxin ingested; various en-vironmental factors including host plant species (Meade & Hare 1994, Moldenke et al.1994); and the presence of Bt spores and other bacteria in the insect gut (Miyasono etal. 1994).

RESISTANCE

Over several decades of commercial use, the continued efficacy of Bt-based insec-ticides led to considerable skepticism that resistance to Bt was possible (Burges 1971,Krieg & Langenbruch 1981). However, recent field and laboratory evidence suggestotherwise. The slow development of field resistance in the past may have resultedfrom low selection pressure exerted by early formulations and usage patterns (Stoneet al. 1991). Other researchers believed that the complex mode of action of Bt, ofteninvolving multiple toxins and Bt spores, provided protection against resistance be-cause a single mutation in the insect would be unlikely to affect susceptibility (Boman1981, Briese 1981, de Barjac 1987). McGaughey & Whalon (1992), however, suggestedthat at high levels of selection, the multicomponent-toxicity pathway merely expandsbehavioral and/or physiological opportunities for adaptation to Bt. Technological ad-vancements in Bt toxicity, host range, stability, formulation, application, and ulti-mately the expression in transgenic plants, are greatly improving biopesticidepotency and efficacy (Feitelson et al. 1992, Stone & Sims 1993, Carlton & Gawron-Burke 1993). Unfortunately, while providing high levels of pest suppression, im-proved efficacy will rapidly help select for the segment of the population that is capa-ble of withstanding Bt intoxication.

Possible shifts in susceptibility to Bt were first reported by Kinsinger & Mc-Gaughey (1979), with a 42-fold difference among “natural” populations of Plodia in-terpunctella (Hübner), the Indianmeal moth, and up to a 15-fold difference in Cadracautella (Walker), the almond moth. However, the underlying cause of this variation


is unknown because the history of Bt applications in these grain storage facilities wasnot reported. In a subsequent study over a five-state area, the mean median lethalconcentration (LC50) for populations of P. interpunctella from grain bins treated withBt was 1.2-fold higher than the mean LC50 for populations from untreated bins (Mc-Gaughey 1985). When these Bt-exposed populations were selected further in the lab-oratory, 30-fold resistance developed in 2 generations, and 100-fold resistance in 15generations. Initially, resistance to Bt in the Indianmeal moth was considered some-what unique because exposure to natural infestations of Bt in stored grains may in-crease the genetic variability in Bt susceptibility (Kinsinger & McGaughey 1979). Inaddition, the dark, stable and closed environment of grain bins favored selection forresistance by long toxin residual times because of no UV exposure and by minimizingthe potential of outbreeding with susceptibles from other populations.

It was not until Tabashnik et al. (1990) first reported field resistance to Bt in Ha-waiian populations of Plutella xylostella (L.), the diamondback moth, that the poten-tial for widespread resistance to Bt was generally acknowledged. Resistantpopulations of P. xylostella have also been documented in field populations from Flor-ida (Jansson & Lecrone 1990), New York (Shelton et al. 1993), the Philippines (Kirsch& Schmutterer 1988, Ferré et al. 1991), Japan (Hama et al. 1992), Thailand, and Ma-laysia (Georghiou 1994). These recent reports of resistance to Bt in the field have pro-vided credibility to the results of laboratory selections for Bt resistance (for reviews,see Briese 1981, Georghiou 1990, Stone et al. 1991, McGaughey & Whalon 1992,Tabashnik 1994a). As laboratory and field data accumulate, concern is growing thatthese unique bacterial toxins may be rendered useless as pest management tools, par-ticularly with the imminent commercialization of transgenic plants expressing singleactivated toxic fragments (May 1993, Whalon & McGaughey 1993).

Laboratory Selections

The increased effort researchers are now devoting to Bt resistance is reflected inthe many experiments designed to select for Bt resistance in the laboratory (Table 2).The results summarized in Table 2 represent selection experiments, performed andongoing during the last decade, which achieved significant levels of resistance to Btpreparations containing either a mixture of spores and native δ-endotoxin crystals, orindividual Cry toxins in various forms. These results represent only a few of the morethan 50 laboratory selection experiments performed with at least 16 insect species(Tabashnik 1994a). Significant levels of resistance have been documented in nine spe-cies of Lepidoptera and two species each of Diptera and Coleoptera, with resistanceratios ranging from 1.1 to >1000.

Concern about rapid adaptation of insects to δ-endotoxin-based transgenic plantshas led to a steady increase in the number of researchers performing laboratory se-lection experiments with single toxins and in some cases with the specific gene prod-ucts that insects will ingest when consuming the foliage from bioengineered plants(Estada & Ferré 1994). Researchers found that resistance ratios were consistentlyhigher, and resistance developed more rapidly, in insect populations selected with in-dividual toxins than in populations selected with Bt-insecticides that contain livespores and multi-toxin crystals, such as Bt subsp. kurstaki (Btk) (Tables 1 and 2). Theresults of these experiments suggest that deployment of the high dose, single toxinstrategy in the design of transgenic plants will quickly generate resistant populationsof the target pest.

The selection experiment reported by Moar et al. (1994) best illustrates the differ-ential response of Spodoptera exigua populations subjected to selection pressure fromeither Btk spore/crystal preparations or purified CryIC toxin (Table 2). After 20 gen-


erations, larvae selected with Btk spore/crystal preparations were only 3- to 4-fold re-sistant. However, the cohort selected with pure CryIC protoxin was 1000-foldresistant after 21 generations. This confirms that the complex mode of action, presentin most commercially Bt-based insecticides available today, is important in the lowprevalence of field resistance for pest populations treated with Bt. It may also explainthe lack of Bt-resistance generated in laboratory experiments which used crude or for-mulated Bt preparations containing both δ-endotoxins and spores for selections(Tabashnik 1994a). Before 1985, large quantities of a single, pure Cry toxin, madepossible through advances in molecular biology, were simply not available.

Although laboratory-generated resistance within a susceptible species may not re-flect the rates and mechanisms generated in field populations, laboratory populationsare invaluable tools for the study of potential risk of resistance, physiological and be-havioral mechanisms of resistance, cross-resistance, genetics, stability, fitness costs,and, ultimately, for the development of methods for monitoring, managing, and delay-ing resistance.

Heritability

It is apparent that the genetic capacity to evolve resistance to Bt δ-endotoxin iswidespread in insects. Although some efforts to select for resistance have failed, thismay reflect either insufficient selection pressure, a genetic bottleneck due to lack orloss of genetic diversity in the laboratory colony, or both (McGaughey & Whalon 1992,Whalon & McGaughey 1993). Selection experiments often provide the necessary in-formation to estimate heritability (h2), the proportion of the observed variability thatis caused by additive genetic variation (Falconer 1989). Estimated heritability wasused by Tabashnik (1992, 1994a) to estimate the ability of populations to develop re-sistance in 27 selection experiments. He showed that P. interpunctella has a relativelyhigh h2 compared to other moths. This reflects low phenotypic variation, perhaps re-sulting from its stable environment, and high additive genetic variation for the resis-tance trait, perhaps resulting from exposure to Bt in its environment.

Tabashnik (1994a) also discussed the potential usefulness and limitation of theheritability estimates in assessing resistance risk, i.e., predicting the rate at which apest will evolve resistance (Tabashnik 1992, Keiding 1986, Tabashnik & McGaughey1994). Plodia interpunctella has a high estimated h2 and adapts readily to Bt in thelaboratory, but high levels of field resistance are unknown. McGaughey (1985) sug-gested that the infrequent applications of Bt in grain bins, as well as Bt’s limited effi-cacy, helped preserve the susceptible individuals within the treated population,thereby generating only low levels of resistance.

In contrast, susceptible populations of P. xylostella with a comparatively low h2 donot achieve significant levels of resistance to Bt in laboratory selections (Devriendt &Martouret 1976, Krieg & Langenbruch 1981). However, moderately resistant fieldpopulations of P. xylostella quickly reach high levels of resistance during laboratoryselections (Tabashnik et al. 1991). This shows that selection is occurring in pest pop-ulations that are being intensively managed with Bt-based insecticides. Similar re-sults of rapid laboratory adaptation to Bt after intensive field exposure were found inpopulations of mosquitoes (Gill et al. 1992), P. interpunctella (McGaughey & Johnson1992), and Leptinotarsa decemlineata (Say) (Whalon et al. 1993) (Table 2).

Intraspecific Susceptibility

In most species, phenotypic variations in Bt tolerance have a strong genetic basis,and species with high variability in this trait will develop resistance more quickly un-


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der intensive selection pressure (Tabashnik 1994a). Baseline data on target pest sen-sitivity to Bt are essential in assessing the risk of resistance. Tabashnik (1994a)estimated variations in Bt-susceptibility among populations of 15 insect specieswithin three orders and found minimal variation except in three species of moths.Variation in two stored product moths (up to 42-fold) was not associated with Bt treat-ments, but presumably resulted from natural exposure to Bt in grain bins (Kinsinger& McGaughey 1979). Variation in the third moth, P. xylostella, however, was attribut-able to repeated exposure to Bt foliar sprays. Tabashnik et al. (1990) determined thatthe susceptibility of intensively treated populations varied up to 40-fold, whereas thesusceptibility for populations receiving minimal Bt exposure in the field and in labo-ratory colonies varied no more than 7-fold.

Increasing reliance on Btk-based products for the suppression of spruce budworm,Choristoneura fumiferana (Clemens), an important defoliator of coniferous forests inNorth America, stimulated interest in assessing the risk of field resistance (van Fran-kenhuyzen et al. 1995). Studies on the variation in Bt tolerance within and amongnine spruce budworm field populations from Ontario with no previous Bt exposure re-veal substantial familial differences in sensitivity to Bt, whereas differences betweenpopulations are minimal. This finding suggests that Bt tolerance is genetically based,and that spruce budworm has the potential to adapt to Bt. One laboratory selectionexperiment (Table 2) illustrates an increase in frequency of the resistance trait (vanFrankenhuyzen et al. 1995).

To gain baseline data on the Bt susceptibility of two pests targeted by transgeniccotton, Heliothis virescens (F.) and Helicoverpa zea (Boddie), Stone & Sims (1993) bio-assayed populations from 14 states with activated CryIA(c) toxin and a commercialBtk-based insecticide. Although previous exposure to Bt was not identified, significantdifferences within populations of both species were detected. Moreover, the variabilityin Bt tolerance was consistently higher for the activated toxin in both species. Thisagain suggests that an individual toxin as expressed in transgenic cotton would stim-ulate adaptation more rapidly than the commercial preparations that contain a mix-ture of spores and crystals. Stone & Sims (1993) acknowledged that monitoring andmanaging for resistance will be an exciting challenge when insect-resistant cotton, ex-pressing Bt δ-endotoxin, is deployed.

Mechanisms

An understanding of the mechanisms of resistance to Bt toxins will prove essentialin the future design and management of transgenic plants containing Bt toxin genes.At present, the primary mechanism of resistance reported for P. interpunctella (VanRie et al. 1990b) and P. xylostella (Ferré et al. 1991) is a reduction in the binding oftoxin to receptors on the midgut brush border membrane. This is the same mecha-nism known to account for much of the host specificity to various δ-endotoxins in Lep-idoptera (Hoffman et al. 1988b).

Plodia interpunctella, selected for resistance to Btk (see Table 1), shows a 50-foldreduction in midgut brushborder membrane receptor-binding affinity to CryIA(b)(Van Rie et al. 1990b). However, in vivo toxicity of CryIC (not present in Btk) in-creased when bioassayed in the Btk-resistant P. interpunctella. This increase in tox-icity resulted from an increase in the CryIC binding sites. These results show that atleast two distinct molecular changes occurred in the midgut receptor population in re-sponse to Bt selection. Although the role of these receptor molecules in the insect mid-gut are, as yet, poorly characterized, Van Rie et al. (1990b) suggest the increase in onereceptor population may compensate for diminished function in the other. In a strain


of P. interpunctella selected for resistance to Bt subsp. entomocidus (Bte), Oppert et al.(1994) reported reduced proteolytic activation of CryI(A)c.

Using similar methodologies, resistance mechanisms studies of Btk-resistant fieldpopulations of P. xylostella demonstrated greatly reduced or lack of toxin binding tomidgut receptors, suggesting a change or complete loss of the receptor (Ferré et al.1991, Tabashnik et al. 1994). Using a different method for quantifying receptor bind-ing, Masson et al. (1995) reported a loss of receptors in a strain of resistant P. xylos-tella, although not adequate to explain the high level of resistance.

The results from studies of other resistant insect species suggest that factors otherthan receptor binding can contribute to resistance. For example, two separate studiesof resistance in H. virescens found no relation between resistance to CryIA(b) orCryIA(c) and toxin-receptor binding (MacIntosh et al. 1991, Gould et al. 1992). In ad-dition, studies of Trichoplusia ni (Hübner) determined that CryIA(b) and CryIA(c)share the same receptor, but a strain of T. ni selected for resistance to CryIA(b)showed cross-resistance to CryIA(c) (Estada and Ferré 1994). Resistance mechanismsin these insects may involve changes in post-binding events such as channel forma-tion, leakage, and repair rate.

Cross-resistance

The apparent specificity, diversity, and genetic versatility of Bt δ-endotoxins sug-gest that resistance might be managed by deploying toxins in mixtures or sequences(Georghiou 1990, Stone et al. 1991, Van Rie 1991). Although at least 12 lepidopteran-active δ-endotoxins are available (Adang 1991), evidence is mounting that selectionfor resistance to one or more δ-endotoxins causes resistance to others (Tables 3 and 4).This phenomenon, known as cross-resistance, typically occurs when mechanisms oftoxicity are similar.

Many of the resistant insect populations generated in the laboratory and the fieldwere selected with Bt-based insecticides containing multiple toxins; the most studiedproducts are formulated with Btk strain HD-1, which is comprised of live spores anda mixture of five toxins (Table 1). In many studies, cross-resistance is referred to as anincrease in tolerance of a population, selected with one Bt isolate, to an isolate con-taining a different mixture of toxins. For example, efforts to find other Bt isolates tocontrol P. interpunctella resistant to Btk (140-fold) in grain bins showed that these in-sects were also resistant to 32 of the 57 Bt isolates assayed (McGaughey & Johnson1987). Resistance was highest among various Btk isolates, suggesting some degree ofspecificity. Many Bt isolates overlap considerably in their δ-endotoxin composition. Inthe context of the following discussion, cross-resistance is defined as an increase inthe tolerance of a population to a toxin absent in the preparation used for selection.Resistance, on the other hand, refers to increasing tolerance to a toxin that is presentin the Bt isolate used in selection.

Strains of P. interpunctella, selected for resistance to Btk, Bte, Bt subsp. aizawai(Bta) strain HD-112, Bta strain HD-133, or a mixture of Btk and Bta HD-133, showedsome level of resistance and cross-resistance to six δ-endotoxins tested (McGaughey &Johnson 1994) (Table 3). Btk tended to select for high levels of resistance to the entirecomplex of CryIA toxins which are 82 to 90% homologous in their amino acid se-quences (Höfte & Whitley 1989). Resistance was highest to CryIA(b) and CryIA(c).Evidence suggesting that they share the same binding site (Wolfersberger 1990) isalso supported by the high levels of cross-resistance to CryIA(c) in populations se-lected with Bta HD133 and Bte, which is likely derived from the presence of CryIA(b).The specificity of the target receptor appears to be greater for CryIA(c) than forCryIA(b) because cross-resistance is significantly greater than the selected resis-


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1 Res

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ted

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nt

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t is

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(se

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lon

y.3 R

esu

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augh

ey &

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1994

).4 n

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not

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le.

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ult

s co

mpi

led

from

Tab

ash

nik

et

al. (

1993

, 199

4b).

6 y, s

ign

ifica

nt

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l of

resi

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ce, b

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o w

as n

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epor

ted.


tance. Cross-resistance to CryIB and CryIC was significant, although very low, whichis consistent with their relatively low amino acid sequence similarity to the CryIA tox-ins (ranging from 55 to 67%) (Tabashnik et al. 1994b). As expected, Bt isolates con-taining more dissimilar toxins, such as found in Bta and Bte, selected for a broaderspectrum of cross-resistance in the treated insects than did isolates producing similarCry toxins such as Btk HD-1. The complexity in the pattern of resistance results, inpart, from differential selective pressure exerted by different levels of toxicity and theamounts of each toxin in each Bt strain. Overall, the results of this selection experi-ment clearly contradict the claim that toxin mixtures will prevent or retard the devel-opment of resistance (Tabashnik & McGaughey 1994).

Field-resistant P. xylostella from Hawaii, further selected with Btk in the labora-tory, show a similar pattern of broad, but highly variable, resistance and cross-resis-tance among the Cry toxins tested (Tabashnik et al. 1993, 1994b) (Table 3). Again,resistance was highest to the CryIA toxins, and cross-resistance to CryIB and CryICwas low. Intermediate cross-resistance to CryIF is supported by its amino acid se-quence homology (70 to 72% to the CryIA toxins).

A different field population of P. xylostella from the Philippines, also resistant toCryIA(b) (resistance ratio 236), showed no resistance to Btk, CryIA(a), CryIA(c) andno cross-resistance to CryIB, and CryIC (Ferré et al. 1991, Ballester et al., in press).These researchers determined that CryIA(b) has a single binding site that is also rec-ognized by CryIA(a) and CryIA(c). Loss or modification of the CryIA(b) binding site re-sults in the loss of its toxicity. Ballester et al. (in press) noted that this narrowspectrum of resistance is somewhat unique and suggested that this population mayrepresent a biotype present in the Phillipines, not a case of field selection by Btk.

To understand the complexity of these emerging cross-resistance patterns, wemust select insect populations with individual toxins. In the case of most Bt isolates,which produce several toxins, this requires cloning and transferring the cry gene intoan acrystalliferous strain of Bt, or into another species of bacteria, such as Escheri-chia coli or Pseudomonas fluorescens. In some studies, the gene is modified to simu-late the specific Cry products being expressed in transgenic plants or microorganisms.

The first such study that also included cross-resistance data was the selection ofH. virescens with purified CryIA(c) (Gould et al. 1992) (Table 4). After 17 generations,the population was 50-fold resistant to CryIA(c) and, as reported for other lepidopter-ans, was 13-fold cross-resistant to CryIA(b). This population, however, was also cross-resistant to CryIIA, CryIB, and CryIC. In a subsequent paper, Gould et al. (in press)also reported high levels of cross-resistance to CryIF. No measurable differences inthe concentration of CryIA(c), or CryIA(b) binding sites or binding affinities, were de-tected between selected and unselected H. virescens, suggesting other mechanisms ofresistance are involved. Similar broad-spectrum cross-resistance was also reportedfor two species of Spodoptera and two coleopterans, Chrysomela scripta F. (Table 4)and L. decemlineata (Whalon, unpublished results).

In contrast, cross-resistance in Trichoplusia ni (Hübner) selected with CryIA(b)had a higher degree of specificity within the CryIA group of toxins (Estada & Ferre1994). This specificity was similar to that reported for P. xylostella from the Phil-lipines (Ballester et al. in press), although no data were presented on unrelated toxinsfor T. ni. As determined for several other lepidopterans, receptor-binding assays re-veal that CryIA(b) and CryIA(c) share the same high-affinity binding sites, and resis-tance to one would imply cross-resistance to the other. However, no cross-resistanceto CryIA(c) was detected, suggesting CryIA(c) toxicity results from other binding sitesor alternative mechanisms.

Overall, cross-resistance patterns and their underlying physiological mechanismare very complex and somewhat unpredictable, even within a closely related group of


TA

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toxins and susceptible insects. A more complete understanding of how each toxin in-teracts within a particular target species at the molecular level is critical to selectingδ-endotoxins for design of transgenic plants that do not favor broad-spectrum cross-resistance.

Adaptation to δ-Endotoxin

The recent appearance of resistance to conventional Bt-based insecticides was cor-related with improved formulation and more intensive usage patterns (Tabashnik etal. 1990). In addition to operational uses of the Bt toxins, pest genetics, behavior,physiology, and ecology are critical factors in predicting resistance risk.

The best studied examples of Bt-resistance, P. interpunctella and P. xylostella,share many attributes that leave their populations particularly vulnerable to rapidselection (Table 5). These attributes include multivoltinism, short generation time,and populations that tend to be isolated and stable. These attributes are typical ofmany agricultural and medical pests, and they temper enthusiasm for bioengineeredplants which produce continuous and high levels of δ-endotoxin. It is apparent, how-ever, that strategies are needed to delay or avoid resistance in pests that are inten-sively managed with Bt, regardless of the deployment method.

Inheritance

The development of strategies to manage resistance requires some understandingof the inheritance of a resistance trait in the pest population (Gould 1986). This wasdemonstrated by Tabashnik (1994b) using a population genetics model to simulatethe response of a pest population to different resistance management strategies. Thebest strategy, i.e., one that delays resistance the longest, must be customized to thenumber of alleles and inheritance of the trait within the population.

Studies of the genetics of resistance typically involve determining the susceptibil-ity of progeny from crosses between individuals from the selected and unselected pop-

TABLE 5. FACTORS ASSOCIATED WITH DEVELOPMENT OF RESISTANCE TO BT IN TWOLEPIDOPTERANS.

Factors P. interpunctella P. xylostella

Pest Attributes and Management

Short generation yes yesGenerations/year 5-6 8-10Population isolation yes yesRealized heritability high lowCrop rotation no no

Operational Use of Bt

Multiple toxins yes yesApplications/generation 5-6 5-8Frequency/generation 2-4 days 1.5-2 daysSelection pressure high high


ulations. In both P. interpunctella and P. xylostella, resistance is autosomallyinherited (no maternal effects or sex linkage), partially recessive (progeny suscepti-bility more similar to unselected parent), and apparently due to one or a few majorloci (McGaughey 1985, McGaughey & Beeman 1988, Hama et al. 1992, Tabashnik etal. 1992). The genetic basis of resistance to the CryIA toxin-complex in H. virescens ispartially recessive and due to a single locus or set of tightly linked loci (Gould et al. inpress). In L. decemlineata, resistance to CryIIIA is also autosomally inherited, andconferred by one incompletely dominant gene (Rahardja & Whalon 1995). Knowledgeof the genetic basis of resistance is critical to our understanding of the stability of thetrait within the selected population.

Stability

Perhaps one of the simplest resistance management strategies involves providingthe pest population with intermittent time periods in which Bt is not used for control.The success of such temporal refuges is dependent on the stability of the resistancetrait after Bt exposure ceases. The rate of reversion is dependent on the inheritanceof the resistance trait and the fitness costs associated with resistance.

Different populations of resistant P. xylostella revert at different rates when selec-tion with Bt is relaxed. Typically, the decline is slow and incomplete; for example, oneBtk-resistant population declined from 29-fold to 1-fold after 32 generations withoutselection (Tabashnik et al. 1991). Similar results were reported in other Btk-resistantpopulations of P. xylostella (Tabashnik et al. 1991, 1994, Hamma et al. 1992) and inother resistant species, including P. interpunctella (McGaughey & Beeman 1988), H.virescens (Sims & Stone 1991), and L. decemlineata (Rahardja & Whalon 1995). In arecent study, rapid and complete reversal of resistance occurred after 13 generationswithout selection in a population of P. xylostella with 2800-fold resistance to Btk(Tabashnik et al. 1994a). This result suggests that resistance is achieved with signif-icant loss in fitness, perhaps related to the alteration in the midgut binding sites doc-umented in the study. Groeters et al. (1993, 1994) quantified reduced egg hatch,survival to the adult stage, fecundity, and mating success in male moths as significantfitness costs of resistance. Despite the rapid reversion to the susceptible genotype, thepopulation responded rapidly to reselection (Tabashnik et al. 1994a). Rapid resur-gence of resistance in relaxed populations is typical, indicating the persistence of alow number of highly resistant individuals. If the alleles for resistance become fixedin the population, or other alleles compensate for losses in fitness, resistance becomesstable and reversion to susceptibility is unlikely.

RESISTANCE MANAGEMENT

In an effort to preserve the utility of these unique insecticidal proteins, knowledgeof resistance management to conventional pesticides is useful (Gould 1988a, 1988b,Stone et al. 1991, McGaughey & Whalon 1992, Whalon & McGaughey 1993, Mc-Gaughey 1994, Tabashnik 1994a). Unfortunately, selection for the resistance trait ina pest population is probably the inevitable consequence of insecticide use (Denholm& Rowland 1992). The goal then becomes how to design and manipulate operationalstrategies that best conserve susceptibility, thereby delaying resistance. The imple-mentation of integrated pest management (IPM) strategies that optimize the goals ofresistance management involves 1) diversifying the sources of mortality to avoid se-lection for a single mechanism, 2) reducing selection pressure for the major mortalityfactors, 3) maintaining susceptible individuals by providing refuges and encouragingimmigration, 4) monitoring for increasing resistance to any one of the mortalityagents, and 5) responding to resistance through management strategies designed to


reduce the frequency of the resistance trait (Whalon & McGaughey 1993). Unfortu-nately, IPM is rarely implemented before a resistance crisis occurs, and generally an-other insecticide is available to replace the old one.

Conventionally applied Bt-based insecticides are more amenable to such IPMstrategies, because the short residual time and host specificity help reduce selectionpressure associated with the Bt toxins. Insects surviving Bt exposure are generally ina weakened condition, facilitating their exploitation by other mortality factors such asbeneficial insects (Tabashnik 1986, 1994a) and pathogens (Krieg 1971, Jacques &Morris 1981). Other stressing agents, such as adverse weather conditions and lowplant nutritional quality, will also cause higher mortality in insects recovering fromBt exposure. The cumulative suppression exerted by these factors also reduces selec-tion pressure by reducing the frequency of pesticide applications. However, recent im-provements in Bt formulations involve increasing toxicity, increasing residual times,and in some cases broadening host range, thereby bringing selection pressure for re-sistance more in line with conventional insecticides (Tabashnik 1994a).

The expression of δ-endotoxins in transgenic plants is considered analogous, insome respects, to intrinsic host plant defenses selected for by classical plant breeders(Gould 1988a). Unfortunately, pests adapt to resistant cultivars, and without appro-priate deployment strategies, they are rendered ineffective (Gould 1986, Cox &Hatchett 1986). At present, bioengineering insect-resistant plants involves the incor-poration of Bt δ-endotoxin genes into plants. There was considerable optimism severalyears ago that these plants would remain durable (Gould 1986,Gould 1988b, Rousch1989).

Early optimism and excitement over genetically-engineered plants expressing δ-endotoxin, perhaps bolstered by the presumption of an unlimited variety of these pro-teins, have given way to serious concern over the durability of these plants, and withit, conventional uses of Bt. Resistance management strategies in the context of Btwere recently reviewed in some detail by McGaughey & Whalon (1992), Whalon &McGaughey (1993), Robison et al. (1994), and Tabashnik (1994a). These strategies in-clude 1) mixtures of toxins with different mechanisms, either within the same plantor in different plants, or expressed serially over time (Gould 1986); 2) synergists toincrease toxicity (MacIntosh et al. 1990); 3) rotations to alternative toxins tempo-rally to reduce the frequency of resistant individuals (Tabashnik 1989); 4) refuges,temporal and spatial, to facilitate survival of susceptible individuals (Gould & Ander-son 1991); 5) low doses of toxin that produce sublethal effects, such as reduced fe-cundity and slowed development, favoring other mortality factors; 6) ultrahighdoses of toxin that kill resistant heterozygotes and homozygotes (Denholm & Row-land 1992, Tabashnik 1994a); and 7) gene regulation of toxin titre, location, and in-duction (Whalon & McGaughey 1993).

Tabashnik (1994b) used a population genetics model (Mallet & Porter 1992) tosimulate the effect of several resistance management strategies on resistance (singlelocus with two alleles) in a pest such as Heliothis. Transgenic plants expressing δ-en-dotoxin were planted as 1) pure stands of toxic plants, 2) seed mixtures with varyingproportions of toxic and toxin-free plants, 3) toxin-free plants in refugia, and 4) seedmixtures + refugia. Across a range of conditions, seed mixtures always delayed the on-set of insect resistance to the toxin, when compared to pure stands of toxic plants. Ref-ugia will delay resistance as long, or in some cases longer, than seed mixtures becauserefugia reduce selection without altering dominance (Mallet & Porter 1992). Refugiawill delay resistance longer than mixtures + refugia only under specific conditions.However, the implementation of pest control within refugia limits their ability to de-lay resistance in proportion to the efficacy of the controls.


The distinction between seed mixtures and refugia is the spatial distribution oftoxin-free plants relative to the dispersal capability of pest larvae. Due to close prox-imity, larvae may move easily between toxic and toxin-free plants in fields plantedwith a seed mixture, whereas movement from toxic stands to toxic-free refugia is lesslikely. The tissue-specific expression of toxin through gene regulation is analogous toseed mixtures, because larvae can move freely between toxic and toxin-free tissue(Gould 1988a). Tabashnik (1994b) recommends maximizing spatial refuges, as well astemporal refuges, such as alternative crops and controls. The only method known toprolong the efficacy of any insecticide is to minimize the exposure of the target peststo the toxin in space and time (Denholm & Rowland 1992). Resistance managementtactics must be validated in the field, under the inevitable practical, economic, and po-litical constraints imposed by agriculture today (May 1993).

CONCLUSIONS

The genetic capacity of insect populations to evolve resistance to Bt δ-endotoxinsis now well documented in many species within eight different insect orders. Althoughhigh-level field resistance is known only in P. xylostella, much has been learned fromstudying the many insect populations selected for resistance to Bt in the laboratory.We now know that 1) Bt resistance alleles are present at varying levels indifferent insect species and populations, 2) within a single species, the genetics,mechanisms, level, and stability of resistance vary between selected populations, 3)selection with a blend of toxins can select for resistance to each toxin in the blend, 4)resistance occurs more rapidly with purified toxins than with spore/crystal prepara-tions, 5) cross-resistance to δ-endotoxins is almost ubiquitous and often unpredict-able, and 6) reselection of revertant populations is rapid.

Today, Bt-based insecticides are frequently used in intensive agriculture, either inconjunction with conventional insecticides as a backup for control failure, or, as a lastresort once resistance to other registered insecticides has occurred. Many insect pests,therefore, are already adapted to mixtures of δ-endotoxins. It is probable that the de-ployment of transgenic plants will precede the development of resistance manage-ment strategies, because advances in resistance management technology have notkept pace with those made in biotechnology. Developing and validating realistic resis-tance management plans that preserve the durability of δ-endotoxins deployed intransgenic plants may prove far more complex than the theory and techniques thatactually generated the plants. Unfortunately, experimentally developed tactics to de-lay resistance have often been too naive or unrealistic for large-scale field implemen-tation (Hoy 1995).

Fortunately, many researchers with experience and knowledge of resistance man-agement with conventional insecticides have shifted the emphasis of their researchinto the development of strategies designed to prolong the durability of these uniquebacterial toxins in transgenic plants. In general, resistance management seeks tominimize the exposure of the target pest to a toxin in time and space. This can be ac-complished by developing IPM plans for these crops that include synergists, seed mix-tures and refugia, tissue-specific and inducible toxin expression, and alternatingcrops or control measures (Hoy 1995). Substantial benefits to the environment will begained if insects can be successfully managed through the careful deployment of ge-netically-engineered insect-resistant plants. However, only few Bt δ-endotoxins, allwith similar modes of action, are now available to plant molecular biologists. Deploy-ment of these plants before the management tactics are validated will result in theloss of Bt-based insecticides for many of the pests they are targeted to control.


ACKNOWLEDGMENTS

An overview of this paper was presented in a symposium entitled “The Myths ofManaging Resistance” at the Florida Entomological Society held 8-11 August 1994 inStuart, Florida. I am grateful to M. Hoy for inviting me to participate in that sympo-sium. I wish to thank M. Adang, N. Dubois, F. Gould, W. McGaughey, W. Moar, K. vanFrankenhuyzen, B. Tabashnik, and M. Whalon for their help with preprints and var-ious discussion, and D. Bradley for providing CryIB for laboratory bioassay. I am ap-preciative of the critical editorial comments provided by R. Haack, M. Hoy, N. Koller,and M. Whalon. I also want to give special acknowledgment to the excellent technicalsupport provided by D. Miller in my laboratory with selection and bioassay of Chry-somela scripta.

REFERENCES CITED

ADANG, M. J. 1991. Bacillus thuringiensis insecticidal crystal proteins: gene struc-ture, action, and utilization, pp. 3-24 in K. Maramorosch [ed], Biotechnology forbiological control of pests and vectors. Boca Raton, Florida:CRC Press.

ALY, C., M. S. MULLA, AND B. A. FEDERICI. 1985. Sporulation and toxin production byBacillus thuringiensis var. israelensis in cadavers of mosquito larvae (Diptera:Culicidae). J. Invertebr. Pathol. 46: 251-258.

ANGUS, T. A. 1954. A bacterial toxin paralyzing silkworm larvae. Nature (London)173: 545-546.

ARONSON, A. I. 1993. Insecticidal toxins, pp. 953-963 in A. L. Sonenshein, J. A. Hoch,and R. Losick [eds], Bacillus subtilus and other gram-positive bacteria: bio-chemistry, physiology, and molecular biology. Washington, DC:American Soci-ety for Microbiology.

ARONSON, A. I., E. S. HAN, W. MCGAUGHEY, AND D. JOHNSON. 1991. The solubility ofinclusion proteins from Bacillus thuringiensis is dependent upon protoxin com-position and is a factor in toxicity to insects. Appl. Environ. Microbiol. 57:981-986.

BALLESTER, V., B. ESCRICHE, J. L. MÉNSUA, G. W. RIETHMACHER, AND JUAN FERRÉ.Lack of cross-resistance to other Bacillus thuringiensis crystal proteins in apopulation of Plutella xylostella highly resistant to CryIA(b). Biocontrol Sci.Technol. In Press.

BAUER, L. S., AND H. S. PANKRATZ. 1992. Ultrastructural effects of Bacillus thuring-iensis var san diego on midgut cells of the cottonwood leaf beetle. J. Invertebr.Pathol. 60: 15-25.

BAUER, L. S., C. N. KOLLER, D. L. MILLER, AND R. M. HOLLINGWORTH. 1994. Labora-tory selection of the cottonwood leaf beetles, Chrysomela scripta, for resistanceto Bacillus thuringiensis var. tenebrionis δ-endotoxin. p. 68 in Abstracts VIth In-tern. Coll. Invertebr. Pathol.

BECKER, N., AND J. MARGOLIT. 1993. Use of Bacillus thuringiensis israeliensis againstmosquitoes, pp. 147-170 in P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs[eds], Bacillus thuringiensis, an environmental biopesticide: theory and prac-tice. New York:Wiley.

BERLINER, E. 1911. Über die Schlaffsucht der Mehlmottenraupe (Ephestia kuehniellaZell), und ihren Erreger Bacillus thuringiensis n. sp. Zeitschrift für an-gewandtes. Entomol. 2: 29-56.

BOMAN, H. G. 1981. Insect Responses to Microbial Infections, pp. 769-784 in H. D.Burges and N. W. Hussey [eds], Microbial control of pests and plant diseases1970-1980. New York:Academic Press.

BORGONIE, G., R. VAN DRIESSCHE, F. LEYNS, G. ARNAUT, D. DE WAELE, AND A.COOMANS. 1995. Germination of Bacillus thuringiensis spores in bacteriopha-gous nematodes (Nematoda: Rhabditida). J. Invertebr. Pathol. 65: 61-67.


BRADLEY, D., M. A. HARKEY, M. K. KIM, K. D. BIEVER, AND L. S. BAUER. 1995. Insec-ticidal CryIB crystal protein of Bacillus thuringiensis has dual specificity to co-leopteran and lepidopteran larvae. J. Invertebr. Pathol. 65: 162-173.

BRAVO, A., S. JANSENS, AND M. PEFEROEN. 1992. Immunocytochemical localization ofBacillus thuringiensis insecticidal crystal proteins in intoxicated insects. J. In-vertebr. Pathol. 60: 237-246.

BREWER, G. J. 1991. Resistance to Bacillus thuringiensis subsp. kurstaki in the sun-flower moth (Lepidoptera: Pyralidae). Environ. Entomol. 20:316-322.

BRIESE, G. J. 1981. Resistance of insect species to microbial pathogens, pp. 511-545 inE. W. Davidson [ed], Pathogenesis of Invertebrate Microbial Diseases. Totowa,New Jersey:Allanheld Osmun.

BURGES, H. D. 1971. Possibilities for pest resistance to microbial control agents, pp.445-457 in H. D. Burges and N. W. Hussey [eds], Microbial Control of Insectsand Mites. New York:Academic Press.

BURGES, H. D., AND J. A. HURST. 1977. Ecology of Bacillus thuringiensis in storagemoths. J. Invertebr. Pathol. 30: 131-139.

CARLTON, C. C., AND C. GAWRON-BURKE. 1993. Genetic improvement of Bacillus thu-ringiensis, pp. 43-61 in L. Kim [ed], Advanced Engineered Pesticides. NewYork:Marcel Dekker.

CARROLL, J., AND D. J. ELLAR. 1993. Analysis of Bacillus thuringiensis δ-endotoxin ac-tion on insect-membrane permeability using a light-scattering assay. Eur. J.Biochem. 214: 771-778.

CARROLL, J., J. LI, AND D. J. ELLAR. 1989. Proteolytic processing of acoleopteran-specific delta-endotoxin produced by Bacillus thuringiensis var.tenebrionis. Biochem. J. 261:99-105.

CHILCOTT, C. N., B. H. KNOWLES, D. J. ELLAR, AND F. A. DROBNIEWSKI. 1990. Mech-anism of action of Bacillus thuringiensis parasporal body, pp. 44-65 in H. deBarjac and D. Sutherland [eds], Bacterial Control of Mosquitoes and Blackflies:Biochemistry, Genetics, and Applications of Bacillus thuringiensis and Bacillussphaericus. New Brunswick:Rutgers University Press.

COX, T. S., AND J. H. HATCHETT. 1986. Genetic model for wheat/Hessian fly (Diptera:Cecidomyiidae) interaction: strategies for deployment of resistance genes inwheat cultivars. Environ. Entomol. 15: 24-31.

CUMMINGS, C. E., AND D. J. ELLAR. 1994. Chemical modification of Bacillus thuring-iensis activated δ-endotoxin and its effect on toxicity and binding to Manducasexta midgut membranes. Microbiol. 140: 2737-2747.

DE BARJAC, H. 1987. Operational bacterial insecticides and their potential for futureimprovements, pp. 63-73 in K. Maramorosch [ed], Biotechnology in Inverte-brate Pathology and Cell Culture. San Diego, California:Academic Press.

DENHOLM, I., AND M. W. ROWLAND. 1992. Tactics for managing pesticide resistance inarthropods: theory and practice. Annu. Rev. Entomol. 37: 91-112.

DENOLF, P., S, JANSENS, M. PEFEROEN, D. DEGHEELE, AND J. VAN RIE. 1993. Two dif-ferent Bacillus thuringiensis delta-endotoxin receptors in the midgut brushborder membrane of the european corn borer, Ostrinia nubilalis (Hübner) (Lep-idoptera: Pyralidae). Appl. Environ. Microbiol. 59: 1828-1837.

DEVRIENDT, M., AND D. MARTOURET. 1976. Absence de résistance à Bacillus thuring-iensis chez la teigne des crucifères, Plutella maculipennis. Entomophaga 21:189-199.

DU, C., P. A. MARTIN, AND K. W. NICKERSON. 1994. Comparison of disulfide contentsand solubility at alkaline pH of insecticidal and noninsecticidal Bacillus thur-ingiensis protein crystals. Appl. Environ. Microbiol. 60: 3847-3853.

ELLAR, D. J. 1994. Structure and mechanism of action of Bacillus thuringiensis endo-toxins and their receptors. Biocon. Sci. Technol. 4: 445-447.

ELY, S. 1993. The engineering of plants to express Bacillus thuringiensis δ-endotox-ins, pp. 105-124 in P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs [eds],Bacillus thuringiensis, an environmental biopesticide: theory and practice.New York:Wiley, 311 pp.


ENGLISH, L., AND S. L. SLATIN. 1992. Mode of action of delta-endotoxin from Bacillusthuringiensis: a comparison with other bacterial toxins. Insect Biochem. Molec.Biol. 22: 1-7.

ESCRICHE, B., A. C. MARTINEZ-RAMIREZ, M. D. REAL, F. J. SILVA, AND J. FERRÉ. 1994.Occurrence of three different binding sites for Bacillus thuringiensis δ-endot-oxins in the midgut brush border membrane of potato tuber moth, Phthorimaeaoperculella (Zeller). Arch. Insect Biochem. Physiol. 26: 315-327.

ESTADA, U., AND J. FERRÉ. 1994. Binding of insecticidal crystal proteins of Bacillusthuringiensis to the midgut brush border of the cabbage looper, Trichoplusia ni(Hübner) (Lepidoptera: Noctuidae), and selection for resistance to one of thecrystal proteins. Appl. Environ. Microbiol. 60: 3840-3846.

FALCONER, D. S. 1989. Introduction to Quantitative Genetics. New York:Longman.FEDERICI, B. A. 1993. Insecticidal bacterial proteins identify the midgut epithelium as

a source of novel target sites for insect control. Arch. Insect Biochem. Physiol.22: 357-371.

FERRÉ, J., M. D. REAL, J. VAN RIE, S. JANSENS, AND M. PERFEROEN. 1991. Resistanceto the Bacillus thuringiensis bioinsecticide in a field population of Plutella xy-lostella is due to a change in a midgut membrane receptor. Proc. Natl. Acad. Sci.USA 88: 5119-5123.

FEITELSON, J. S. 1993. The Bacillis thuringiensis family tree, pp. 63-71 in L. Kim[ed.], Advanced Engeered Pesticides, New York: Marcel Dekker.

FEITELSON, J. S., J. PAYNE, AND L. KIM. 1992. Bacillus thuringiensis: insects and be-yond. Bio/technology 10: 271-276.

GARCZYNSKI, S. F., AND M. J. ADANG. Bacillus thuringiensis CryIA(c) δ-endotoxinbinding aminopeptidase in the Manduca sexta midgut has a glycosyl-phos-phatidylinositol anchor. Insect Biochem. Molec. Biol. In Press.

GARCZYNSKI, S. F., J. W. CRIM, AND M. J. ADANG. 1991. Identification of putative in-sect brush border membrane-binding molecules specific to Bacillus thuringien-sis δ-endotoxin by protein blot analysis. Appl. Environ. Microbiol. 57: 2816-2820.

GASSER, C. S., AND R. T. FRALEY. 1989. Genetically engineering plants for crop im-provement. Science 244: 1293-1299.

GELERNTER, W., AND G. E. SCHWAB. 1993. Transgenic bacteria, viruses, algae andother microorganisms as Bacillus thuringiensis delivery systems, pp. 89-104 inP. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs [eds], Bacillus thuringien-sis, an environmental biopesticide: theory and practice. New York:Wiley, 311pp.

GEORGHIOU, G. P. 1990. Resistance potential to biopesticides and consideration ofcountermeasures, pp. 409-420 in J. E. Casida [ed], Pesticides and Alternatives.New York:Elsevier.

GEORGHIOU, G. P. 1994. Mechanisms and microbial characteristics of invertebrate re-sistance to bacterial toxins, pp. 48-50. in Proc. VIth Intern. Coll. Invertebr.Pathol., Montpellier, France:Soc. Invertebr. Pathol.

GEORGHIOU, G. P., AND A. LAGUNES. 1988. The occurrence of resistance to pesticides:cases of resistance reported worldwide through 1988. Food and AgriculturalOrganization, Rome, 325 pp.

GILL, S. S., E. A. COWLES, AND P. V. PIETRANTONIO. 1992. The mode of action of Ba-cillus thuringiensis endotoxins. Annu. Rev. Entomol. 37: 615-36.

GOLDBERG, L. J., AND J. MARGALIT. 1977. A bacterial spore demonstrating rapid lar-vicidal activity against Anopheles serengetii, Uranotaenia unguiculata, Culexunivittatus, Aedes aegypti and Culex pipiens. Mosq. News. 37: 355-358.

GOLDMAN, I. F., J. ARNOLD, AND B. C. CARLTON. 1986. Selection for resistance to Ba-cillus thuringiensis subspecies israelensis in field and laboratory populations ofthe mosquito Aedes aegypti. J. Invertebr. Pathol. 47: 317-324.

GOULD, F. 1986. Simulation models for predicting durability of insect-resistant germ-plasm: Hessian fly (Diptera: Cecidomyiidae) resistant winter wheat. Environ.Entomol. 15: 1-10.


GOULD, F. 1988a. Evolutionary biology and genetically engineered crops: Consider-ation of evolutionary theory can aid in crop design. BioScience 38: 26-33.

GOULD, F. 1988b. Genetic engineering, integrated pest management and the evolu-tion of pests, pp. 15-18 in Planned Release of Genetically Engineered Organ-isms. Cambridge (UK):Elsevier.

GOULD, F., AND A. ANDERSON. 1991. Effects of Bacillus thuringiensis and HD-73delta-endotoxin on growth, behavior, and fitness of susceptible andtoxin-adapted strains Heliothis virescens (Lepidoptera: Noctuidae). Environ.Entomol. 20: 30-38.

GOULD, F., A. ANDERSON, D. LANDIS, AND H. VAN MELLAERT. 1991. Feeding behaviorand growth of Heliothis virescens larvae on diets containing Bacillus thuring-iensis formulations or endotoxins. Entomol. Exp. Appl. 58: 199-210.

GOULD, F., A. ANDERSON, A. REYNOLDS, L. BUMGARNER, AND W. MOAR. Selection andgenetic analysis of a Heliothis virescens (Lepidoptera: Noctuidae) strain withhigh levels of resistance to some Bacillus thuringiensis toxins. J. Econ. Ento-mol. In Press.

GOULD, F., A. MARTINEZ-RAMIREZ, A. ANDERSON, J. FERRE, F. J. SILVA, AND W. J.MOAR. 1992. Broad-spectrum resistance to Bacillus thuringiensis toxins in He-liothis virescens. Proc. Natl. Acad. Sci. 88: 7986-7990.

GROETERS, F. R., B. E. TABASHNIK, N. FINSON, AND M. W. JOHNSON. 1993. Resistanceto Bacillus thuringiensis affects mating success of the diamondback moth (Lep-idoptera: Plutellidae). J. Econ. Entomol. 86: 1035-1039.

GROETERS, F. R., B. E. TABASHNIK, N. FINSON, AND M. W. JOHNSON. 1994. Fitnesscosts of resistance to Bacillus thuringiensis in the diamondback moth (Plutellaxylostella). Evol. 48: 197-201.

HAMA, H., K. SUZUKI, AND H. TANAKA. 1992. Inheritance and stability of resistance toBacillus thuringiensis formulations of the diamondback moth, Plutella xylos-tella L. (Lepidoptera: Yponomeutidae). Appl. Entomol. Zool. 27: 355-362.

HEIMPEL, A. M., AND T. A. ANGUS. 1959. The site of action of crystalliferous bacteriain Lepidoptera larvae. J. Insect Pathol. 1: 152-170.

HOFFMAN, C., P. LUTHY, R. HUTTER, AND V. PLESKA. 1988a. Binding of the delta-en-dotoxin from Bacillus thuringiensis to brush border membrane vesicles of thecabbage butterfly (Pieris brassicae). Eur. J. Biochem. 173:85-91.

HOFFMAN, C., H. VANDERBRUGGEN, H. HÖFTE, J. VAN RIE, AND S. JANSENS. 1988b.Specificity of Bacillus thuringiensis delta-endotoxin is correlated with the pres-ence of high-affinity binding sites in the brush border membrane of target in-sect midguts. Proc. Natl. Acad. Sci. USA 85: 7844-7848.

HÖFTE, H., AND H. R. WHITELEY. 1989. Insecticidal crystal proteins of Bacillus thur-ingiensis. Microbiol. Rev. 53: 242-255.

HONÉE, G., AND B. VISSER. 1993. The mode of action of Bacillus thuringiensis crystalprotein. Entomol. exp. appl. 69: 145-155.

HOY, M. A. 1995. Multitactic resistance management: an approach that is long over-due? Florida Entomol. 78:443-451.

ISHIWATA, S. 1901. On a kind of severe flasherie (sotto disease). Dainihan SanbshiKaiho 9: 1-5.

JAQUES, R. P., AND O. N. MORRIS. 1981. Compatibility of pathogens with other meth-ods of pest control and with different crops, pp. 695-715 in H. D. Burges [ed],Microbial control of pests and plant diseases 1970-1980. New York:AcademicPress.

JANSSON, R. K., AND S. H. LECRONE. 1990. Management of diamondback moth, Plu-tella xylostella, with nonconventional chemical and biological insecticides.Proc. Florida State Hortic. Soc. 103: 122-126.

KEIDING, J. 1986. Prediction of resistance risk assessment, pp. 279-297 in NationalResearch Council. Pesticide Resistance: Strategies and Tactics for Manage-ment. Washington, DC:National Academy.

KELLER, B., AND G. LANGENBRUCH. 1993. Control of coleopteran pests by Bacillus thu-ringiensis and resistance management, pp. 171-191 in P. F. Entwistle, J. S.


Cory, M. J. Bailey, and S. Higgs [eds], Bacillus thuringiensis, an environmentalbiopesticide: theory and practice. New York:Wiley.

KINSINGER, R. A., AND W. H. MCGAUGHEY. 1979. Susceptibility of populations of In-dianmeal moth and almond moth to Bacillus thuringiensis isolates (Lepi-doptera: Pyralidae). J. Econ. Entomol. 72: 346-349.

KIRSCH, K., AND H. SCHMUTTERER. 1988. Low efficacy of a Bacillus thuringiensis(Berl.) formulation in controlling the diamondback moth, Plutella xylostella(L.), in the Philippines. J. Appl. Entomol. 105: 249-255.

KNIGHT, P. J. K., N. CRICKMORE, AND D. J. ELLAR. 1994. The receptor for Bacillus thu-ringiensis CryIA(c) delta-endotoxin in the brush border membrane of the lepi-dopteran Manduca sexta is aminopeptidase N. Mol. Microbiol. 11: 429-436.

KNOWLES, B. H., AND J. A. T. DOW. 1993. The crystal delta-endotoxins of Bacillus thu-ringiensis: models for their mechanism of action on the insect gut. Bioessays15: 469-475.

KNOWLES, B. H., AND D. J. ELLAR. 1987. Colloid-osmotic lysis is a general feature ofthe mechanism of action of Bacillus thuringiensis delta-endotoxin with differ-ent insect specificity. Biochem. Biophys. Acta 924: 509-518.

KOLLER, C. N., L. S. BAUER, AND R. M. HOLLINGWORTH. 1992. Characterization of thepH-mediated solubility of Bacillus thuringiensis var. san diego native delta-en-dotoxin crystals. Biochem. Biophys. Res. Comm. 184:692-699.

KONI, P. A., AND D. J. ELLAR. 1994. Biochemical characterization of Bacillus thuring-iensis cytolytic δ-endotoxin. Microbiol. 140: 1869-1880.

KRIEG, A. 1971. Interactions between pathogens, pp. 459-468 in H. D. Burges and N.W. Hussey [eds], Microbial control of insects and mites. New York:AcademicPress.

KRIEG, A., AND G. A. LANGENBRUCH. 1981. Susceptibility of arthropod species to Ba-cillus thuringiensis, pp. 837-898 in H. D. Burges and N. W. Hussey [eds], Mi-crobial Control of Pests and Plant Diseases 1970-1980. New York: AcademicPress.

KRIEG, A., A. M. HUGER, G. A. LANGENBROOK, AND W. SCHNETTER. 1983. Bacillus thu-ringiensis var. tenebrionis: ein neuer gegenuber larven von Coleopteren wirk-samer. Pathotyp. Z. Ang. Entomol. 96: 500-508.

LAMBERT, B., AND M. PEFEROEN. 1992. Insecticidal promise of Bacillus thuringiensis.Bioscience 42: 112-122.

LI. J., J. CARROLL, AND D. J. ELLAR. 1991. Crystal structure of insecticidaldelta-endotoxin from Bacillus thuringiensis at 2.5 Å resolution. Nature 353:815-821.

MACINTOSH, S. C., G. M. KISHORE, F. J. PERLAK, P. G. MARRONE, AND T. B. STONE.1990. Potentiation of Bacillus thuringiensis insecticidal activity by serine pro-tease inhibitors. J. Agric. Food Chem. 38: 1145-1152.

MACINTOSH, S. C., T. B. STONE, R. S. JOKERST, AND R. L. FUCHS. 1991. Binding of Ba-cillus thuringiensis proteins to a laboratory-selected line of Heliothis virescens.Proc. Natl. Acad. Sci. USA 88: 8930-8933.

MALLET, J., AND P. PORTER. 1992. Preventing insect adaptation to insect-resistantcrops: are seed mixtures or refugia the best strategy? Proc. R. Soc. London B250: 165-169.

MARRONE, P. G., AND S. C. MACINTOSH. 1993. Resistance to Bacillus thuringiensisand resistance management, pp. 221-235 in P. F. Entwistle, J. S. Cory, M. J.Bailey, and S. Higgs [eds], Bacillus thuringiensis, an environmental biopesti-cide: theory and practice. New York:Wiley.

MARTIN, P. A. W., AND R. S. TRAVERS. 1989. Worldwide abundance and distribution ofBacillus thuringiensis isolates. Appl. Environ. Microbiol. 55: 2437-2442.

MASSON, L., A. MAZZA, R. BROUSSEAU, AND B. TABASHNIK. 1995. Kinetics of Bacillusthuringiensis toxin binding with brush border membrane vesicles from suscep-tible and resistant larvae of Plutella xylostella. J. Biol. Chem. 270: 1-10.

MAY, R. M. 1993. Resisting resistance. Nature 361: 593-594.


MCGAUGHEY, W. H. 1985. Insect resistance to the biological insecticide Bacillus thu-ringiensis. Science 229:193-195.

MCGAUGHEY, W. H. 1994. Problems of insect resistance to Bacillus thuringiensis. Agr.Ecosystem. Environ. 49: 95-102.

MCGAUGHEY, W. H., AND R. W. BEEMAN. 1988. Resistance to Bacillus thuringiensis incolonies of Indianmeal moth and almond moth (Lepidoptera:Pyralidae). J.Econ. Entomol. 81: 28-33.

MCGAUGHEY, W. H., AND D. E. JOHNSON. 1987. Toxicity of different serotypes and tox-ins of Bacillus thuringiensis to resistant and susceptible Indianmeal moths. J.Econ. Entomol. 80: 1122-1126.

MCGAUGHEY, W. H., AND D. E. JOHNSON. 1992. Indianmeal moth (Lepidoptera: Pyral-idae) resistance to different strains and mixtures of Bacillus thuringiensis. J.Econ. Entomol. 85: 1594-1600.

MCGAUGHEY, W. H., AND D. E. JOHNSON. 1994. Influence of crystal protein composi-tion of Bacillus thuringiensis strains on cross-resistance in Indianmeal moths(Lepidoptera: Pyralidae). J. Econ. Entomol. 87: 535-540.

MCGAUGHEY, W. H., AND M. E. WHALON. 1992. Managing insect resistance to Bacil-lus thuringiensis toxins. Science 258: 1451-1455.

MEADE, T., AND J. D. HARE. 1994. Effects of genetic and environmental host plantvariation on the susceptibility of two noctuids to Bacillus thuringiensis. Ento-mol. exp. appl. 70: 165-178.

MEADOWS, M. P. 1993. Bacillus thuringiensis in the environment: ecology and risk as-sessment, pp. 193-220 in P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs[eds], Bacillus thuringiensis, an environmental biopesticide: theory and prac-tice. New York:Wiley.

MIYASONO, M, M. YAMAMOTO, S. INAGAKI, K. OHBA, T. ISHIGURO, Y. HAYASHI, AND R.TAKEDA. 1994. Effects of several microorganisms on the insecticidal activity ofδ-endotoxin of Bacillus thuringiensis serovar kurstaki HD-1. Jpn. J. Appl. En-tomol. Zool. 38: 101-108.

MOAR, W., M. PUSZTAI-CAREU, H. VAN FAASSEN, R. MCCOLLUM, AND S. CLEMENT.1994. Resistance to Bacillus thuringiensis CryIC in Spodoptera exigua. p. 65 inAbstracts VIth Intern. Coll. Invertebr. Pathol.

MOLDENKE, A. F., R. E. BERRY, J. C. MILLER, J. G. WERNZ, X. H. LI. 1994. Toxicity ofBacillus thuringiensis subsp. kurstaki to gypsy moth, Lymantria dispar, fedwith alder or Douglas fir. J. Invertebr. Pathol. 64: 143-145.

MÜLLER-COHN, J., J. CHAUFAUX, C. BUISSON, N. GILOIS, V. SANCHIS, AND D. LERE-CLUS. 1994. Spodoptera littoralis resistance to the Bacillus thuringiensis cryICtoxin and cross-resistance to other toxins. p. 66 in Abstracts VIth Intern. Coll.Invertebr. Pathol.

NATIONAL RESEARCH COUNCIL. 1986. Pesticide Resistance: strategies and tactics formanagement. National Academy, Washington, DC. 471 pp.

NAVON, A. 1993. Control of lepidopteran pests with Bacillus thuringiensis, pp. 125-146 in P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs [eds], Bacillus thu-ringiensis, an environmental biopesticide: theory and practice. NewYork:Wiley.

OGIWARA, K., L. S. INDRASITH, S. ASANO, AND H. HORI. 1992. Processing of delta-en-dotoxin from Bacillus thuringiensis subsp. kurstaki HD-1 and HD-73 by gutjuices of various insect larvae. J. Invertebr. Pathol. 60: 121-126.

OPPERT, B., K. J. KRAMER, D. E. JOHNSON, S. C. MACINTOSH, AND W. H. MCGAUGHEY.1994. Altered protoxin activation by midgut enzymes from a Bacillus thuring-iensis resistant strain of Plodia interpunctella. Biochem. Biophys. Res. Comm.198: 940-947.

PARENTI, P, V. MANUELA, G. M. HANOZET, M. TASCA, AND B. GIORDANA. 1995. Inter-action of the insecticidal crystal protein CryIA from Bacillus thurigiensis withamino acid transport into brush border membranes from Bombyx mori larvalmidgut. J. Invertebr. Pathol. 65: 35-42.


PEFEROEN, M. 1992. Engineering of insect-resistant plants with Bacillus thuringien-sis crystal protein genes, pp. 135-153 in A. M. R. Gatehouse, V. A. Hilder, andD. Boulter [eds], Plant genetic manipulation for crop protection. Wallingford,United Kingdom:CAB International.

RAHARDJA, U., AND M. E. WHALON. 1995. Inheritance of resistance to Bacillus thur-ingiensis subsp. tenebrionis δ-endotoxin in the Colorado potato beetle (Co-leoptera: Chrysomelidae) J. Econ. Entomol. 88: 21-26.

RAMACHANDRIN R., K. F. RAFFA, M. J. MILLER, D. D. ELLIS, AND B. H. MCCOWN. 1993.Behavioral responses and sublethal effects of spruce budworm (Lepidoptera:Tortricidae) and fall webworm (Lepidoptera: Arctiidae) to Bacillus thuringien-sis CryIA(s) toxin in diet. Environ. Entomol. 22: 197-211.

ROBISON, D. J., B. H. MCCOWN, AND K. F. RAFFA. 1994. Responses of gypsy moth (Lep-idoptera: Lymanitriidae) and forest tent caterpillar (Lepidoptera: Lasiocampi-dae) to transgenic poplar, Populus spp., containing a Bacillus thuringiensis δ-endotoxin gene. Environ. Entomol. 23: 1030-1041.

ROUSCH, R. T. 1989. Designing resistance management programs: how can youchoose? Pestic. Sci. 26: 423-441.

SALAMA, H. S., AND M. M. MATTER. 1991. Tolerance level to Bacillus thuringiensisBerliner in the cotton leafworm Spodoptera littoralis Biosduval (Lepidoptera:Noctuidae). J. Appl. Entomol. 111: 225-230.

SALEH, M. S. 1987. Effect of larval selection with two bioinsecticides on susceptibilitylevels and reproductive capacity of Aedes aegypti. Anz. Schadingskd. Pflanz.Umweltschutz 60: 55-57.

SANCHIS, V., J. CHAUFAUX, AND D. PAURON. 1994. A comparison and analysis of thetoxicity and receptor binding properties of Bacillus thuringiensis CryIC δ-en-dotoxin on Spodoptera littoralis and Bombyx mori. FEBS Letters 353: 259-263.

SANGADALA, S., F. W. WALTERS, L. H. ENGLISH, AND M. J. ADANG. 1994. A mixture ofManduca sexta aminopeptidase and phosphatase enhances Bacillus thuring-iensis insecticidal CryIA(c) toxin binding and 86Rb+-K+ efflux in vitro. J. Biol.Chem. 269: 10088-10092.

SCHWARTZ, J. M., B. E. TABASHNIK, AND M. W. JOHNSON. 1991. Behavioral and phys-iological responses of susceptible and resistant diamondback moth larvae toBacillus thuringiensis. Entomol. Exp. Appl. 61: 179-187.

SHELTON, A. M., J. L. ROBERTSON, J. D. TANG, C. PEREZ, AND S. E. EIGENBRODE. 1993.Resistance of diamondback moth to Bacillus thuringiensis subspecies in thefield. J. Econ. Entomol. 86: 697-705.

SIMS, S. R., AND T. B. STONE. 1991. Genetic basis of tobacco budworm resistance to anengineered Pseudomonas fluorescens expressing the δ-endotoxin of Bacillusthuringiensis var. kurstaki. J. Invertebr. Pathol. 57: 206-210.

SMITH, R. A., AND G. A. COUCHE. 1991. The phylloplane as a source of Bacillus thur-ingiensis variants. Appl. Environ. Microbiol. 57: 311-315.

STONE, T. B., AND S. R. SIMS. 1993. Geographic susceptibility of Heliothis virescensand Helicoverpa zea (Lepidoptera: Noctuidae) to Bacillus thuringiensis. J.Econ. Entomol. 86: 989-994.

STONE, T. B., S. R. SIMS, S. C. MACINTOSH, R. L. FUCHS, AND P. G. MARRONE. 1991.Insect resistance to Bacillus thuringiensis, pp. 53-66 in K. Maramorosch [ed],Biotechnology for biological control of pests and vectors. Boca Raton, Flor-ida:CRC Press.

STONE, T. B., S. R. SIMS, AND P. G. MARRONE. 1989. Selection of tobacco budworm forresistance to a genetically engineered Pseudomonas fluorescens containing thedelta-endotoxin of Bacillus thuringiensis subsp. kurstaki. J. Invertebr. Pathol.53: 228-234.

TABASHNIK, B. E. 1986. Evolution of pesticide resistance in predator-prey systems.Bull. Entomol. Soc. America 32: 156-161.

TABASHNIK, B. E. 1989. Managing resistance with multiple pesticide tactics: theory,evidence and recommendations. J. Econ. Entomol. 82: 1263-1269.


TABASHNIK, B. E. 1992. Resistance risk management: realized heritability of resis-tance to Bacillus thuringiensis in diamondback moth (Lepidopteran: Plutel-lidae), tobacco budworm (Lepidopteran: Noctuidae), and Colorado potato beetle(Coloeptera: Chrysomelidae). J. Econ. Entomol. 85: 1551-1559.

TABASHNIK, B. E. 1994a. Evolution of resistance to Bacillus thuringiensis. Annu. Rev.Entomol. 39: 47-79.

TABASHNIK, B. E. 1994b. Delaying insect adaptation to transgenic plants: seed mix-tures and refugia reconsidered. Proc. R. Soc. London B 255: 7-12.

TABASHNIK, B. E., AND W. H. MCGAUGHEY. 1994. Resistance risk assessment for sin-gle and multiple insecticides: responses of Indianmeal moth (Lepidoptera:Pyralidae) to Bacillus thuringiensis. J. Econ. Entomol. 87:834-841.

TABASHNIK, B. E., N. L. CUSHING, N. FINSON, AND M. W. JOHNSON. 1990. Field devel-opment of resistance to Bacillus thuringiensis in diamondback moth (Lepi-doptera: Plutellidae). J. Econ. Entomol. 83: 1671-1676.

TABASHNIK, B. E., N. FINSON, C. F. CHILCUTT, N. L. CUSHING, AND M. W. JOHNSON.1993. Increasing efficiency of bioassays: evaluation of resistance to Bacillusthuringiensis in the diamondback moth (Lepidoptera: Plutellidae). J. Econ. En-tomol. 86: 635-644.

TABASHNIK, B. E., N. FINSON, F. R. GROETERS, W. J. MOAR, M. W. JOHNSON, K. LUO,AND M. J. ADANG. 1994a. Reversal of resistance of Bacillus thuringiensis inPlutella xylostella. Proc. Natl. Acad. Sci. USA 91: 4120-4124.

TABASHNIK, B. E., N. FINSON, AND M. W. JOHNSON. 1991. Managing resistance to Ba-cillus thuringiensis: lessons from the diamondback moth (Lepidoptera: Plutel-lidae). J. Econ. Entomol. 84: 49-55.

TABASHNIK, B. E., N. FINSON, M. W. JOHNSON, AND D. G. HECKEL. 1994b. Cross-re-sistance to Bacillus thuringiensis toxin CryIF in the diamondback moth (Plu-tella xylostella). Appl. Environ. Microbiol. 60: 4627-4629.

TABASHNIK, B. E., J. M. SCHWARTZ, N. FINSON, AND M. W. JOHNSON. 1992. Inherit-ance of resistance to Bacillus thuringiensis in the diamondback moth (Lepi-doptera: Plutellidae). J. Econ. Entomol. 85: 1046-1055.

TOJO, A., AND K. AIZAWA. 1983. Dissolution and degradation of Bacillus thuringiensisdelta-endotoxin by gut juice protease of the silkworm, Bombyx mori. Environ.Microbiol. 45: 576-580.

VAN FRANKENHUYZEN. 1993. The challenge of Bacillus thuringiensis, pp. 1-35 in P. F.Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs [eds], Bacillus thuringiensis, anenvironmental biopesticide: theory and practice. New York:Wiley.

VAN FRANKENHUYZEN, K., C. W. NYSTROM, AND B. E. TABASHNIK. 1995. Variation intolerance to Bacillus thuringiensis among and within populations of the sprucebudworm (Lepidoptera: Tortricidae) in Ontario. J. Econ. Entomol. 88: 97-105.

VAN RIE, J. S. 1991. Insect control with transgenic plants: resistance proof ? TrendsBiotechnol. 9: 177-179.

VAN RIE, J., S. JANSENS, H. HÖFTE, D. DEGHEELE, AND H. VAN MELLAERT. 1989. Spec-ificity of Bacillus thuringiensis delta-endotoxins. Eur. J. Biochem. 186: 239-247.

VAN RIE, J., S. JANSENS, H. HÖFTE, D. DEGHEELE, AND H. VAN MELLAERT. 1990a. Re-ceptors on the brush border membrane of the insect midgut as determinants ofthe specificity of Bacillus thuringiensis delta-endotoxins. Appl. Environ. Micro-biol. 56: 1378-1385.

VAN RIE, J., W. H. MCGAUGHEY, D. E. JOHNSON, B. D. BARNETT, AND H. VAN MEL-LAERT. 1990b. Mechanisms of insect resistance to the microbial insecticide Ba-cillus thuringiensis. Science 247: 72-74.

VISSER, B., D. BOSCH, AND G. HONÉE. 1993. Domain-function studies of Bacillus thu-ringiensis crystal protein, pp. 71-88 in P. F. Entwistle, J. S. Cory, M. J. Bailey,and S. Higgs [eds], Bacillus thuringiensis, an environmental biopesticide: the-ory and practice. New York:Wiley.


WALTERS, F. S., C. A. KULESZE, A. T. PHILLIPS, AND L. H. ENGLISH. 1994. A stable oli-gomer of Bacillus thuringiensis delta-endotoxin, CryIIIA. Insect Biochem.Molec. Biol. 24: 963-968.

WALTERS, F. S., S. L. SLATIN, C. A. KULESZE, AND L. H. ENGLISH. 1993. Ion channelactivity of n-terminal fragments from cryIA(c) delta-endotoxin. Biochem. Bio-phys. Res. Comm. 196: 921-926.

WATKINSON, I. 1994. Global view of present and future markets for Bt products. Agri.Ecosys. Environ. 49: 3-7.

WHALON, M. E., AND W. H. MCGAUGHEY. 1993. Insect resistance to Bacillus thuring-iensis, pp. 215-232 in L. Kim [ed], Advanced Engineered Pesticides, NewYork:Marcel Dekker.

WHALON, M. E., D. L. MILLER, R. M. HOLLINGWORTH. E. J. GRAFIUS, AND J. R.MILLER. 1993. Selection of a Colorado potato beetle (Coleoptera: Chrysomel-idae) strain resistant to Bacillus thuringiensis. J. Econ. Entomol. 86: 226-233.

WILSON, G. R., AND T. G. BENOIT. 1990. Activation and germination of Bacillus thur-ingiensis spores in Manduca sexta larval gut fluid. J. Invertebr. Pathol. 56: 233-236.

WOLFERSBERGER, M. G. 1990. The toxicity of two Bacillus thuringiensis δ-endotoxinsto gypsy moth larvae is inversely related to the affinity of binding sites on themidgut brush border membranes for the toxins. Experientia 46: 475-477.

WOLFERSBERGER, M. G. 1991. Inhibition of potassium-gradient-driven phenylalanineuptake in larval Lymantria dispar midgut by two Bacillus thuringiensis δ-en-dotoxins correlates with the activity of the toxins as gypsy moth larvicides. J.Exp. Biol. 161: 519-525.

WU, D., J. J. JOHNSON, AND B. A. FEDERICI. 1994. Synergism of mosquitocidal toxicitybetween CytA and CryIVD proteins using inclusions produced from clonedgenes of Bacillus thuringiensis. Mol. Microbiol. 13: 965-972.

YAMAMOTO, T., AND G. K. POWELL. 1993. Bacillus thuringiensis crystal proteins: re-cent advances in understanding insecticidal activity, pp. 215-232 in L. Kim

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

[ed], Advanced Engineered Pesticides, New York:Marcel Dekker.

Hoy: Symposium on Managing Resistance

443

MULTITACTIC RESISTANCE MANAGEMENT: AN APPROACH THAT IS LONG OVERDUE?

M

ARJORIE

A. H

OY

Department of Entomology and Nematology University of Florida, Gainesville, FL 32611-0620

A

BSTRACT

“Resistance management” tactics have been much discussed, but such tactics haveprovided surprisingly limited practical results for pest management programs todate. We have learned a great deal about pesticide resistance mechanisms, the modeof inheritance of resistances, the molecular basis of resistance and cross resistancemechanisms, and how to evaluate the impact of resistance on fitness. However, it re-mains difficult to “manage” resistance once resistant individuals make up more than5 to 10% of the population. Generally, the best that can be achieved is to

delay

the de-velopment of high levels of resistance for a few years, most often by using the productless often.

A more effective resistance management strategy will combine a variety of effec-tive pest management tactics along with a reduction in numbers and rates of pesti-cides applied. Effective pest management tactics include monitoring, evaluating

444



economic injury levels so that pesticides are applied only when needed, biological con-trol, host plant resistance, cultural controls, biorational pest controls, and geneticcontrol methods. As a part of this multi-tactic strategy, it is crucial to evaluate the im-pact of pesticides on natural enemies. Sometimes, pesticide-resistant natural enemiescan be effective components of a program to delay the development of resistance inpest arthropods.

Key Words: Pesticide resistance, integrated pest management, biological control, pes-ticide selectivity, resistance management.

R

ESUMEN

Las tácticas de manejo de la resistencia han sido muy discutidas pero sorprenden-temente, hasta la fecha, tales tácticas han dado resultados prácticos limitados en pro-gramas de manejo de plagas. Hemos aprendido mucho sobre los mecanismos deresistencia, el modo de heredarse, sus bases moleculares y los mecanismos de resis-tencia cruzada, así como a evaluar el impacto de la resistencia en el ajuste genético.Sin embargo, es aún difícil manejar la resistencia cuando los individuos resistentesintegran más del 5-10% de la población. Generalmente, lo mejor que se ha logrado hasido retrasar en varios años el desarrollo de altos niveles de resistencia en la pobla-ción, más a menudo usando menos f recuentemente el producto que la provoca. Unaestrategia más efectiva de manejo de la resisitencia combinaría una variedad de tác-ticas eficaces de manejo de plagas con la de reducir el número y dosis de los pesticidasaplicados. Las tácticas efectivas de manejo inculyen el monitoreo, la evaluación deldaño económico de modo que los pesticidas sean aplicados solamente cuando es nece-sario, el control biológico, la resistencia de las plantas hospedantes, el control cultu-ral, el control biorracional de plagas, y los métodos de control genético. Como parte deesta estrategia multitáctica, es crucial evaluar el impacto de los pesticidas en los ene-migos naturales. A veces, los enemigos naturales resistentes pueden ser conponentesefectivos de un programa para retrasar el desarrollo de la resistencia en los artrópo-

dos plagas.

This commentary will make the following argument: “integrated pest manage-ment” (IPM) and “management of pesticide resistance in pest arthropods” (MPR),which are usually perceived to be distinct topics for research, should be considered tohave equivalent goals and methods. When we accept that the goals and tactics aresimilar, we will develop effective resistance management programs for arthropods.Effective management of resistance and effective IPM programs require an holisticand multitactic strategy. A key component of this holistic and multitactic approach in-cludes enhancing the

compatibility

of pesticides and biological control agents (Hoy1992).

Resistance to pesticides is an extremely significant problem internationally, na-tionally (Georghiou 1986, Roush & Tabashnik 1990), and within Florida (Leibee &Capinera, this volume). At least 440 arthropod species have become resistant to insec-ticides and acaricides, with many species having become resistant to all the majorclasses of such products (Georghiou & Saito 1983, Georghiou 1986, Roush & Tabash-nik 1990). Resistance to pesticides in weeds, plant pathogens, and nematodes also isincreasing, although somewhat more slowly (National Academy of Sciences 1986,Denholm et al. 1992). While my commentary focuses on resistance to insecticides andacaricides, it will probably be applicable to fungicides, herbicides, and nematicides.

Developing and registering a new pesticide is an elaborate, and increasingly ex-pensive, business in the USA (Georghiou 1986) with costs estimated to be more than


445

$60 million per successful compound. Thus, pesticide producers should be increas-ingly interested in extending the economic life of their products in order to maximizea return on their investment.

Likewise, most pest management specialists want to preserve registered pesti-cides. This is especially true for products that are effective against arthropod pests inminor crops, which are increasingly being ignored by pesticide companies becausethey are such a small market. These so-called ‘minor’ crops of fruits and vegetablesare a major component of Florida’s agriculture. Registration of new pesticides is likelyto be more difficult and expensive in the future, which could leave some pest manage-ment specialists with extremely limited options for managing certain recalcitrantpests.

A few environmentalists have argued that we do not need pesticides, that they willsoon be outlawed, and that pesticide resistance will no longer be an important issue.However, it is unrealistic to eliminate all pesticides from agriculture; there are signif-icant arthropod pests for which we have no other effective control tactic. Pesticidesare the most effective tools for fighting outbreaks and emergency arthropod pest prob-lems, and they are often required to control plant pathogens, weeds, or nematodesthat cannot be controlled by alternative methods.

R

ESEARCH

A

PPROACHES

TO

R

ESISTANCE

M

ANAGEMENT

Scientists have approached the problem of pesticide resistance in a variety ofways. Fundamental research over the past 40 years has produced insights into resis-tance mechanisms (Corbett et al. 1984, Scott 1990, Soderlund & Bloomquist 1990)and the mode of inheritance of resistance in arthropods (Georghiou & Saito 1983,Scott 1990, Soderlund & Bloomquist 1990). Simulation models have been developedto evaluate different options for managing resistance (for a recent review, see Tabash-nik 1990), but the debate over whether to recommend (1) alternation of different pes-ticides or (2) mixtures of different pesticides for slowing the development of resistanceremains controversial and field-tested experimental data available are not strongenough to support either model (Roush & Daly 1990, Tabashnik et al. 1992). The hy-pothesis that reduced fitness, which is often associated with resistance alleles, couldbe used in management programs continues to be controversial and may have limitedapplication (Tabashnik 1990). Not all resistance alleles confer lowered fitness (for ex-ample, Hoy & Conley 1989, Hoy 1990) and natural selection can select for modifyinggenes that restore fitness to individuals carrying resistance alleles. Various monitor-ing techniques have been developed to identify resistant individuals and detect theirestablishment and spread (ffrench-Constant & Roush 1990). These methods are par-ticularly useful for documenting that resistance has occurred. However, monitoringmethods that would allow us to detect rare resistant individuals in populations in suf-ficient time that operational programs could be altered remain difficult and expensiveto execute (Brent 1986).

Resistance management research programs and IPM research programs have hadfairly distinct identities to date (Denholm et al. 1992, Croft 1990b, Hoy 1992). Becausethey have been distinct, an effective paradigm for resistance management has notbeen adopted in US agriculture. The current scenario usually goes something likethis: A pesticide is registered and used, resistant individuals are detected in popula-tions, people begin to discuss developing and implementing a resistance managementprogram. With this short-sighted approach, it is exceedingly difficult to develop andexecute a program in sufficient time to have the desired results.

Developing a resistance management program can take several years; studies typ-ically are conducted to develop an appropriate monitoring method, estimate the fre-

446



quency of resistant individuals in populations, detect cross resistances, and evaluatemode of inheritance and stability of the resistance (Roush & Daly 1990). Meanwhile,unless pesticide applications are discontinued, selection for resistance continues. Ini-tial detection of resistance usually requires that resistant individuals comprise atleast 5% of the population (Brent 1986). Thus, by the time resistant individuals aredetected, selection by additional pesticide applications is likely to increase their fre-quency in the population. This scenario is particularly familiar with multivoltine andhighly fecund pests such as aphids, spider mites, whiteflies, and leafminers.

Although we can learn from the experiences of scientists studying pesticide resis-tance in ubiquitous pests in other geographic regions and thus be alerted to a poten-tial problem, this seems to be an inefficient method for managing resistance inarthropods. Furthermore, this approach may be misleading because geneticists rec-ognize that different species or different geographic populations may develop resis-tance to a particular toxic chemical by a variety of mechanisms. The mechanisms,their mode of inheritance, and the degree of reduction in fitness associated with themmay vary, because the resistance alleles at each site are different. Because it is diffi-cult to sample for rare individuals in natural populations, monitoring programs maynot be cost effective if employed other than as a method to document a problem onceit has developed. Waiting until the pest becomes resistant before instituting a resis-tance management program is ineffectual (Hoy 1992).

R

ESISTANCE

AND

IPM

A better paradigm for managing resistance in arthropods involves altering pesti-cide use patterns, and nearly everyone will agree that reducing pesticide use is an ef-fective resistance management tactic (Croft 1990a, Tabashnik 1990, Leeper et al.1986). What has not been widely acknowledged is that resistance management pro-grams should include: 1) Altering the way pesticides are developed and registered,and 2) Recognizing that resistance management must be a broad-based, multitacticendeavor (Hoy 1992, Fig. 1).

It seems reasonable, conservative, and fiscally-responsible to assume that nearlyall major insect and mite pests will eventually become resistant to all classes of pes-ticides given sufficient selection pressure over sufficient time. There may be some ex-ceptions, but this generalization is reasonable given the documented record ofresistance development in arthropod pests during the past 40 years. Resistance tostress is a fundamental and natural response by living organisms (Scott, this volume).On an evolutionary time scale, it is apparent that insects will develop multiple and di-verse mechanisms to survive extreme temperatures, allelochemicals, and other envi-ronmental stresses. Thus, we should expect most insects to develop resistance to mostpesticides, if subjected to appropriate and sustained selection. While new pesticideclasses have been proclaimed to be potential ‘silver bullets’, and not amenable to re-sistance development, these hopes have been misplaced to date. It seems appropriateto assume that the development of resistance is nearly inevitable and the issue is not

whether

resistance will develop, but

when

. With this assumption, resistance manage-ment programs have the goal of

delaying

rather than preventing resistance (Hoy1992).

Growers and pest management experts can not afford to rely on pesticides as theirprimary management tool, as has been done for the past 30 years. There are increas-ing social, economic, and ecological pressures to reduce pesticide use and to increasethe use of nonchemical control tactics such as host plant resistance, biorational meth-ods, cultural controls, and biological controls (National Research Council 1989, Officeof Technology Assessment 1992). There is an increasing priority by research scien-


447

tists, regulatory agencies, legislators, and the public on using pesticides that are non-toxic to biological control agents and have minimal impact on the environment.Compatibility of pesticides with natural enemies and other nonchemical tactics iscritical for improving pest management and environmental quality, and for managingresistance to pesticides. Enhancing the compatibility of pesticides and biological con-trol agents is complex and sometimes difficult (Croft 1990a, Hoy 1985a, 1990), but canreward us with handsome dividends in improved pest control (Metcalf 1994) and pes-ticide resistance management (Tabashnik & Croft 1985).

The way pesticides are registered should be changed as part of an effective resis-tance management strategy (Hoy 1992). These changes also are essential in achievingimproved integrated pest management. For example, some pesticides are relativelynontoxic to important natural enemies in cropping systems at

low rates

, but the rec-ommended application rates are too high (Hoy 1985b). These high rates disrupt effec-tive biological control, leading to additional pesticide applications, which exertunnecessary selection for resistance in the pest. Under these circumstances, it may beappropriate for the label to contain two different directions for use; one rate could berecommended for the traditional strategy of relying on pesticides to provide control(although this is becoming a less and less viable option). A lower rate could be recom-mended for use in an IPM program that employs effective natural enemies. This ap-proach to labeling could reduce the number of pesticide applications and reduce rates,resulting in reduced selection for resistance in both target and nontarget pests.

Another innovation in pesticide registration would require that the toxicity of thepesticide to a selected list of biological control agents be determined for each croppingsystem. This information should be provided, either on the label or in readily-availablecomputerized data bases, perhaps via the internet. Without such information, use ofthese pesticides could disrupt benefits derived from effective biological control agents.

Figure 1. Effective resistance management programs will incorporate multipletactics as part of a fully integrated IPM program. Pesticides should be used only whenneeded, at the lowest rates possible, and in a manner to reduce negative impacts onarthropod natural enemies. Pesticide labels should include information about its im-pact on natural enemies.

448



This often results in unnecessary use of pesticides, leading to more rapid evolution ofpesticide resistance. Enhancing biological control not only leads to improved pest man-agement, but is also an essential tool in managing pesticide resistance.

How could information about the toxicity of pesticides to biological control agentsbe made available to the end user most effectively? How should bioassays be con-ducted to evaluate pesticide selectivity? There are no simple answers to these ques-tions. Theiling & Croft (1988) and Croft (1990b) compiled an extensive set of data onthe impact of pesticides on natural enemies, but additional data are also buried inmany publications or reports that are difficult to find. Unfortunately, even if the datacan be found, it is not always easy to interpret bioassay data obtained by different sci-entists using different methods. Thus, different conclusions about the toxicity of pes-ticides to natural enemies can be drawn. Also, it is often difficult to predict the impactof pesticides under field conditions based on laboratory assays (Hoy 1990, Hassan etal. 1991, Robertson & Preisler 1992). Consequently, the recommendation that labelsor data bases be developed with information on the impact of pesticides on natural en-emies requires considerable discussion and additional research. Should pesticidecompanies conduct the research using standard bioassay methods? Should a consor-tium of pest management scientists conduct the assays? Who should pay for the re-search? What species of natural enemies should be tested? These questions are notnew, and in Europe standardized bioassays already are being conducted on selectednatural enemy species by a scientific working group (Hassan et al. 1991). Whetherthis concept can be imported to the USA and Florida should be explored. Increased in-ternational consultation and cooperation between scientists, regulatory agencies, andpesticide companies could resolve many of the questions raised above.

Evolution of pesticide resistance has been shown by computer simulations of pred-ator and prey systems to be slowed by reduced pesticide use (Tabashnik & Croft 1985,Tabashnik 1990). There is general agreement that reduced pesticide use is one of theessential elements of any resistance management program (Croft 1990b, Tabashnik1990, Metcalf 1994). Thus, the compatibility of pesticides and biological controlagents is a crucial issue in pesticide resistance management.

Attempts to manage pesticide resistance generally has involved making relativelyminor tactical shifts in use patterns. What we need is a major shift in thinking aboutpesticide development and use, if we are to develop effective resistance managementtactics. It is time to recognize that effective resistance management

begins

before theproduct is even registered (Figure 1). The strategy thus should be to manage the pes-ticide, even before it is fully developed and registered, with the goal of delaying resis-tance development. If this farsighted strategy is adopted, decisions on applicationrates and the numbers of applications per growing season will be made with the un-derstanding that they affect the speed with which resistance will develop. In somecases, new products may not be developed because they are toxic to biological controlagents and thus could disrupt effective IPM programs already in place. This wouldonly speed up the development of resistance in specific pests.

Some people suggest that financial incentives may have to be provided to inducepesticide companies to develop and register products that are harmless to biologicalcontrol agents (= physiological selectivity). Others argue that many pesticides can beapplied in a manner that affords substantial selectivity (= ecological selectivity) if thetiming, location, rates, and methods of application are altered (Hull & Beers 1985).Relying on ecological selectivity is more likely to be cost effective than developinglarge numbers of special use products with physiological selectivity. However, a com-plete financial analysis may indicate that, over the long term, selective pesticides(based both on physiological and ecological selectivity) are among the most cost effec-tive approaches to managing resistance to pesticides.


449

T

HE

R

OLE

OF

P

ESTICIDE-

R

ESISTANT

N

ATURAL

E

NEMIES

Pesticide-resistant natural enemies are a special category of pesticide selectivity.Relatively few natural enemies have developed resistance to pesticides through natu-ral selection, but several have been employed in effective IPM programs (Croft 1990a,Hoy 1990). Artificial selection of phytoseiid predators for pesticide resistance can be apractical and cost effective tactic for the biological control of spider mites (Hoy 1985a,b, 1990). Field tests have been conducted with several laboratory-selected phytoseiidspecies and some are being used in IPM programs. Laboratory selection of resistantstrains of parasitoids and insect predators currently lags behind efforts with predatorymites, but several laboratory-selected insect natural enemies are being evaluated forincorporation into integrated pest management programs (Hoy 1994). The use of mu-tagenesis and recombinant DNA techniques could improve the efficiency of genetic im-provement projects. However, development of pesticide-resistant natural enemies istime consuming and expensive and should not be considered before exploring other,less expensive, options for IPM and pesticide resistance management.

C

ONCLUSIONS

We need to maintain a source of pesticides because they are powerful and effectivepest management tools. Pesticides can be highly selective, rapid in their impact,adaptable to many situations, and relatively economical. Thus, preserving pesticidesis essential.

Effective paradigms for resistance management are not yet deployed in US andFlorida agriculture. This is because resistance management and IPM have been con-sidered separate issues. We need to recognize that effective resistance management isbased on the development of effective, fully-integrated, multitactic IPM programs.Such programs will acknowledge the role of biological control, host plant resistance,cultural controls, and biorational controls, such as mating disruption, insect growthregulators, and mass trapping. A key issue should always be whether pesticides canbe used in a precise and selective manner without disrupting the impact of natural en-emies.

While altering the way in which pesticides are registered and labeled is difficult,the potential benefits are great for both IPM and resistance management programs.The dialogue should begin on how best to change pesticide labeling and develop data-bases on the selectivity of pesticides to natural enemies.

A

CKNOWLEDGMENTS

This is University of Florida, Institute of Food and Agricultural Sciences, journalseries number R-04510. I thank D. C. Herzog for comments on the manuscript and theFlorida Entomological Society for the opportunity to organize the symposium, “Mythsof Managing Resistance”. A brief essay containing many of the ideas presented herehas been published (Hoy 1992). The references cited provide an entre to the extensiveliterature on resistance and are not complete.

R

EFERENCES

C

ITED

B

RENT

, K. J. 1986. Detection and monitoring of resistant forms: an overview, pp.298-312

in

Pesticide Resistance: Strategies and Tactics for Management. Na-tional Academy Press, Washington, D.C.

C

ORBETT

, J. R., K. W

RIGHT

AND

A. C. B

AILLIE

. 1984. The Biochemical Mode of Actionof Pesticides. 2nd ed., Academic Press, New York.

450



C

ROFT

, B. A. 1990a. Arthropod Biological Control Agents and Pesticides.Wiley-Interscience Publ., New York.

C

ROFT

, B. A. 1990b. Developing a philosophy and program of pesticide resistancemanagement, pp. 277-296

in

R. T. Roush and B. E. Tabashnik [eds.] PesticideResistance in Arthropods, Chapman and Hall, New York.

D

ENHOLM

, I., A. L. D

EVONSHIRE

AND

D. W. H

OLLOMON

[eds.]. 1992.Resistance 91:Achievements and Developments in Combating Pesticide Resistance, ElsevierApplied Science, London.

FFRENCH-

C

ONSTANT

, R. H.

AND

R. T. R

OUSH

. 1990. Resistance detection and docu-mentation: the relative roles of pesticidal and biochemical assays. pp. 4-38.

G

EORGHIOU

, G. P. 1986. The magnitude of the resistance problem, pp. 14-43

in

R. T.Roush and B. E. Tabashnik [eds.] Pesticide Resistance in Arthropods, Chap-man and Hall, New York.

G

EORGHIOU

, G. P.

AND

T. S

AITO

[eds.] 1983. Pest Resistance to Pesticides, PlenumPress, New York.

H

ASSAN

, S. A., F. B

IGLER

, H. B

OGENSCHUTZ

, E. B

OLLER

, J. B

RUN

, J. N. M. C

ALIS

, P.C

HIVERTON

, J. C

OREMANS-

P

ELSENEER

, C. D

USO

, G. B. L

EWIS

, F. M

ANSOUR

, L.M

ORETH

, P. A. O

OMEN

, W. P. J. O

VERMEER

, L. P

OLGAR

, W. R

IECKMANN

, L.S

AMSOE-

P

ETERSEN

, A. S

TAUBLI

, G. S

TERK

, K. T

AVARES

, J. J. T

USET

AND

G. V

IG-GIANI

. 1991. Results of the fifth joint pesticide testing programme carried outby the IOBC/WPRS-Working Group “Pesticides and Beneficial Organisms”.Entomophaga 36: 55-67.

H

OY

, M. A. 1985a. Almonds: integrated mite management for California almond or-chards, pp. 299-310

in

W. Helle and M. W. Sabelis [eds.]. Spider Mites, Their Bi-ology, Natural Enemies, and Control, Vol. 1B. Elsevier, Amsterdam.

H

OY

, M. A. 1985b. Recent advances in genetics and genetic improvement of the Phy-toseiidae. Annu. Rev. Entomol. 30: 345-370.

H

OY

, M. A. 1990. Pesticide resistance in arthropod natural enemies: variability andselection responses, pp. 203-236

in

R. T. Roush and B. E. Tabashnik [eds.] Pes-ticide Resistance in Arthropods, Chapman and Hall, New York.

H

OY

, M. A. 1992. Guest editorial: Proactive management of pesticide resistance in ag-ricultural pests. Phytoparasitica 20(2): 93-97.

H

OY

, M. A. 1994. Transgenic pest and beneficial arthropods for pest management pro-grams: an assessment of their practicality and risks, pp. 641-670

in

D. Rosen,F. D. Bennett and J. L. Capinera [eds.] Pest Management in the Subtropics: Bi-ological Control a Florida Perspective. Intercept Ltd., Andover, U.K.

H

OY

, M. A.

AND

J. C

ONLEY

. 1989. Propargite resistance in Pacific spider mite (Acari:Tetranychidae): stability and mode of inheritance. J. Econ. Entomol. 82: 11-16.

H

ULL

, L. A.

AND

E. H. B

EERS

. 1985. Ecological selectivity: modifying chemical controlpractices to preserve natural enemies, pp. 103-122 in M. A. Hoy and D. C. Her-zog [eds.], Biological Control in Agricultural IPM Systems. Academic, Orlando.

LEEPER, J. R., R. T. ROUSH AND H. T. REYNOLDS. 1986. Preventing or managing resis-tance in arthropods, pp. 335-346 in Pesticide Resistance: Strategies and Tacticsfor Management, National Academy Press, Washington, D.C.

METCALF, R. L. 1994. Insecticides in pest management, pp. 245-284 in R. L. Metcalfand William H. Luckmann [eds.], Introduction to Insect Pest Management,third ed., John Wiley and Sons, New York.

NATIONAL ACADEMY OF SCIENCES. 1986. Pesticide Resistance: Strategies and Tacticsfor Management. National Academy Press, Washington, D.C.

NATIONAL RESEARCH COUNCIL. 1989. Alternative Agriculture, National AcademyPress, Washington, D. C.

OFFICE OF TECHNOLOGY ASSESSMENT. 1992. A New Technological Era for AmericanAgriculture, U. S. Congress, U.S. Government Printing Office, Washington,D.C.

ROBERTSON, J. L. AND H. K. PREISLER. 1992. Pesticide Bioassays with Arthropods.CRC Press, Boca Raton, Florida.

Hoy: Symposium on Managing Resistance 451

ROUSH, R. T. AND J. C. DALY. 1990. The role of population genetics in resistance re-search and management, pp. 97-152 in R. T. Roush and B. E. Tabashnik [eds.]Pesticide Resistance in Arthropods, Chapman and Hall, New York.

ROUSH, R. T. AND B. E. TABASHNIK [eds.] 1990. Pesticide Resistance in Arthropods.Chapman and Hall, New York, NY.

SCOTT, J. G. 1990. Investigating mechanisms of insecticide resistance: methods, strat-egies, and pitfalls, pp. 39-57 in R. T. Roush and B. E. Tabashnik [eds.] PesticideResistance in Arthropods, Chapman and Hall, New York.

SODERLUND, D. M. AND J. R. BLOOMQUIST. 1990. Molecular mechanisms of insecticideresistance, pp. 58-96 in R. T. Roush and B. E. Tabashnik [eds.] Pesticide Resis-tance in Arthropods, Chapman and Hall, New York.

TABASHNIK, B. E. 1990. Modeling and evaluation of resistance management tactics,pp. 153-182 in R. T. Roush and B. E. Tabashnik [eds.] Pesticide Resistance inArthropods, Chapman and Hall, New York.

TABASHNIK, B. E. AND B. A. CROFT. 1985. Evolution of pesticide resistance in applepests and their natural enemies. Entomophaga 30: 37-49.

TABASHNIK, B. E., J. A. ROSENHEIM AND M. A. CAPRIO. 1992. What do we really knowabout management of insecticide resistance? pp. 124-135 in I. Denholm, A. L.Devonshire and D. W. Hollomon [eds.]. 1992. Resistance 91: Achievements andDevelopments in Combating Pesticide Resistance, Elsevier Applied Science,London.

THEILING, K. M. AND B. A. CROFT. 1988. Pesticide side-effects on arthropod naturalenemies: a database summary. Agric. Ecosys. Environm. 21: 191-218.

452



IMMUNOLOGICAL STUDY OF JUVENILE HORMONE BINDING PROTEIN FROM HEMOLYMPH OF THE FALL

WEBWORM (LEPIDOPTERA: ARCTIIDAE)

I. H. L

EE

1

, H

AK

R. K

IM

1

AND

R

ICHARD

T. M

AYER

2

1

Department of Biology, Korea University, Seoul, Korea

2

USDA, ARS, U.S. Horticultural Research Laboratory,2120 Camden Road, Orlando, FL 32803-1419

A

BSTRACT

The juvenile hormone binding protein (JHBP) was purified from the hemolymphof

Hyphantria cunea

(Lepidoptera: Arctiidae) using anion exchange, gel filtration, andMono P FPLC chromatofocusing chromatography. The protein is a single polypeptide(M

r

32,000) with an apparent dissociation constant of 0.43

µ

M for juvenile hormoneIII. An antibody developed against hemolymph JHBP (hJHBP) was prepared, and useof it showed that a protein immunologically identical with hJHBP occurs in fat bodyand ovary. The hJHBP of

H. cunea

was neither immunologically related to the lipo-phorin from

H. cunea

nor similar to the hJHBP of

Bombyx mori

and

Periplaneta amer-icana

.

Key Words: Hemolymph, hemolymph proteins, insect, immunodetection, juvenile hor-mone III, lipoprotein, protein purification.

R

ESUMEN

La proteína de unión de la hormona juvenil (JHBP) fue purificada a partir de la he-molinfa de

Hyphantria cunea

. Para purificar la JHBP fueron usados el intercambio deaniones, la filtración en gel y cromatografías cromatofocales Mono P FPLC. La pro-teína es un polipéptido simple (M

r

32,000) con una constante de disociación aparentede 0.43

µ

M para la hormona juvenil III. Fue preparado un antibiótico desarrolladocontra la hemolinfa JHBP (hJHBP) y su uso demostró que una proteína inmunológi-camente idéntica a la hJHBP está presente en el cuerpo graso y el ovario. La hJHBPde

H. cunea

no estuvo inmunológicamente relacionada con la lipoforina de

H. cunea

ni fue similar a la hJHBP de

Bombyx mori

y

Periplaneta americana.

Juvenile hormone is synthesized by the corpora allata, released into hemolymph,and transported to target tissues by the juvenile hormone binding protein (JHBP)(Whitmore & Gilbert 1972; Rudnica et al. 1979; Ozyhar et al. 1983). JHBP is synthe-sized by fat body and released into the hemolymph (Nowock et al. 1976; Ferkovich etal. 1977). Generally, high M

r

(over 200,000) and low M

r

(20,000-40,000) JHBPs arepresent in the hemolymph (de Kort & Granger 1981; de Kort et al. 1983). JHBP fromhemolymph was first purified from

Manduca sexta

(Kramer et al. 1976) and then from

Diatraea grandiosella

(Dillwith et al. 1985),

Diploptera punctata

(King & Tobe 1988),and

Platyprepia virginalis

(Prestwich & Atkinson 1990). It has been reported thathemolymph juvenile hormone binding protein (hJHBP) is closely related toapoprotein-I of lipophorin (Rayne & Koeppe 1988; Koopmanschap & Dekort 1988).

Proteins with JH affinity were found to be present in ovary and fat body and theircharacteristics were also reported (van Mellaert et al. 1985; Koeppe et al. 1987; Shem-

Lee et al.: JHBP of

H. cunea 453

shedini & Wilson 1993). However, information on the relationship between hJHBPand JH binding protein in these tissues is still limited, thus additional information isneeded for a better understanding of the role of JHBP.

Here, we describe the presence of a JHBP in ovary and fat body which is immuno-logically identical with hJHBP by using an antibody against low M

r

hJHBP from thefall webworm,

Hyphantria cunea

Drury (Lepidoptera: Arctiidae). In addition, thephysicochemical characteristics of hJHBP are described.

M

ATERIALS

AND

M

ETHODS

Insects

Fall webworms,

Hyphantria cunea

, were reared on artificial diet (Dong Bang OilCo., Ltd., Seoul, Korea) at 28

±

1

°

C and 70

±

5% relative humidity and with a photo-period of 16:8 (L:D).

Collection of Hemolymph and Extraction of Ovary and Fat Body

Hemolymph was collected into a cold Eppendorf microcentrifuge tube from last in-star larvae by cutting the forelegs. A few crystals of phenylthiourea were added to pre-vent melanization, and the hemolymph was centrifuged at 10,000 g for 10 min at 4

°

Cto remove hemocytes and cell debris. The supernatant was stored at -70

°

C.Ovaries were dissected from 5-day-old pupae in cold Ringer’s solution (128 mM

NaCl, 1.8 mM CaCl

2

, 1.3 mM KCl; pH 7.4). They were used directly for electron mi-croscopic observation or homogenized and centrifuged at 10,000 g for 10 min and thesupernatant used as the sample.

Fat body was dissected from last instar larvae in Ringer’s solution. These tissueswere stored at -70

°

C until used.

Dextran Coated Charcoal (DCC) Binding Assay

A DCC suspension (0.5% active charcoal and 0.05% dextran, M

r

80,000) in buffer(10 mM Tris, 5 mM MgCl

2

, 50 mM KCl, pH 7.4) was used to separate protein-boundJH from unbound hormone (Engelmann 1981). [

3

H]JH III (NEN Corp., Wilmington,DE; specific activity: 440 GBq/mmol; 50,000 cpm) was put into a microcentrifuge tubeand the solvent evaporated with a gentle stream of nitrogen gas. Buffer (100

µ

l; 10mM Tris, 5 mM MgCl

2

, 50 mM KCl, pH 7.4), and the sample used for the binding as-say, were added to the hormone and incubated for 1 h at 4

°

C. DCC suspension (100

µ

l)was added to the tube, the contents incubated for 2 min and then centrifuged at10,000 g to remove free hormone. An aliquot (10

µ

l) of supernatant was transferred to10 ml of scintillation cocktail solution (toluene 2 liters, triton X-100 1 liter, 12 g Om-nifluor, NEN) and the radioactivity was measured by liquid scintillation counter(Beckman LS 100

°

C, Palo Alto, CA).

Purification of JHBP

Anion exchange chromatography.

Hemolymph (10 ml) was diluted 1:1 with TPNbuffer [10 mM Tris, 0.1 mM phenylmethylsulfonyl fluoride (PMSF), 0.01% NaN

3

, pH8.2] and dialyzed against the same buffer for 4 h. Dialyzed hemolymph was applied toa DEAE cellulose (DE 52, Whatman, Hillsboro, OR) column (1.8

×

20 cm) and elutedwith 1 column volume of TPN buffer. This was followed by a NaCl concentration gra-dient (0-0.5 M) in buffer (10 mM Tris, 0.01% NaN

3

, 0.1 mM PMSF, pH 8.2, total 300

454



ml) at a flow rate of 18 ml per h and then monitored for the presence of JHBP by theDCC binding assay. Fractions containing JHBP were pooled, dialyzed against 10 mMTris buffer (0.01% NaN 0.1 mM PMSF, pH 8.2) and applied to a Sepharose CL-6B(Pharmacia, Piscataway, NJ) column (0.9

×

15 cm). The column was washed with thesame buffer using a gradient of 0.05-0.35 M NaCl. Each fraction (2.6 ml) was moni-tored for JHBP by the DCC binding assay. Fractions containing JHBP were lyo-philized before the next purification step.

Gel Filtration.

The lyophilized hemolymph sample was dissolved in 2 ml water andapplied to a Sephadex G-100 column (1.8

×

55 cm) and eluted with buffer (25 mMbis-Tris, pH 7.2) at a flow rate of 12 ml per h. Each fraction (2.6 ml) was monitored forJHBP by the DCC binding assay. Fractions containing JHBP were pooled, concen-trated by lyophilization, and applied to a Sephadex G-75 column (0.9

×

20 cm) andeluted with buffer (25 mM bis-Tris, pH 7.2) at a flow rate of 8 ml per h. Fractions of1.5 ml were collected.

Mono P FPLC Chromatofocusing Chromatography.

Fractions containing JHBPfrom the Sephadex G-75 column were subjected to FPLC chromatofocusing chroma-tography on a Mono P column (Mono P HR 5120, 5

×

200 mm, Pharmacia). Free pro-teins were eluted from the column with 25 mM bis-Tris buffer (pH 7.2, 10 ml), andthen Polybuffer (pH 7.4, 52 ml) was used to elute resin-bound proteins. Each fractionwas monitored for JHBP by the DCC binding assay.

Determination of Molecular Weight and Isoelectric Point

The molecular weights were determined on sodium dodecyl sulfate-polyacrylamidegel (10%) as described by Weber & Osborn (1969). Standard molecular weight markerswere: bovine serum albumin, M

r

66,200; hen egg albumin, M

r

45,000; bovine carbonicanhydrase, M

r

31,000; soybean trypsin inhibitor, M

r

21,500; hen egg white lysozyme,M

r

14,400.Isoelectric focusing was performed on 5% polyacrylamide gel according to Klages

& Emmerich (1979) by using ampholytes in the pH range 3-10 (Sigma, St. Louis, MO).

Production of Antibody Against JHBP

Fractions (0.5 ml) containing JHBP after Mono P FPLC chromatofocusing weremixed thoroughly with Freund’s complete adjuvant (0.5 ml) and injected subcutane-ously into a rabbit. Injections were repeated every other day for the first week. Thefourth injection was conducted 1 week after the third injection, and a booster injectionwas given 2 weeks later. JHBP-containing fractions (0.5 ml) were mixed with incom-plete adjuvant (0.5 ml) for the booster injection. Blood was taken 1 week after thebooster injection and centrifuged at 10,000 g for 10 min. Antibody against JHBP waspurified by immunoprecipitation of extraneous antibodies using JHBP-free fractions.

Immunodiffusion

Immunodiffusion was conducted using 1% agarose gel containing 0.1% (w/v) so-dium azide in veronal buffer (pH 8.6) as described by Ouchterlony (1949). The plateswere stained in 1% amido black 10B and destained in 2% acetic acid.

Electrophoresis and Immunodetection

Fractions from each purification step were electrophoresed on 10%SDS-polyacrylamide gel as described by Laemmli (1970) to determine purity. Gelswere stained with silver nitrate according to Wray et al. (1981).

Lee et al.: JHBP of

H. cunea 455

Western blotting was used to detect JHBP on SDS-polyacrylamide gels. Sampleswere electrophoresed as above and transferred to nitrocellulose sheets in Tris-glycinebuffer (25 mM Tris, 92 mM glycine, 30% methanol, pH 8.3) at 100 V for 2 h (Towbinet al. 1979). After transfer, the nitrocellulose sheets were equilibrated in TBS (20 mMTris, 500 mM NaCl, pH 7.5) for 10 min and incubated in blocking solution (3% gelatinin TBS) for 30 min. The sheet was then washed with TTBS (0.05% tween 20 in TBS)twice, each time for 5 min, and incubated for 1 h in a solution containing a 300-folddilution of the primary antibody against JHBP. This sheet was again washed withTTBS twice and incubated for 1 h in a solution containing a 3000-fold diluted second-ary antibody (goat antirabbit-horseradish peroxidase conjugated IgG). After incuba-tion, the sheet was again washed with TTBS twice and submerged in horseradishperoxidase color development reagent containing 4-chloro-1-naphthol in 20 mlice-cold methanol + 0.015% H

2

O

2

in 100 ml TBS for development of purple color.

Electron Microscopic Observation by Immunogold Labeling

Ovaries and fat body were dissected from pupae in Ringer’s solution and prefixedin 2.5% glutaraldehyde for 2 h at 4C. Tissues were washed with 0.1 M phosphatebuffer (pH 7.2) 3 times at 15 min intervals and dehydrated in an ethanol series. De-hydrated tissues were then put into propylene oxide and embedded in anEpon-Araldite mixture. Embedded tissues were semithin-sectioned using an ultrami-crotome (Sorvall MT-II, Wilmington, DE) and stained in 1% toluidine blue and at-tached to a grid. The thin section-attached grid was washed with TBS. This grid wasreacted with the primary antibody solution 30-fold diluted with antibody buffer andagain washed with TBS 3 times to remove nonspecifically attached antibody. This gridwas then reacted with antirabbit IgG combined with protein-A gold particles (30 nmin diam) for 40 min, washed with TBS and distilled water 3 times each, stained in 2%uranyl acetate and observed under JEOL JEM 100 CX-II electron microscope at 80kV.

R

ESULTS

Purification of JHBP from Hemolymph

The DCC binding assay for each fraction showed that JHBP was eluted from theDE-52 column with a linear NaCl gradient (0.06-0.1 M) (Fig. 1A). Fractions contain-ing JHBP were subjected to a second anion exchange chromatography treatment(Sepharose CL-6B). JHBP was found to be present in the backside of the first peak(0.1 M NaCl) (Fig. 1B). Fractions containing JHBP from the second ion exchange chro-matography run were subjected to gel filtration (Sephadex G-100) and fractions con-taining radioactivity were located (Fig. 1C) and pooled. The pooled fractions wereapplied to Mono P FPLC chromatofocusing columns and 4 peaks isolated by using A

280

(Fig. 1D). The fourth peak exhibited high binding activity for [

3

H]JH III (Fig. 1D, in-set). Because the JHBP fractions were contaminated with other hemolymph proteinsafter Mono P chromatofocusing, gel filtration on Sephadex G-75 was necessary (datanot shown). Fractions were applied to a 10% SDS gel and completely purified JHBPwas confirmed (Fig. 2A).

Characterization of hJHBP

JHBP was electrophoresed with a low molecular weight standard marker to deter-mine the M

r

of the JHBP subunit. The M

r

was estimated to be 32,000 (Fig. 2B). Also,

456



the pI was determined to be 5.3 (data not shown). Saturation analysis of the purifiedhJHBP revealed a high affinity for JH III (K

D

of 0.43

µ

M) (Fig. 3).

Immunological Studies

Antibodies were made against fractions containing JHBPs (Fig. 4A, lane b) thathad been separated by Mono P FPLC chromatofocusing. Extraneous antibodies, i.e.,other than antibody against hJHBP, were precipitated with JHBP-free fractions (Fig.4A, lane a). The resulting supernatant from this precipitation showed pure hJHBPantiserum (Fig. 4B).

A protein that is immunologically identical with hJHBP was found in ovaries byimmunodiffusion and western blotting (Fig. 5). However, the hJHBP of

H. cunea

wasneither immunologically related to the lipophorin from

H. cunea

nor was it similar tothe hJHBP of

Bombyx mori

and

Periplaneta americana

(Fig. 5). The western blots

Figure 1. Chromatography elution profiles for purification of H. cunea hemolymphJHBP. A, the first anion exchange chromatography of dialyzed hemolymph proteinson DE52. Hatched region (35-46) indicates fractions containing JHBP; B, the secondanion exchange chromatography on Sepharose CL-6B. Fractions (47-55, hatched re-gion) showing radioactivity in DCC binding assay were pooled and lyophilized for thenext purification step; C, gel (Sephadex G-100) filtration of partially purified JHBP.JHBP containing fractions were confirmed (hatched region, 31-37); D, Mono P FPLCchromatofocusing column. Four major protein peaks were resolved and collected(fractions 26-29, 32, 35, 37-38). Each peak was also analyzed by DCC binding assay(inset). Arrow indicates peak containing JHBP.

Lee et al.: JHBP of

H. cunea 457

shown in Figure 5 (left) show a crossreaction for the

H. cunea

hemolymph, ovarian ex-tracts, and JHBP with the antibody against hJHBP. Likewise, the immunodiffusiontest (Fig. 5, right) showed an immunopreciptation line extending from the JHBP wellthrough the ovarian extract and ending at the

H. cunea

hemolymph well, thus corrob-orating the western blot data.

Electron microscopic observations showed that gold particles accumulated in ova-rian protein bodies as well as the fat body (Fig. 6). These results clearly indicate pro-tein that is immunologically identical with hJHBP is present in protein bodies ofovary.

D

ISCUSSION

The DCC binding assay was employed to purify a low M

r

JHBP. Fractions showingradioactivity according to the DCC binding assay were electrophoresed and silverstained and found to be a single polypeptide with an M

r

32,000. The protocols em-ployed to purify hemolymph JHBP from

H. cunea

are similar to those used to purifyJHBP of

Diatraea grandiosella

(Dillwith et al. 1985) and

Manduca sexta

(Kramer etal. 1976). The hJHBP of

H. cunea

has a pI of 5.3 (data not shown) and can be effec-tively separated from other hemolymph proteins with chromatofocusing. Low M

r

Figure 2. A, electrophoretic profile of fractions from final purification step (gel fil-tration on Sephadex G 75). Anterior portions of the chromatographic profile still con-tain other proteins in addition to JHBP (A: a, b, c, d). Completely purified JHBP fromthe posterior portion of the chromatography was confirmed on lane e. The gel (10%)was stained with silver nitrate. B, SDS-polyacrylamide gel (15%) electrophoresis todetermine the molecular weight of JHBP. The gel was stained with coomassie bril-liant blue. Right lane contains the low molecular weight standard markers and thearrow in left lane indicates JHBP.

458



JHBP has an M

r

range of 20,000-40,000 in several insects (de Kort & Granger 1981;de Kort et al. 1983).

The dissociation constant of

H. cunea hJHBP was estimated to be 0.43 µM. Thisvalue is similar to those of pyralid moth’s 0.08-0.28 µM (Lenz et al. 1986), D. grandi-osella 0.31 µM, (Dillwith et al. 1985), and M. sexta, 0.44 µM (Kramer et al. 1976). TheJHBP purified from hemolymph of H. cunea is of low Mr and high affinity.

Thus far, most JHBP antibody production employed large Mr proteins. Prepara-tion of antibodies for the low Mr JHBP was difficult because of the small amount of

Figure 3. The saturation analysis (Scatchard plot) of the binding of JH III to thepurified hJHBP. •••, total binding; °°°, nonspecific binding; ▲▲▲, specific binding.The specific binding was used for the Scatchard analysis (inset).

Lee et al.: JHBP of H. cunea 459

Figure 4. Western blot of purified JHBP. The JHBP band was detected by enzymeimmunoassay using antibody against JHBP (B,b). Left panel (A) shows the electro-phoretic patterns of fractions from the Mono P FPLC chromatofocusing chromatogra-phy. The JHBP-containing fractions (the fourth peak that the arrow indicates in Fig.1D, lane b) were used for the production of antibody and fractions (anterior threepeaks in Fig. 1D, lane a) that lack JHBP were used for the immunoprecipitation to re-move other antibodies except antibody against JHBP.


material in hemolymph. In the present work with H. cunea, immunoprecipitation wasvery effective in purifying antibody against low Mr JHBP from other antibodies.

hJHBP was confirmed to be lipophorin in Leucophaea maderae (Koeppe et al.1988) and Locusta migratoria (Koopmanschap & de Kort, 1988). Immunodiffusionstudies conducted here indicate that the hJHBP was neither immunologically relatedto lipophorin in H. cunea nor the hJHBP of Bombyx mori and Periplaneta americana.The hJHBP of H. cunea appeared to be species-specific.

hJHBP is synthesized by fat body and released into hemolymph. Protein with JHaffinity (JH binding protein) is also present in cytosol and nucleus of fat body (Shem-shedini & Wilson 1993). A JH binding protein has been reported in the ovary (vanMellaert et al. 1985), but it is not known how it is related to hJHBP. We found a pro-tein that is immunologically related to hJHBP that appears in fat body and the cyto-sol of ovarian protein body of H. cunea. This protein is assumed to be a JHBP. Moreinformation on the relationship between hJHBP and JHBP in the cytosol is requiredto better understand the transport of JH from secretory cells to target tissues and re-ceptors.

ENDNOTE

Mention of a trademark, warranty, proprietary product, or vendor does not consti-tute a guarantee by the U.S. Department of Agriculture and does not imply its ap-proval to the exclusion of other products or vendors that may also be suitable.

Figure 5. Left panel: Western blots of proteins from the hemolymph (H), JHBP (J),ovarian extracts (O) of H. cunea, the hemolymph (B) of B. mori, the hemolymph (P) ofP. americana. The JHBP bands were detected by enzyme immunoassay using anti-body against the hemolymph JHBP. In three lanes (H,J,O), a single Mr 32,000 proteincorresponding to the JHBP was detected by the antibody. Right panel: Immunodiffu-sion test to determine the relationship of several antigens (J, JHBP, 6 µg protein; Ov,ovarian extracts, 1.8 mg protein; H, hemolymph of H. cunea, 1 mg protein; B,hemolymph of B. mori 1.4 mg protein; L, lipophorin of H. cunea, 20 µg protein; P,hemolymph of P. americana, 1.2 mg protein; Ab, antibody against JHBP inhemolymph of H. cunea).


REFERENCES CITED

DE KORT, C. A. D., AND N. A. GRAGER. 1981. Regulation of the juvenile hormone titer.Annu. Rev. Entomol. 26: 1-28.

Figure 6. Immunoelectron micrographs of fat body (A) and ovary (B) from H. cu-nea. Immunogold particles were observed in large protein bodies (PB). Bar length =0.2 µm (A), 0.5 µm (B).


DE KORT, C. A. D., M. G. PETER, AND A. B. KOOPMANSCHAP. 1983. Binding and deg-radation of juvenile hormone III by hemolymph proteins of the Colorado potatobeetle: a re-examination. Insect Biochem. 13: 481-487.

DILLWITH, J. W., S. D. MANE, AND G. M. CHIPPENDALE. 1985. High affinity juvenilehormone binding protein of the hemolymph of the southwestern corn borer, Di-atraea grandiosella. Characteristics and relation to diapause. Insect Biochem.15: 233-246.

ENGELMANN, F. 1981. The identification of juvenile hormone binding protein in the fatbodies of Leucophaea maderae, pp. 263-270 in G. E. Pratt and G. T. Brooks[eds.], Juvenile Hormone Biochemistry, Elsevier North Horland Biomedical,New York.

FERKOVICH, S. M., H. OBERLANDER, AND R. R. RUTTER. 1977. Release of juvenile hor-mone binding protein by fat body of the Indian meal moth, Plodia interpunc-tella, in vitro. J. Insect Physiol. 23: 297-302.

KING, L. E., AND S. S. TOBE. 1988. The identification of an enantioselective JH IIIbinding protein from the hemolymph of the cockroach, Diploptera punctata. In-sect Biochem. 18: 793-805.

KLAGES, G., AND H. EMMERICH. 1979. Juvenile hormone binding protein in thehemolymph of third instar larvae of Drosophila hydei. Nature 286: 282-285.

KOEPPE, J. K., R. C. RAYNE, M. D. SHEARIN, D. J. CARVER, E. A. WHITSEL, AND R. C.VOGLER. 1988. Synthesis and secretion of a precursor hemolymph juvenilehormone-binding protein in the adult female cockroach Leucophaea maderae.Insect Biochem. 18: 661-666.

KOEPPE, J. K., R. C. RAYNE, E. A. WHITSEL, AND M. E. POOLER. 1987. Synthesis andsecretion of juvenile hormone-binding protein by fat body from the cockroachLeucophaea maderae: protein a immunoassay. Insect Biochem. 17: 1027-1032.

KOOPMANSCHAP, A. B., AND C. A. D. DE KORT. 1988. Isolation and characterization ofa high molecular weight JH III transport protein in the hemolymph of Locustamigratoria. Arch. Insect Biochem. Physiol. 7: 105-118.

KRAMER, K. J., P. E. DUNN, R. C. PETERSON, H. L. SEBALLOS, L. SANBURG, AND J. H.LAW. 1976. Purification and characterization of the carrier protein for juvenilehormone from the hemolymph of the tobacco hornworm Manduca sexta Jo-hannson (Lepidoptera: Sphingidae). J. Biol. Chem. 251: 4979-4985.

LAEMMLI, U. K. 1970. Cleavage of structural proteins during assembly of head of bac-teriophage T4. Nature 227: 680-685.

LENZ, C. J., J. W. DILLWITH, AND G. M. CHIPPENDALE. 1986. Comparison of some prop-erties of the high juvenile hormone binding protein from the larval hemolymphof pyralid moth. Arch. Insect Biochem. Physiol. 3: 61-73.

NOWOCK, J., B. HAMMOCK, AND L. I. GILBERT. 1976. The binding protein as a modu-lator of juvenile hormone stability and uptake, pp. 354-372 in L. I. Gilbert [ed.],The Juvenile Hormone, Plenum Press, New York.

OUCHTERLONY, O. 1949. Antigen-antibody reactions in gels. Acta Path. Microbiol.Scan. 26: 507-515.

OZYHAR, A., J. R. WISNIEWSKI, F. SEHNAL, AND M. KOCHMAN. 1983. Age dependentchanges in the binding and hydrolysis of juvenile hormone in the hemolymphof last instar larvae of Galleria mellonella. Insect Biochem. 13: 435-441.

PRESTWICH, G. D., AND J. K. ATKINSON. 1990. Rapid purification and N-terminalamino acid sequence of a photoaffinity-labeled juvenile hormone binding pro-tein from an arctiid moth larva, Platyprepia virginalis. Insect Biochem. 20:801-807.

RAYNE, R. C., AND J. K. KOEPPE. 1988. Relationship of hemolymph juvenile hormonebinding protein to lipophorin in Leucophaea maderae. Insect Biochem. 18:667-673.

RUDNICA, M., F. SEHNAL, V. JAROLIN, AND M. KOCHMAN. 1979. Hydrolysis and bind-ing of the juvenile hormone in the hemolymph of Galleria mellonella. InsectBiochem. 9: 569-575.


SHEMSHEDINI, L., AND T. G. WILSON. 1993. Juvenile hormone binding proteins in lar-val fat body nuclei of Drosophila melanogaster. J. Insect Physiol. 39: 563-569.

TOWBIN, H., T. STAEHELIN, AND J. GORDON. 1979. Electrophoretic transfer of proteinsfrom polyacrylamide gels to nitrocellulose sheets: procedure and some applica-tions. Proc. Natl. Acad. Sci. USA 76: 4350-4355.

VAN MELLAERT, H., S. THEUNIS, AND A. DE LOOF. 1985. Juvenile hormone bindingproteins in Sarcophaga bullata hemolymph and vitellogenic ovaries. InsectBiochem. 15: 655-661.

WEBER, K. AND M. OSBORN. 1969. The reliability of molecular weight determinationby dodecyl sulfate-polyacrylamide gel electrophoresis. J. Biol. Chem. 244:4406-4412.

WHITMORE, E., AND L. I. GILBERT. 1972. Haemolymph lipoprotein transport of juve-nile hormone. J. Insect Physiol. 18: 1153-1167.

WRAY, W., T. BOULIKAS, V. P. WRAY, AND R. HANCOCK. 1981. Silver staining of pro-

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

teins in polyacrylamide gels. Anal. Biochem. 118: 197-203.

BA & Phillips: Fire Ant Yolk Sphere Degradation

463

DEGRADATION OF RED IMPORTED FIRE ANT (HYMENOPTERA: FORMICIDAE) YOLK SPHERES

A

MADOU

S. BA

AND

S

HERMAN

A. P

HILLIPS

, J

R

.Department of Plant and Soil Science

Texas Tech UniversityLubbock, TX 79409-2134

A

BSTRACT

Transmission electron microscopy reveals that the eggs of the red imported fireant,

Solenopsis invicta

Buren, contain yolk spheres that are evenly distributedthroughout the center of the egg. Yolk degradation occurs sequentially as evidencedby the simultaneous observation of three stages of yolk platelet degradation withinthe eggs.

Key Words: Insecta,

Solenopsis invicta

, egg, oogenesis.

R

ESUMEN

La microscopía electrónica de transmisión revela que los huevos de la hormiga defuego

Solenopsis invicta

Buren, contienen esferas de yema que se distribuyen pareja-mente a lo largo del centro del huevo. La observación simultánea de tres etapas de de-gradación en las esferas de yema dentro de los huevos evidencia que la degradación

de la yema ocurre secuencialmente.

The most abundant subcellular organelles of mature oocytes in arthropods areyolk granules, constituting approximately 80% of the total protein present (Medina etal. 1988). Arthropod yolk spheres (granules or platelets) are phosphoglycolipoproteinswith varying amounts of phosphorus, lipids, and carbohydrates (Yamashito & Indra-

464



sith 1988). Much of the yolk material consists of vitellin, a protein that is derived fromvitellogenin (Kunkel & Nordin 1985; Tata & Smith 1979; Wallace 1985). Vitellin,which comprises 60-90% of the total mass of yolk proteins (Kunkel & Nordin 1985), istransported through insect haemolymph and accumulates as yolk granules in the de-veloping oocyte by pinocytosis (Postlethwait & Giorgi 1985). The yolk protein compo-sition of numerous arthropods, including many orders of the class Insecta, have beendetermined (Bownes & Hames 1977; Yamashita & Indrasith 1988). For example, yolkspheres of the fruit fly (

Drosophila

) constitute about 80% of the total protein of themature oocyte (Vallejo et al. 1981). However, the degradation of yolk in arthropods isnot a process that is well understood.

The red imported fire ant,

Solenopsis invicta

Buren, was introduced into theUnited States from South America around 1940 and has rapidly become a major pestin the southern U.S.A. Although much is known of its basic biology and behavior, rel-atively little attention has been given to oogenesis, vitellogenesis, and embryogenesis.However, one study indirectly addressing these subjects was initiated to understandthe mode of action of the insect growth regulator fenoxycarb on the red imported fireant (Glancey & Banks 1988). Fenoxycarb caused a lack of cell differentiation involv-ing oocytes, trophocytes, and follicular epithelial cells. In addition, because nurse cellsdid not develop, oocytes did not contain the necessary yolk spheres. Therefore, eggsdid not develop, and the colony died due to lack of worker replacement (Glancey &Banks 1988). In that study, the sequestration of yolk in untreated eggs and the lackof yolk spheres in treated eggs with concomitant egg reabsorption were clearly dis-played. A review of the literature reveals that study as the only one portraying yolksphere formation in oocytes of the red imported fire ant. Our paper, herein, is the firstto describe the appearance of naturally occurring yolk degradation in fertilized eggsof ants, and in particular, of the red imported fire ant.

M

ATERIALS

AND

M

ETHODS

Red imported fire ant colonies were collected from Abilene and Victoria, Texas, inthe spring of 1991 and maintained at approximately 28

°

C. Oviposited eggs were re-moved from colonies with a camel hair brush after the ants were immobilized by car-bon dioxide. Samples were fixed for three hours at room temperature in 2%glutaraldehyde, 0.1 M phosphate buffer (pH 7.0) and several drops of the wettingagent Triton-X. Eggs were subsequently washed in 0.1 M phosphate buffer (pH 7.0)and post-fixed with osmium tetroxide. Eggs were next washed in 0.1 M phosphatebuffer (pH 7.0) and dehydrated through a graded ethanol series. To remove the etha-nol, specimens were rinsed twice with 100% acetone and stored over a dehydratingmaterial (CuSO

4

). Eggs were embedded in Spurr’s

low viscosity medium (Spurr 1969).Sorvall MT-2B and glass knives were used for ultramicrotomy. Sections were stainedwith methanolic uranyl acetate and lead citrate, and micrographs were taken usinga Hitachi HS-9 transmission electron microscope.

R

ESULTS

AND

D

ISCUSSION

Electron micrographs showed the eggs to be filled with large and dense storage or-ganelles (yolk spheres). Yolk platelets were distributed evenly throughout the centerof eggs, whereas the periphery of eggs were relatively free of yolk granules (Fig. 1B).However, early embryonic cells were located at the periphery of the egg near the vi-telline membrane (Fig. 1A and B). The yolk is thus somewhat initially separated fromtissue of the developing embryo. We observed three categories of yolk granules: 1)

BA & Phillips: Fire Ant Yolk Sphere Degradation

465

large, membrane bound, undegraded dense yolk spheres; and 2) partially degradedand 3) degraded yolk spheres (Fig. 1C). Not all yolk platelets in red imported fire anteggs are degraded at the same time, but rather, yolk sphere utilization proceeds se-quentially.

Our cytological observations of yolk degradation (Fig. 1C) in red imported fire anteggs support earlier studies showing sequential yolk degradation in other arthropods.For example, proteinase activity is involved in yolk granule organelle degradationduring embryogenesis in

Drosophila

, and this activity is developmentally regulated(Medina et al. 1988). Fagotto (1991) states that yolk spheres of the eggs of an Africansoft tick (

Ornithodoros moubata

) are dense and neutral and, that during later stages,these spheres are degraded through acidification. Furthermore, acidic protease activ-ity is responsible for yolk degradation in the migratory locust,

Locusta migratoria

(McGregor & Loughton 1974). Although Glancey & Banks (1988) observed less yolkduring reabsorption of developing oocytes in red imported fire ant queens after treat-

Figure 1. Transmission electron micrographs of cross sections of red imported fireant eggs showing sequential degradation of yolk spheres: us = undegraded yolkspheres; ps = partially degraded yolk spheres; ds = degraded yolk spheres; ex = exo-chorion; en = endochorion; vtm = vitellin membrane (Fig. 1A, bar = 36 µm; Fig. 1B, bar= 50 µm; Fig 1C, bar = 52 µm; Fig. 1D, bar = 67 µm).

466



ment with an insect growth regulator, proteinase is inactive during those stages of eggformation. Therefore, normal yolk degradation would not be observable. Proteinase isonly activated in the egg subsequent to fertilization (Medina et al. 1988).

Although common mechanisms of yolk degradation may exist among evolution-arily distant species (Fagotto 1991), the assumption should not be made that thesedegradation mechanisms are applicable to other systems (Yamashita & Indrasith1988). However, the general mechanism for yolk degradation in many insects hasbeen shown to be sequentially controlled (Yamamoto & Takahashi 1993), and proteinswithin these yolk spheres are differentially hydrolyzed.

A

CKNOWLEDGMENT

This study was supported by the Texas State Line Item for Fire Ant Research. Wethank H. Thorvilson, E. White, and C. Landry for reviewing our manuscript.

R

EFERENCES

C

ITED

B

OWNES

, M.,

AND

B. D. H

AMES

. 1977. Accumulation and degradation of three majoryolk proteins in

Drosophila melanogaster

. J. Exp. Zool. 200: 149-156.F

AGOTTO

, F. 1991. Yolk degradation in tick eggs: III. Developmentally regulated acid-ification of the yolk spheres. Develop. Growth and Differ. 33: 57-66.

G

LANCEY

, B. M.,

AND

W. A. B

ANKS

. 1988. Effect of the insect growth regulator fenox-ycarb on the ovaries of queens of the red imported fire ant (Hymenoptera: For-micidae). Ann. Entomol. Soc. America 81: 642-648.

K

UNKEL

, J. G.,

AND

J. H. N

ORDIN

. 1985. Yolk proteins, pp. 83-111

in

G. A. Kerkut andL. I. Gilbert [eds.] Comprehensive Insect Physiology, Biochemistry and Phar-macology, Vol. 1. Pergamon Press, Oxford.

M

C

G

REGOR

, D. A.,

AND

B. G. L

OUGHTON

. 1974. Yolk protein degradation during em-bryogenesis of the African migratory locust. Canadian J. Zool. 52: 907-917.

M

EDINA

, M., P. L

EON

,

AND

C. G. V

ALLEJO

. 1988.

Drosophila

cathepsin B-like protein-ase: A suggested role in yolk degradation. Arch. Biochem Biophysics 263: 355-363.

P

OSTLETHWAIT

, J. H.,

AND

F. G

IORGI

. 1985. Vitellogenesis in insects, pp. 85-126

in

W.L. Browder [ed.]. Developmental Biology: A Comprehensive Synthesis, Vol. 1:Oogenesis. Plenum Press, New York.

S

PURR

, A. R. 1969. A low-viscosity epoxy resin medium for electron microscopy. J. Ul-trastructure Res. 26: 31-43.

T

ATA

, J. R.,

AND

D. F. S

MITH

. 1979. Vitellogenesis: A versatile model for hormonal reg-ulation of gene expression. Rec. Prog. Horm. Res. 35: 49-95.

V

ALLEJO

, C. G., R. P

ERONA

, R. G

ARESSE

,

AND

R. M

AROCO

. 1981. The stability of theyolk granules of

Artenia

. An improved method for their isolation and study. CellDiff. 10: 345-356.

W

ALLACE

, R. W. 1985. Vitellogenesis and oocyte growth in nonmammalian verte-brates, pp. 127-177

in

W. L. Browder [ed.]. Developmental Biology: A compre-hensive synthesis, Vol. 1: Oogenesis. Plenum Press, New York.

Y

AMASHITA

, O.,

AND

L. S. I

NDRASITH

. 1988. Metabolic fates of yolk proteins duringembryogenesis in arthropods. Develop. Growth and Differ. 30: 337-346.

Y

AMAMOTO

, Y.,

AND

S. Y. T

AKAHASHI

. 1993. Cysteine proteinase from

Bombyx

eggs:role in programmed degradation of yolk proteins during embryogenesis. Comp.Biochem. Physiol. 106B: 35-45.

Moraes et al.: Salivary Gland Cultures

467

ORGAN CULTURE OF SALIVARY GLANDS OF MALE

ANASTREPHA SUSPENSA

(DIPTERA:TEPHRITIDAE)

R

EJANE

R. D

E

M

ORAES

1

, J

AMES

L. N

ATION

AND

J

AMES

E. M

ARUNIAK

Department of Entomology and Nematology Hull Road, Building 970University of FloridaGainesville, FL 32611

1

Fellow of National Research Council (CNPq). Brasilia, Brazil

A

BSTRACT

Salivary glands from male Caribbean fruit flies,

Anastrepha suspensa,

were iso-lated individually by dissection and cultured in Schneider’s medium containing 10%Fetal Bovine Serum and antibiotics. A total of 103 salivary glands were successfullycultured. Two different procedures were used to evaluate viability. A trypan blue dyetest revealed that no more than 10% of the cultured cells were non-viable, since theyacquired blue coloration. In the second set of experiments, acid phosphatase activityof the cultured glands was measured colorimetrically from absorbance at 415 nm ofliberated p-nitrophenol. This method indicated that the cultured glands stayed met-abolically active, with enzyme activity equal to that of freshly excised glands on thefifth, eighth, and fifteenth days of the experiment. Scanning electron microscopy(SEM) showed no visual differences in tissue organization and size of individual cellswhen freshly excised salivary glands and cultured glands were compared. SEMshowed that the cells of cultured glands were neither swollen nor shrunken, andclose-up views showed no evidence of cell deterioration or lysing.

Key Words: Salivary glands, organ culture, Caribbean fruit fly, Tephritidae,

Anas-trepha suspensa

, acid phosphatase.

R

ESUMEN

Las glándulas salivares de los machos de la mosca caribeña de las frutas,

Anas-trepha suspensa

, fueron aisladas individualmente por disección y cultivadas en mediode Schneider que conteniene 10% de suero fetal bovino y antibióticos. Un total de 103glándulas salivares fueron cultivadas con éxito. Dos procesos diferentes fueron usa-dos para evaluar la viabilidad. Una prueba de tinción de trypan azul reveló que menosdel 10% de las células cultivadas no era viable porque adquirían una coloración azul.En una segunda serie de experimentos, la actividad de fosfatasa ácida de las glándu-las cultivadas fue estudiada colorimétricamente mediante la absorbancia de p-nitro-fenol liberado a 415 nm. Este método indicó que las glándulas cultivadas permanecíanmetabólicamente activas, con actividad enzimática en los días quinto, octavo y déci-moquinto del experimento igual a la de las glándulas recién extirpadas. La Microsco-pía Electrónica de Barrido (MEB) no mostró diferencias visuales en la organización delos tejidos y en el tamaño de las céllulas individuales, cuando fueron comparadas lasglándulas frescas y las cultivadas. La MEB mostró que las células de las glándulascultivadas no estaban hinchadas ni contraídas, y las vistas tomadas de cerca no evi-

denciaron deterioración o rompimiento celular.

Organ culture is a technique in which whole organs or representative parts aremaintained as tissues and retain their intrinsic distribution, numerical and spatial

468



orientation of explanted cells (Freshney 1987). Most organ cultures do not grow or, ifthey do, proliferation is limited to the outer cell layers. Thus, organ culture is essen-tially a technique for studying the behavior of integrated tissues rather than isolatedcells.

The in vitro culture of insect organs has made much progress in the last 30 years.This success is due mainly to the development of adequate tissue culture media by sci-entists, e.g., Schneider (1964). Insect organ culture has been associated primarilywith insect endocrinological studies (Marks 1976). In vitro cultures of imaginal discsof

Drosophila melanogaster

Meigen (Raikow & Fristrom 1971),

Sarcophaga peregrina

(Robineau-Desvoidy) (Ohmori & Ohtaki 1972),

Plodia interpunctella

(Hubner) (Ober-lander 1976) and others have aided in the understanding of the role of20-hydroxyecdysone and ecdysone in development and metabolic activity. Organ cul-tures of chironomid salivary glands have been extensively utilized in investigations ofpolytenic cell differentiation, cytogenetics, biochemistry and physiology (Firling &Kobilka 1978; Firling & Hou 1980).

The salivary glands in

Anastrepha suspensa

(Loew) are sexually dimorphic, withmales having much enlarged glands that terminate on each side of the body in a largeball of convoluted tubules in the pleural region of abdominal segments 3, 4 and 5 (Na-tion 1974, 1981). The glands in males reach maximum size at approximately the sametime that males are maximally active in sexual behavior and in producing a phero-mone that attracts females (Nation 1972). Pheromonal components have been de-tected in dissected salivary glands by gas chromatographic analysis (Nation 1989),but conclusive evidence that the glands participate in pheromone production is lack-ing. Organ cultures of the salivary glands might be used to determine whether or notthey are involved in pheromone production, if a culture technique for these glandswas available. For example, Srinivasan et al. (1979) successfully cultured sex phero-mone glands of

P. interpunctella

for 10 days and were able to recover sex pheromonefrom the culture medium.

The objectives of this study were to establish a protocol for sterile dissection of sal-ivary glands from male Caribbean fruit flies; to maintain salivary glands in artificialmedium for at least seven days; and to establish protocols for monitoring the viabilityof cultured glands.

M

ATERIALS

AND

M

ETHODS

Male

A. suspensa

were obtained from the mass rearing facility at the Florida StateDepartment of Agriculture and Consumer Services, Gainesville, FL. Organ culturesof the male salivary glands were established in four separate experiments after dis-secting the glands from 7- to 15-day-old males. All equipment and tools used to dissectthe glands were sterilized. Microscopes, lamps and wax dishes were sterilized with70% ethanol and left under a UV lamp for 20-30 min before dissections began. Scis-sors and forceps were autoclaved before each series of dissections and flamed aftereach fly was dissected. Glands were dissected under sterile conditions in a sterilehood. Male flies were surface-sterilized by immersion in 70% ethanol for 10 min andrinsed in sterile saline solution for 10 min. Just prior to each dissection, the head andlegs of each fly were removed to diminish movement and to facilitate salivary glandremoval. Each fly was pinned ventral side up in a wax dish containing sterile salinesolution. A longitudinal cut was made through the cuticle and the lateral sides of theabdomen were gently pressed to release the salivary glands, which were then care-fully excised. The excised glands were placed in sterile saline solution until a seriesof dissections were complete, then they were transferred to the culture medium.


469

Organ Culture

Schneider’s medium (Schneider, 1964) containing 10% Fetal Bovine Serum andthe antibiotics penicillin (100 units per ml), streptomycin (100

µ

g per ml), amphoteri-cin B (0.2

µ

g per ml), gentamycin (50

µ

g per ml), geocillin (100

µ

g per ml), kanamycin(100

µ

g per ml) and tetracycline (10

µ

g per ml) were used to culture the salivaryglands. [Schneider’s medium has been used successfully to culture

Drosophila

cells(Schneider & Blumenthal, 1978).] A 24-cluster-well tissue culture plate (Corning) wasused to culture the glands with 0.5 ml medium and two salivary glands per well. Thesalivary gland cultures were kept in an incubator at 27

°

C. Four experiments wereconducted for eight to 15 days; the medium was replaced every three days in experi-ments 2 to 4.

Viability Tests

In the first experiment, penetration of trypan blue dye was used as an indicator ofviability of the cultured glands, based on the principle that dead cells allow the dye topenetrate while live cells exclude the dye. Each day during one week of culture, 250

µ

l of trypan blue was added to the culture medium in one of the wells. Penetration orexclusion of the dye in the 2 glands was determined by microscopic examination. Inexperiments 2, 3 and 4, viability was monitored by determining the activity of acidphosphatase on the fifth, eighth and (only in experiment 4) on the fifteenth day. Oneach of these days, about 10 glands were removed from culture, rinsed with saline so-lution, and homogenized in 250

µ

l of 0.1 M citrate buffer, pH 5.5. Seventy

µ

l of theglandular homogenate was added to each of two tubes containing 1 ml of 7 mMp-nitrophenol phosphate in 0.1 M citrate buffer, pH 5.5. A third tube served as a con-trol and contained only citrate buffer. Fresh glands were dissected from flies that werethe same age as the cultured glands, and these fresh glands were homogenized as de-scribed above for the cultured glands. Assay tubes identical to those for culturedglands were prepared to receive 70

µ

l of the fresh gland homogenate positive controls.All tubes were incubated at 30

°

C for 10 min. The reaction was stopped by adding 2 mlof 10% Na

2

CO

3

to each tube. This altered the pH, stopped acid phosphatase activityand caused a shift in the electron structure of the released p-nitrophenol so that itgave a yellow color and absorbed light strongly at 415 nm. The amount ofp-nitrophenol released was determined at 415 nm in a Spectronic 20D colorimeter. Astandard curve was prepared from pure p-nitrophenol, and final results were ex-pressed as pg p-nitrophenol released per gland per h.

The above protocol was changed slightly for the fourth experiment as follows. Thesubstrate was dissolved in 0.1 M citrate buffer (pH 5.5) just before use. Incubationtubes contained 2 ml of 7 mM p-nitrophenol phosphate and 0.1 ml of glandular homo-genate. Tubes were incubated for 20 minutes at 30

°

C. The reaction was stopped byadding 3 ml of 10% Na

2

CO

3

solution. The control contained 7mM p-nitrophenol in ci-trate buffer.

Scanning Electron Microscopy

Freshly dissected and cultured salivary glands were fixed for about 24 h in Bouin’sfixative, washed in 70% ethanol to remove excess picric acid, and dehydrated throughan alcohol series to 100% ethanol. Glands were transferred from 100% ethanol to hex-amethyldisilasane (HMDS) (Nation 1983), transferred once to fresh HMDS and airdried. The dried tissues were mounted on stubs, gold coated with a sputter-coater, andobserved in a Hitachi S 570 instrument with filament voltage at 15 or 20 KV.

470



R

ESULTS

AND

D

ISCUSSION

A total of 103 excised salivary glands from adult male flies were successfully cul-tured in isolated organ cultures for up to 1 week, and, in one experiment, up to 15days. Scanning electron microscopy (SEM) showed no visual differences in tissue or-ganization and size of individual cells between freshly excised salivary glands andglands cultured for 8 days (Fig. 1). Cells in the cultured glands did not shrink, collapseor lyse. Each of the bilateral salivary glands has the appearance of a cluster of grapes,and each “grape” is a cell enclosing a cavity, with cells interconnected by a central ca-nal (Nation 1974). The SEM was useful in viewing the surface appearance of thegland cells and in ascertaining that the individual cells were not swelling, shrinking,or lysing.

Cultured glands excluded trypan blue dye, an indication that the cells of theglands were alive and apparently functioning normally. Some blue cells were ob-served on the periphery of the glands, but their percentage was never greater than10%. In a preliminary experiment to measure acid phosphatase activity, we foundthat 57 pg of p-nitrophenol per gland per h was released by glands cultured for 5 dayspost-dissection, and 61 pg was released by fresh glands of comparable age to the cul-tures. After 8 days of culture, 56 pg per gland per h of p-nitrophenol was released bycultured glands and 55 pg per gland per h by fresh glands of comparable age. The con-trol was changed to contain 7mM p-nitrophenol in citrate buffer following this prelim-inary experiment and subsequent results are shown in Table 1. Those data show thatacid phosphatase activity of freshly excised glands and cultured glands did not differin glands that were 5, 8 or 15 days old. The substrate, p-nitrophenol phosphate, addedto the standard caused a slight yellow color, lowering the values in Table 1. However,these values more accurately represent the true enzyme activity than the preliminarydata above.

The surface sterilization of adult flies, and the antibiotics in the culture mediumwere effective in maintaining sterility, since only one preparation showed evidence ofcontamination by becoming cloudy and discolored. Contamination also could be re-duced by submerging adult flies in 70% ethanol. For example, Ohmori & Ohtaki(1972) surface-sterilized larvae of

S. peregrina

by submerging them in 70% ethanolfor several min prior to isolating their wing discs for organ cultures, and Oberlander& Leach (1979) surfaced sterilized larvae of

P. interpunctella

(Hubner) by submergingthem in 70% ethanol for 20 min prior to dissecting imaginal discs for organ cultures.Schneider’s medium, which was developed for

Drosophila

imaginal disc cultures(Schneider 1972), was adequate for culturing the glands of male Caribbean fruit flies.

The exclusion of trypan blue and the activity of the enzyme, acid phosphatase, ap-pear to be useful indicators of viability in salivary gland cultures. Enzyme activityand protein synthesis have often been used to measure the viability of cultured organsand cells. Firling & Kobilka (1978) measured the ability of cultured

Chironomus ten-tans

(Fabricius) salivary glands to incorporate

14

C-leucine into trichloroacetic acid(TCA) precipitable proteins. Acid phosphatase as well as amylase and protease activ-ity were monitored in cultured salivary glands of

Calliphora

larvae (Price 1974). Or-gan cultures of

C. tentans

salivary glands were capable of excluding a solution of0.12% trypan blue, while injured cells did not exclude the dye (Firling & Kobilka1978). The few blue cells observed on the surface of the male salivary glands in thepresent experiments may well have been injured in the dissection or handling proce-dures, thus causing them to leak and allow uptake of trypan blue dye. Monitoring fortrypan blue dye uptake is easier than measuring acid phosphatase activity and mightbe the method of choice in any future experiments with salivary glands from fruitflies.


471

A

CKNOWLEDGMENTS

We thank Kathy Milne for technical assistance in the laboratory, Drs. HerbertOberlander and S. M. Ferkovich for helpful criticism of earlier drafts, and the Na-tional Research Council-CNPq, Brazil, for financial support to R. R. de Moraes. Flor-ida Agricultural Experiment Station Journal Series No. R-04079.

Figure 1. Scanning electron microscopy comparing cultured (8-days-old) andfreshly excised male salivary glands (from 8-day-old males) from Caribbean fruit flies(Anastrepha suspensa). A and B, 150 × magnification; C and D, 500 × magnification.

472



R

EFERENCES

C

ITED

F

IRLING

, C. E.,

AND

B. K

OBILKA

. 1979. A medium for the maintenance of

Chironomustentans

salivary glands in vitro. J. Insect Physiol. 25: 93-103.F

IRLING

, C. E.,

AND

A. H

OU

. 1980. Ecdysterone induced regression of

Chironomus

sal-ivary glands maintained in vitro, p. 187-193

in

E. Kurstak, K. Maramorosch,and A. Dubendorfer [eds.], Invertebrate systems in vitro. Elsevier/North Hol-land Biomedical Press, New York, NY. 598p.

F

RESHNEY

, R. I. 1987. Culture of animal cells: A manual of basic technique. 2nd ed.Alan R. Liss, Inc, New York, NY. 397pp.

M

ARKS

, E. P. 1976. The uses of cell and organ cultures in insect endocrinology,

in

E.Kurstak, and K. Maramorosch, [eds.], Invertebrate tissue culture, applicationsin medicine, biology, and agriculture. Academic Press, New York. 398pp.

N

ATION

, J. L. 1972. Courtship behavior and evidence for a sex attractant in the maleCaribbean fruit fly,

Anastrepha suspensa

. Ann. Entomol. Soc. America. 65:1364-1367.

N

ATION

, J. L. 1974. The structure and development of two sex specific glands in maleCaribbean fruit flies. Ann. Entomol. Soc. America. 67: 731-734.

N

ATION

, J. L. 1981. Sex-specific glands in tephritid fruit flies of the genera

Anas-trepha

,

Ceratitis

,

Dacus

, and

Rhagoletis

(Diptera: Tephritidae). Int. J. InsectMorphol. & Embryol. 10: 121-129.

N

ATION

, J. L. 1983. A new method using hexamethyldisilazane for preparation of softinsect tissues for scanning electron microscopy. Stain Technology. 58:347-351.

N

ATION

, J. L. 1989. The role of pheromones in the mating system of

Anastrepha

fruitflies, pp. 189-205

in

A. S. Robinson and G. Hooper, [eds], Fruit Flies Their Bi-ology, Natural Enemies and Control, Vol. 3A, Elsevier Press.

O

BERLANDER

, H. 1976. Hormonal control of growth and differentiation of insect tis-sues cultured in vitro. In Vitro. 12: 225-235.

O

BERLANDER

, H.,

AND

C. E. L

EACH

. 1979. The action of

β

-ecdysone and juvenile hor-mone in organ cultures of lepidopteran imaginal disks. TCA Manual. 5: 993-995.

O

HMORI

, K.,

AND

T. O

HTAKI

. 1973. Effects of ecdysone analogues on development andmetabolic activity of wing disks of the fleshfly,

Sarcophaga peregrina

, in vitro.J. Insect Physiol. 19: 1199-1210.

PRICE, G. M. 1974. Protein Metabolism by the salivary glands and other organs of thelarva of the blow fly, Calliphora erythrocephala. J. Insect Physiol. 20: 329-347.

RAIKOW, R., AND J. W. FRISTROM. 1971. Effects of β-ecdysone on RNA metabolism ofimaginal disks of Drosophila melanogaster. J. Insect Physiol. 17: 1599-1614.

SCHNEIDER, I. 1964. Differentiation of larval Drosophila eye-antennal discs in vitro.J. Exp. Zool. 156: 91-104.

TABLE 1. COMPARISON OF PHOSPHATASE ACTIVITY BETWEEN CULTURED AND FRESHLYEXCISED SALIVARY GLANDS.

Salivary GlandsP-Nitrophenol

pg per Gland per Hour

Mean1 ± SD

5 Days 8 Days 15 Days

Cultured 7.36 ± 0.21 7.26 ± 0.23 6.69 ± 0.02Fresh 8.38 ± 0.11 8.49 ± 1.57 7.32 ± 0.32

1All values are the mean of two replicates except at 8 days, when four replicates were analyzed.

Moraes et al.: Salivary Gland Cultures 473

SCHNEIDER, I. 1972. Cell lines derived form late embryonic stages of Drosophila mel-anogaster. J. Embryol. Exp. Morphol. 27: 353-365.

SCHNEIDER, I., AND A. B. BLUMENTHAL. 1978. Drosophila cell and tissue culture, pp.206-315 in M. Ashburner, and T. R. F. Wright, [eds], Genetics and Biology ofDrosophila, Vol 2A, Academic Press, New York.

SRINIVASAN, A., J. A. COFELT, AND H. OBERLANDER. 1979. In vitro maintenance of thesex pheromone gland of the female indian meal moth Plodia interpunctella

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

(Hubner). J. Chem. Ecol. 5: 653-661.

Shelly & Villalobos: Melon Fly Behavior

473

CUE LURE AND THE MATING BEHAVIOR OF MALE MELON FLIES (DIPTERA: TEPHRITIDAE)

T

ODD

E. S

HELLY

1

AND

E

THEL

M. V

ILLALOBOS

2

1

Hawaiian Evolutionary Biology Program,University of Hawaii, Honolulu, HI 96822

2

Biology Department,Chaminade University, Honolulu, HI 96816

A

BSTRACT

Laboratory tests were conducted to assess the effect of the parapheromone cuelure on the mating behavior of male

Bactrocera cucurbitae

(Coquillett). Exposure tocue lure resulted in a short-term mating advantage. For wild flies, treated males thatfed on cue lure on the day of testing, or 1 day prior to testing, mated more frequentlythan control males that had no prior exposure to cue lure. However, control andtreated males had similar mating success in tests performed 3 or 7 days after thetreated males were exposed to the lure. Exposure to cue lure also increased the mat-ing success of mass-reared, irradiated males relative to unexposed wild males, thoughthis advantage was evident for only 1 day following exposure. Cue lure appeared toenhance mating performance by increasing male wing-fanning activity but not the at-tractiveness of the signal per se. A field study revealed that irradiated males exposedto cue lure 1 week prior to release were less likely to be captured (in Steiner trapsbaited with cue lure and naled) than unexposed males. These findings suggest thatexposure of sterile males to cue lure might improve the effectiveness of sterile insectrelease as well as enable simultaneous control programs of sterile insect release andmale annihilation.

Key Words:


, parapheromone, sterile insect release, Hawaii.

R

ESUMEN

Se estudió en el laboratorio el efecto de la paraferomona “cue lure” en el compor-tamiento sexual de los machos de


(Coquillett). La exposición ala “cue lure” tuvo como resultado ventajas en los apareamientos a corto plazo. Los ma-chos salvajes que fueron alimentados con “cue lure” el mismo día, o el día anterior alexperimento se aparearon más frecuentemente que los machos del grupo control queno habían sido expuestos a la paraferomona. Sin embargo, los machos de ambos gru-pos tuvieron éxitos de copulación similares en experimentos conducidos 3 o 7 días des-pués de haber sido expuestos a la “cue lure”. La exposición a la “cue lure” de los

474



machos criados en el laboratorio e irradiados también aumentó el éxito de copulaciónen comparación con los machos salvajes que no habían recibido la paraferomona, peroesta ventaja sexual fue efectiva sólo un día después del tratamiento. Al paracer la “cuelure” aumenta la efectividad del comportamiento sexual porque estimula la actividadvibratoria de las alas, pero no afecta la calidad atractiva de la señal química en sí. Unestudio de campo indicó que los machos irradiados expuestos a la “cue lure” la semanaanerior a ser liberados fueron capturados con menor frecuencia (en trampas de Stei-ner cebadas con “cue lure” y naled) que los machos que no fueron tratados. Los resul-tados sugieren que el exponer machos estériles a la “cue lure” podría aumentar laefectividad de las liberaciones de insectos estériles, asícomo permitir el uso simultá-neo de programas de control basados en la liberación de insectos estériles y aniquila-

ción de machos.

The males of many tephritid species are strongly attracted to specific chemicalcompounds, termed male lures or parapheromones, which either occur naturally inplants or are (presumed) synthetic analogues of plant-borne substances (Chambers1977; Sivinski & Calkins 1986; Fletcher 1987). Several well-known examples includethe attraction of male Mediterranean fruit flies,

Ceratitis capitata

(Wiedemann), totrimedlure, male oriental fruit flies,

Bactrocera dorsalis

(Hendel), to methyl eugenol,and male melon flies

B. curcubitae

(Coquillett) to cue lure. Because they are powerfulattractants, parapheromones are used in current control programs of tephritid pestsfor detection and monitoring of populations and for eradication via male annihilation(Chambers 1977; Economopoulos & Haniotakis 1994).

Recent studies (Shelly & Dewire 1994; Shelly 1994) on

B. dorsalis

suggest anotherpotential use of male lures in control efforts. Data collected in the laboratory showedthat (treated) males exposed to methyl eugenol mated more frequently than (control)males that had no prior exposure to the lure. Moreover, the effect of methyl eugenolwas long-lasting, and males that fed for only 30 s on methyl eugenol had a mating ad-vantage as long as 35 days later. Enhanced mating success appeared to result from 2factors: treated males signaled (wing-fanned) more frequently and for a given level ofsignaling attracted more females per min than control males. In addition, field testsshowed that treated males were less likely to be captured (in traps baited with methyleugenol and naled) than control males for as long as 35 days after exposure. Thesestudies suggest that the effectiveness of control efforts might be enhanced by exposingsterile males to parapheromone prior to release, making it possible to combine pro-grams of male annihilation and sterile insect release.

The present study further examines the relationship between parapheromonesand sexual behavior by investigating the influence of cue lure on the mating behaviorof male

B. cucurbitae

under laboratory conditions. Using laboratory-reared wild flies,we initially tested whether exposure to cue lure enhances mating success and, if so,for how long after exposure. We then examined whether exposure to cue lure in-creased the mating competitiveness of irradiated (sterile) males relative to wild malesand, if so, for how long after exposure. As will be described, the mating trials did, infact, show that cue lure enhanced mating performance, and data were then gatheredto assess the effect of cue lure on male signaling effort and signal attractiveness. Fi-nally, to investigate the possibility of conducting sterile insect release and male anni-hilation simultaneously, we tested whether exposure to cue lure reduced theprobability of capturing (in traps baited with cue lure and naled) irradiated males fol-lowing their release in the environment.


475

M

ATERIALS

AND

M

ETHODS

Mating Behavior of

B. cucurbitae

Several workers have described the mating behavior of

B. cucurbitae

(Suzuki &Koyama 1980, 1981; Kuba & Koyama 1982, 1985; Kuba et al. 1984; Iwahashi & Ma-jima 1986; Kuba & Sokei 1988), and the following summary derives from these earlierreports. Mating occurs at dusk in male aggregations termed leks. Males perch singlyon leaf under surfaces of both host and non-host plants and defend their site againstintruding males. While perching, males fan their wings rapidly, producing a high-pitched buzzing sound. Wing-fanning also enhances dispersal of a pheromone, pro-duced in the rectal gland, attractive to females (Schultz & Bousch 1971; Kobayashi etal. 1978; Baker et al. 1982). (Because acoustic, visual, and olfactory cues may be in-volved, the terms “signaling” and “calling”, which are used synonymously with wing-fanning, refer to the composite stimulus produced by all communicative modes.) Afterseveral seconds of wing-fanning, the male pounces on the female, and copulation en-sues.

At present, the role of cue lure in male mating behavior is obscure, particularly be-cause cue lure is a synthetic compound (unlike methyl eugenol; Metcalf et al. 1975)that has not yet been recorded from any natural source (Beroza et al. 1960). However,studies by Nishida and co-workers (Nishida et al. 1990, 1993) suggest that cue lure re-sembles a natural plant-borne substance, whose metabolites are sequestered in therectal gland for pheromone synthesis. Initial work showed that males fed cue lure ac-cumulated in the rectal gland a particular ketone [4-(4-hydroxyphenyl)-2-butanone;also known as Willison’s lure and raspberry ketone] known to occur in various plantsand to be attractive to male

B. cucurbitae

. More recently, male melon flies were ob-served to feed voraciously on flowers of an orchid that contained raspberry ketone andto subsequently sequester it in the rectal gland. Though the exact role of this com-pound in female attraction remains unknown, its accumulation at the site of phero-mone production clearly suggests a role in mate attraction and courtship.

Mating Trials

Wild flies were derived from a laboratory stock started in April 1994 with 200-300adults reared from

Coccinia grandis

(L.) collected in Waimanalo, Oahu, Hawaii. Thepresent study was conducted during July-September, 1994, and consequently the flieswere 3-5 generations removed from the wild. (Though this stock may have undergonegenetic change as a result of colonization, the flies will be referred to as “wild” as ter-minological shorthand). The colony was housed in a large screen cage (1.2 m by 0.6 mby 0.6 m) with superabundant food (a mixture of honey and protein hydrolysate) andwater. Italian squash (

Cucurbita pepo

L.) was provided periodically for oviposition.Room temperature was maintained at 20-22

°

C and relative humidity at 65-75%. In-fested squash were placed in plastic buckets containing vermiculite, and larval devel-opment and pupation proceeded in situ. Adults were separated by sex within 5 daysof eclosion, well before reaching sexual maturity [at 18-20 days of age (Wong et al.1986)]. Adults were maintained in plastic buckets (5 liters volume; 40-60 individualsper bucket) covered with screen mesh and were supplied with ample food and water.

Mass-reared, irradiated flies were obtained from a colony started by the USDA/ARS Tropical Fruit and Vegetable Laboratory, Honolulu, in 1958 (H. Chang, pers.comm.) using standard rearing procedures (Tanaka et al. 1969; males from this stockwill hereafter be referred to as “sterile males”). Pupae that had been exposed to 10krad of gamma radiation from a

60

Co source were obtained 1 day before eclosion. Sexes

476



were separated within 3 days of eclosion [sexual maturity in this stock is attained atabout 8-10 days of age (Vargas et al. 1984; Wong et al. 1986)].

In the mating tests, 3 males from each of the 2 groups being tested were placedwith 3 wild females in screen cages at least 4 h before sunset. Six to 9 cages were usedon a given day. The cages were 30-cm cubes with one side open and fitted with a clothsleeve. Before placement in the cages, males from the 2 groups were cooled for severalminutes and then marked on the thorax with different colors of enamel paint. Fe-males were not marked. The cooling and painting procedures had no apparent ad-verse effects, and males resumed normal activities within minutes of handling.

Three experiments were conducted. In the first, mating success was compared be-tween wild males that had previous exposure to cue lure (treated males) and wildmales denied access to cue lure (control males). Treated males (21-25 days old) weregiven unrestricted access during a 2-h period to a cotton wick (5 cm long) to which 1.5ml of pure cue lure had been applied (cue lure obtained from the USDA/ARS Fruit andVegetable Laboratory, Honolulu). The wicks, held upright in small plastic containers,were placed singly in the plastic buckets during midday. Treated males were used 0,1, 3 or 7 days after exposure to cue lure. In an additional test, treated males were ex-posed to the wick for 2 h, but the wick was covered with screen mesh so that directcontact with it was prohibited. In these tests, control and treated males were approx-imately the same age. Females used in this and all subsequent experiments were 21-28 days old. In the second experiment, the mating frequency of wild vs. sterile maleswas compared without previously exposing either group to cue lure. In the final ex-periment, cue lure was provided to either wild or sterile males during a 2-h middayperiod 1 day or 3 days prior to testing. In the latter two experiments, males of bothgroups were used during their first 2 weeks of sexual maturity (21-30 days and 12-21days of age for wild and sterile males, respectively). In the final experiment, wild andsterile males were exposed to cue lure at 21-23 days and 12-14 days of age, respec-tively.

Signal Production and Attractiveness

To investigate the possibility that exposure to cue lure increased signal productionand attractiveness, we placed treated and control males in “minicages” within alarger flight cage and simultaneously monitored signalling by, and female attractionto, males (following the basic protocol of Poramarcom & Boake [1991]). Because wildmales called only infrequently in the mini-cages, males (and females) used in this ex-periment were from the USDA/ARS colony mentioned above. In this case, however,we used non-irradiated, “normal” flies. Sexes were separated within 3 d of eclosion.

Because single males were reluctant to call, groups of three males were placed inthe minicages. Four groups—two control and two treated—were tested on a given day.All treated males fed for 1-1.5 min on a wick containing 1.5 ml of cue lure one dayprior to use, and in all tests control and treated males were between 12-15 days old.The minicages were transparent, plastic cylinders (12 cm by 7 cm) whose ends werecovered with nylon screening.

The tests were conducted in a large screen cage (1.2 m by 0.6 m by 0.6 m) that con-tained three potted plants (

Ficus

sp.). The minicages were hung at four selectedbranches on these plants, and the individual cages were placed randomly at these lo-cations for each observation day. On a given day, 45 mature virgin females (12-15 daysof age) were placed in the cage 4-6 h before sunset. At the same time, the minicages(containing males) were placed at their assigned locations on the plants. Room lightswere then extinguished, and the flight cage, which was near a west-facing window, re-ceived only natural light.


477

Starting 45 min before sunset, we checked the individual minicages at 1-min in-tervals and recorded the numbers of wing-fanning males and females resting on theminicages. Attractiveness of male signaling was thus based on the number of femalesightings for a given minicage; because females were not marked, no data are avail-able on the number of different females that alighted on the minicages. At the end ofan observation period, individuals of both sexes were discarded, and a new set ofmales and females was used each day of testing. Observations were made on 7 days,and thus data were gathered for 14 minicages of control and treated males, respec-tively.

Capture of Sterile Males Exposed to Cue Lure

A trapping experiment was conducted on the campus of the University of Hawaiiat Manoa, Honolulu, Hawaii. The study area was a large, grassy lawn containing nu-merous ornamental trees and shrubs. Ten Steiner traps were placed singly in trees ina circle (50-m radius) around a central release point. The same plants were used in allreplicates. Traps were suspended in the canopy (2 m above ground) by a 30-cm wirefastened to a branch. Each trap contained a 5-cm long cotton wick to which 2.0 ml ofcue lure (5% naled) had been applied.

Treated sterile flies (12-15 days old) were given access to a wick containing 1.5 mlof cue lure for a 2-h period 1 day prior to release. Similarly-aged control males (18-23day old) that had no exposure to cue lure were also released. Prior to release, controland treated males were marked on the thorax with different color combinations ofenamel paint (a given combination was used only once). Flies were released between1000-1100 hours by removing the screen top and gently tapping the bucket to induceflight. Traps were checked 48 h after release, and captured flies were examined indi-vidually for markings in the laboratory. Seven replicates were conducted with 100males released per group (control or treatment) per replicate. During the study pe-riod, days were generally sunny or only partly cloudy, and ambient daylight temper-atures ranged between 25-33

°

C.

R

ESULTS

Mating trials

Cue lure conferred a short-term mating advantage (Fig. 1). In the first experiment,treated males tested on the same day they were exposed to cue lure achieved 62% (55/88) of all matings recorded (t=2.3; P < 0.05; binomial test). Similarly, treated malestested on the day following exposure obtained 67% (58/86) of the matings (t=3.4; P <0.001; binomial test). However, no difference in mating success was observed betweencontrol and treated males when treated males were tested 3 days (t=0.8) or 7 days af-ter exposure to cue lure (P > 0.05 in both cases; binomial test). Also, treated males thatwere permitted only to approach a cue lure source without contacting it did not havea mating advantage over control males. Treated males obtained only 51% (38/74) of allmatings (t=0.2; P > 0.05; binomial test).

Cue lure also influenced the relative mating competitiveness of wild and sterilemales. In the second experiment (where cue lure was not given to either wild or sterilemales), sterile males had a mating advantage over wild males and obtained 86% (75/87) of all matings (t=9.6; P < 0.001; binomial test). In the final experiment, sterile fliesexposed to cue lure 1 day prior to testing obtained 98% (93/95) of all matings, a pro-portion significantly greater than that observed in the preceding experiment (G=8.0;P < 0.01; G test with Yates correction). Wild flies exposed to cue lure 1 day before test-

478



ing also displayed increased mating success and obtained nearly 1/4 (22/95=23%) ofthe matings (about twice the proportion recorded when cue lure was not provided),though this increase was not statistically significantly (G=2.0; P < 0.20; G test withYates correction). Despite their improved performance, wild males still accounted fora disproportionately small number of matings relative to sterile males (t=6.4; P <0.001; binomial test). Exposure to cue lure 3 days prior to testing had no effect on therelative mating success of wild and sterile males. Sterile males obtained 89% (10/88;t=11.4; P < 0.001; binomial test) and 88% (70/80; t=10.0; P < 0.001; binomial test) ofall matings when wild and sterile males, respectively, were tested 3 days after expo-sure to cue lure. In both cases, the results were similar to those obtained when cuelure was not given to either group of males (P > 0.05 in both comparisons; G test withYates correction).

Signal Production and Attractiveness

Males exposed to cue lure called more frequently than control males (Fig. 2). Anaverage of 70 instances of wing-fanning (maximum value possible=135=3 males perminicage over 45 checks) was recorded for minicages with treated males compared toonly 54 instances for minicages with containing control males (U=149; P < 0.001;Mann-Whitney test). In addition, more females were sighted on minicages withtreated males than control males (Fig. 2). On average, 56 female sightings were re-corded for minicages with treated males but only 34 were observed for minicages withcontrol males (U=161; P < 0.001; Mann-Whitney test). This difference in female arriv-

Figure 1. Numbers of matings obtained by control vs. treated wild males in trialsconducted 0, 1, 3, or 7 days after treated males were exposed to cue lure.


479

als appeared to reflect the differential calling activity between control and treatedmales and not a difference in signal attractiveness per se between the two groups,since the rate at which female sightings increased with male signaling was similar forcontrol and treated males (t=0.7; P > 0.05); Fig. 2).

Capture of Sterile Males Exposed to Cue Lure

Sterile males that had been exposed to cue lure were trapped less frequently thansterile males that had no exposure to the lure. On average, only 6 treated males(range: 2-12) were captured per replicate compared to 19 control males (range: 14-28;U=49; P < 0.001; Mann-Whitney test).

D

ISCUSSION

The present study shows that, under laboratory conditions, exposure to cue lureconfers a mating advantage to male

B. cucurbitae

. However, whereas a positive effectof methyl eugenol was evident one month after exposure (Shelly & Dewire 1994), cuelure provided a mating advantage for only one day following feeding. In the melon fly,

Figure 2. Relationship between female sightings and wing-fanning for control (●)and treated (*) males. Each point represents a minicage that contained three males.Abscissa represents the total number of wing-fanning instances recorded for all 3males/minicage over the 1-min checks. Ordinate represents the total number of fe-male sightings on a minicage over the 1-min checks. Regression equations: controlmales - Y=1.05X-25.2; r2=0.75; treated males - Y=0.81X-0.6; pooled (shown) - Y=1.07X-22.6; r2=0.64.

480



ingestion of cue lure appeared to increase mating success via an increased level of sex-ual signaling. Though metabolites of cue lure (or its natural analogue) may be used forpheromone synthesis (Nishida et al. 1990, 1993), we found no evidence that the sig-nals produced by males exposed to cue lure were more attractive than those generatedby control males: for a given level of wing-fanning, similar numbers of females weresighted on minicages with control and treated males. This situation differs from thatreported for

B. dorsalis

, where (independent of signaling effort) females showed astriking preference for males that fed on methyl eugenol over control males (Shelly &Dewire 1994).

A separate study (Whittier & Shelly 1995) indicates that trimedlure has an effecton male medflies similar to that described here for cue lure and male melon flies.Though they do not feed on the chemical,

C. capitata

males exposed to trimedlure dis-played higher calling levels and mated more often than control males. However, basedon the number of female arrivals to calling males, treated males did not produce moreattractive signals than control males. Also, as with the melon fly, the positive effect ofthe lure was short-lived and was, in fact, evident only on the day that males were ex-posed to the lure.

Cue lure appears to be far less attractive to

B. cucurbitae

males than is methyl eu-genol to

B. dorsalis

males. When exposed to the lure for a 30-min period in screencages, nearly 90% of wild

B. dorsalis

males fed on methyl eugenol at their first oppor-tunity (Shelly 1994), but only 33% (22/64) of wild

B. cucurbitae

males fed on cue lureduring initial exposure (T.E.S., unpublished data). Similarly, trapping experiments(using identical procedures at the same site) with untreated, irradiated males in thetwo species showed a pronounced difference in capture probability: an average of 32%of

B. dorsalis

males were captured per replicate (Shelly, unpublished data) comparedto only 19% of

B. cucurbitae

males. The comparatively low attractiveness of cue lureis consistent with its short-lived effect on mating performance. Alternatively, and in-dependent of its effects on male behavior, cue lure may be an “imperfect” mimic of anatural analogue and hence a less potent attractant than methyl eugenol, a naturalplant product.

As in

B. dorsalis

,

B. cucurbitae

males that fed on cue lure were unlikely to re-visita cue lure source for additional feeding. In the field experiment, an average of only 6previously exposed males were captured per replicate compared to 19 control malesper replicate. Similarly, among the 22 males that fed on cue lure during an initial 30-min trial (see above), not a single individual fed on cue lure during a second trial con-ducted 7 days later. These results are consistent with Chambers et al. (1972), who re-ported reduced trap captures of

B. cucurbitae

males for 3 weeks following exposure tocue lure. Given that cue lure confers only a short-term mating advantage, it remainsunclear why male melon flies show a reduced tendency to re-visit a cue lure source.

Regarding potential application to control efforts, the present study suggests thatpre-release exposure of sterile

B. cucurbitae

males to cue lure may both enhance theirmating competitiveness in the wild and decrease their attraction to lure-baited traps.Thus, pre-release exposure to lure may represent a relatively simple and inexpensivemeans to combine programs of sterile insect release and male annihilation (see alsoChambers et al. 1972). However, compared to the oriental fruit fly, the effectivenessof pre-release exposure may be limited for the melon fly given the short-lived matingadvantage conferred by the lure. For both species, assessing the value of this tech-nique will ultimately require field tests in which wild populations are compared be-tween areas receiving, for example, (1) sterile insect release with or without pre-release exposure to lure or (2) male annihilation alone or in combination with sterileinsect release with pre-release exposure to lure.


481

A

CKNOWLEDGMENTS

We thank S. Fong, K. Luke, and F. Meng for their help in rearing and marking theflies. This research was funded by grants from the USDA/ARS (58-91H2-6-42) andUSDA/CSRS (9401860).

R

EFERENCES

C

ITED

B

AKER

, R., R. H. H

ERBERT

,

AND

R. A. L

OMER

. 1982. Chemical components of the rectalgland secretions of male

Dacus cucurbitae

, the melon fly. Experientia 38: 232-233.

B

EROZA

, M., B. H. A

LEXANDER

, L. F. S

TEINER

, W. C. M

ITCHELL

,

AND

D. H. M

IYASHITA

.1960. New synthetic lure for the male melon fly. Science 131: 1044-1045.

C

HAMBERS

, D. L., K. O

HINATA

, M. F

UJIMOTO

,

AND

S. K

ASHIWAI

. 1972. Treating te-phritids with attractants to enhance their effectiveness in sterile-release pro-grams. J. Econ. Entomol. 65: 279-282.

C

HAMBERS

, D. L. 1977. Attractants for fruit fly survey and control, pp. 327-344

in

H.H. Shorey and J. J. McKelvey [eds.], Chemical control of insect behavior. Wiley,New York.

E

CONOMOPOULOS

, A. P.,

AND

G. E. H

ANIOTAKIS

. 1994. Advances in attractant andtrapping technologies for tephritids, pp. 57-66

in

C. O. Calkins, W. Klassen, andP. Liedo [eds.], Fruit flies and the sterile insect technique. CRC Press, Boca Ra-ton.

F

LETCHER

, B. S. 1987. The biology of dacine fruit flies. Ann. Rev. Entomol. 32: 115-144.

I

WAHASHI

, O.,

AND

T. M

AJIMA

. 1986. Lek formation and male-male competition in themelon fly,

Dacus cucurbitae

Coquillett (Diptera: Tephritidae). Appl. Entomol.Zool. 21: 70-75.

K

OBAYASHI

, R. M., K. O

HINATA

, D. L. C

HAMBERS

,

AND

M. S. F

UJIMOTO

. 1978. Sexpheromones of the oriental fruit fly and the melon fly: mating behavior, bioas-say method, and attraction of females by live males and by suspected phero-mone glands of males. Environ. Entomol. 7: 107-112.

K

UBA

, H.,

AND

J. K

OYAMA

. 1982. Mating behavior of the melon fly,

Dacus cucurbitaeCoquillett (Diptera: Tephritidae): comparative studies of one wild and two lab-oratory strains. Appl. Entomol. Zool. 17: 559-568.

KUBA, H., AND J. KOYAMA. 1985. Mating behavior of wild melon flies, Dacus cucurbi-tae Coquillett (Diptera: Tephritidae) in a field cage: courtship behavior. Appl.Entomol. Zool. 20: 365-372.

KUBA, H., J. KOYAMA, AND R. J. PROKOPY. 1984. Mating behavior of wild melon flies,Dacus cucurbitae Coquillett (Diptera: Tephritidae) in a field cage: distributionand behavior of flies. Appl. Entomol. Zool. 19: 367-373.

KUBA, H., AND Y. SOKEI. 1988. The production of pheromone clouds by spraying in themelon fly, Dacus cucurbitae Coquillett (Diptera: Tephritidae). J. Ethol. 6: 105-110.

METCALF, R. L., W. C. MITCHELL, T. R. FUKUTO, AND E. R. METCALF. 1975. Attractionof the oriental fruit fly, Dacus dorsalis, to methyl eugenol and related olfactorystimulants. Proc. Nat. Acad. Sci. 72: 2501-2505.

NISHIDA, R., O. IWAHASHI, AND K. H. TAN. 1993. Accumulation of Dendrobium super-bum (Orchidaceae) fragrance in the rectal glands by males of the melon fly, Da-cus cucurbitae. J. Chem. Ecol. 19: 713-722.

NISHIDA, R., K. H. TAN, S. TAKAHASHI, AND H. FUKAMI. 1990. Volatile components ofmale rectal glands of the melon fly, Dacus cucurbitae Coquillett (Diptera: Te-phritidae). Appl. Entomol. Zool. 25: 105-112.

PORAMARCOM, R., AND C. R. B. BOAKE. 1991. Behavioural influences on male matingsuccess in the Oriental fruit fly, Dacus dorsalis Hendel. Anim. Behav. 42: 453-460.


SCHULTZ, G. A., AND G. M. BOUSCH. 1971. Suspected sex pheromone glands in threeeconomically important species of Dacus. J. Econ. Entomol. 64: 347-349.

SHELLY, T. E. 1994. Consumption of methyl eugenol by male Bactrocera dorsalis(Diptera: Tephritidae): low incidence of repeat feeding. Florida Entomol. 77:201-208.

SHELLY, T. E., AND A. DEWIRE. 1994. Chemically mediated mating success in maleOriental fruit flies, Bactrocera dorsalis (Diptera: Tephritidae). Ann. Entomol.Soc. America 87: 375-382.

SIVINSKI, J. M., AND C. CALKINS. 1986. Pheromones and parapheromones in the con-trol of tephritids. Florida Entomol. 69: 157-168.

SUZUKI, Y., AND J. KOYAMA. 1980. Temporal aspects of mating behavior of the melonfly, Dacus cuccurbitae Coquillett (Diptera: Tephritidae): a comparison betweenlaboratory and wild strains. Appl. Entomol. Zool. 15: 215-224.

SUZUKI, Y., AND J. KOYAMA. 1981. Courtship behavior of the melon fly, Dacus cucurb-itae Coquillett (Diptera: Tephritidae). Appl. Entomol. Zool. 16: 164-166.

TANAKA, N., L. F. STEINER, K. OHINATA, AND R. OKAMOTO. 1969. Low- cost larval rear-ing medium for mass production of Oriental and Mediterranean fruit flies. J.Econ. Entomol. 62: 967-968.

VARGAS, R. I., D. MIYASHITA, AND T. NISHIDA. 1984. Life history and demographic pa-rameters of three laboratory-reared tephritids (Diptera: Tephritidae). Ann. En-tomol. Soc. America 77: 651-656.

WHITTIER, T. S., AND T. E. SHELLY. 1995. Trimedlure affects mating success and mateattraction in male Mediterranean fruit flies. Entomol. Exp. Appl. In review.

WONG, T. T. Y., D. O. MCINNIS, AND J. I. NISHIMOTO. 1986. Melon fly (Diptera: Te-phritidae): sexual maturation rates and mating responses of laboratory-reared

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and wild flies. Ann. Entomol. Soc. America 79: 605-609.

482



CHEMICAL FACTORS INVOLVED IN SELECTION OF HOST PLANT FOR OVIPOSITION BY THE PICKLEWORM MOTH

(LEPIDOPTERA: PYRALIDAE)

J

OSEPH

K. P

ETERSON

AND

K

ENT

D. E

LSEY

USDA, Agricultural Research ServiceVegetable Laboratory

2875 Savannah HighwayCharleston, SC 29414-5334

A

BSTRACT

Studies with caged gravid females of the pickleworm, [

Diaphania nitidalis

(Stoll.)], revealed that leaves of yellow squash (

Curcurbita pepo

L

.

) contain small (MW<1000), non-volatile and highly polar amphoteric compounds which stimulate ovipo-sition on artificial sites. Several compounds, extracted from paper chromatograms,caused moderate stimulation of oviposition. When these extracts were recombined,the mixture proved highly active. Addition of whole leaf volatiles to the active water-soluble fraction increased oviposition. The whole leaf volatiles mixture could be sub-stituted with volatiles originating from leaf glandular trichomes.

Index Words. Lepidoptera,

Diaphania nitidalis

, oviposition, host plant, squash,

Cur-curbita pepo

, trichomes, volatiles.

Peterson & Elsey: Chemical Factors for Pickleworm Oviposition

483

R

ESUMEN

Estudios sobre el gusano de los pepinos [

Diaphania nitidalis

(Stoll.)] revelaron quelas hojas de la calabaza (

Cucurbita pepo

L.) contienen pequeños (MW<1000) compues-tos anfotéricos no volátiles y altamente polares que estimulan la ovoposición en sitiosartificiales. Varios compuestos, extraidos a partir de cromatogramas en papel, estimu-laron moderadamente la ovoposición. Cuando esos extractos fueron recombinados, lamezcla fue muy activa. La adición de volátiles de la hoja completa a la fracción activahidrosoluble incrementó la ovoposición. La mexcla de volátiles de la hoja completa po-

dría ser sustituida por volátiles originados de los tricomas glandulares de las hojas.

The pickleworm, [

Diaphania nitidalis

(Stoll.)], is a severe pest of cucurbit vegeta-bles in the southeastern United States, mainly because the larvae tunnel into thefruit. Commercially, damage tolerance is low, especially in the pickling industry (El-sey et al. 1989). Infestations in the field vary with cucurbit species as well as with cul-tivar (Corley 1973, Day et al. 1961, Pulliam 1979, Quisumbing & Lower 1975).Bioassays with cucumber cultivars showed that little antibiosis to larvae exists (Weh-ner et al. 1985). However, non-preference to oviposition was shown in glabrous mu-tants of muskmelon and cucumber (Day et al. 1961, Elsey & Wann 1982). Since nearlyisogenic lines of glabrous and pubescent cucumbers were evaluated, it appeared thatresistance (avoidance) was due to the lack of plant trichomes (Elsey & Wann, 1982).Elsey (1985) showed that resistance in

Cucurbita moschata

was due to ovipositionnon-preference. A study with whole plants in large field screen cages showed thatpickleworm oviposition preference decreased in the order of yellow squash, cucumberand watermelon (Elsey 1981). A similar pattern was observed when ethanol extractsof whole leaves were applied to artificial oviposition sites (Elsey & McFadden 1981).

Because chemical factors were suspected to be involved in oviposition preferenceby pickleworms, this study was undertaken to isolate these stimulants. Leaves ofstraightneck yellow squash were used because this yellow squash is a preferred host(Dilbeck & Canerday 1968) and most pickleworm eggs are laid on foliage. Effects ofvolatile and non-volatile compounds were evaluated and attempts to isolate non-vol-atile stimulants are described.

M

ATERIALS

AND

M

ETHODS

Insect Rearing

Diaphania nitidalis

were raised in the laboratory according to methods describedby Elsey et al. (1984).

Culture of Plant Material

Seedlings of yellow squash (

Cucurbita

pepo

L. var. Early Prolific Straightneck)were planted in the field and raised using standard cultural practices. Healthy leaves,approximately one half ultimate size (10 - 12 cm longest dimension) were collectedfrom the field early in the morning and immediately processed or used for bioassays.

Bioassays

Assays involving oviposition were conducted according to Elsey & McFadden(1981) with some modification to improve quantitative aspects. Pads of building insu-

484



lation glasswool (diam 9 cm, thickness 2-3 cm) were placed in petri dishes containingthe solutions to be tested. After adsorption was completed, the solvent was evapo-rated. Subsequently, the pads were suspended from the top of wire mesh cages (120

×

90

×

90 cm), which contained about 100 pairs of 3-day-old moths. The cages were lo-cated in a humidified room held at 23 C, and a photoperiod of 16:8 (L:D). Testing ma-terial was placed in the cages in the late afternoon (before the start of the scotoperiod), removed the next morning and the eggs counted.

Common amino acids were tested for stimulation of oviposition by dissolving themin 50% ethanol and applying 50

µ

g or 200

µ

g per pad. The following amino acids wereused: L-alanine,

γ

-amino butyric acid, L-arginine, L-asparagine, aspartic acid, citrul-line, cysteine, L-glutamic acid, L-glutamine, glycine, histidine, DL-homocysteine, hy-droxyproline, isoleucine, leucine, lysine, DL-methionine, phenylalanine, proline,serine, DL-threonine, tryptophan, L-tyrosine and DL-valine. A test consisted of fourpads containing one (different) amino acid each, one pad with 0.25 g equivalent of anaqueous polyamide fraction (control, contains no volatiles), and one pad without ap-plication (blank).

Volatiles from small whole leaves (6-9 cm longest dimension) or droplets (trichomeexudates) were tested by placing them between glass fiber pads. Alternatively, thefresh material was lyophilized for 72 hrs under high vacuum while the volatiles weretrapped under liquid nitrogen. The leaves, now free of volatiles, were extracted andtested.

Extractions and Separations

Leaves approximately two-thirds full size [4-7 g fresh weight (FW), 14-18 cm long-est dimension of leaf blade] were harvested from greenhouse grown squash plants.The leaves were lyophilized and ground to pass a 0.85 mm mesh screen. The materialwas subsequently extracted with hexane for approximately 12 hours on a shaker andfiltered. The procedure was repeated twice and the residue was extracted with hot95% ethanol in a homogenizer and filtered. After two repetitions, the combined etha-nol extracts were condensed to a syrup, and water was added. The extract was filteredthrough a series of surface membrane molecular filters under nitrogen pressure in astirring cell. The filters had a nominal molecular weight cut-off (MWCO) of 20K, 5K,

T

ABLE

1. O

VIPOSITION

B

IOASSAYS

OF

F

ILTRATES

O

BTAINED

BY

S

EQUENTIAL

F

ILTRA-TION

T

HROUGH

M

EMBRANE

M

OLECULAR

F

ILTERS

.

Filter Average Oviposition

1,2

(MWCO) (Percent of Total)

none 11.5 b20 K 15.8 bc5 K 22.1 de1 K 26.9 e500 19.9 cd

blank

3

3.9 a

1

Average of four independent bioassays.

2

Means followed by the same letter are not significantly different. Means separation by Fisher’s LSD (P =0.05).

3

No fraction applied.


485

1K and 500 respectively. Since the 500 MWCO gave rise to losses of activity (Table 1),filtrations of subsequent extracts were limited to a MWCO of 1K. Molecular filtrationwas followed by partitioning against dichloromethane and ethylacetate respectively.The extraction scheme is presented in Fig. 1.

Chromatography

The aqueous fraction obtained after the last partitioning was subjected to low-pressure column chromatography using polyamide as stationary phase (column di-mensions: 25

×

2.6 cm diam). The polyamide was prewashed with one liter each of0.5% formic acid and water, respectively. An extract aliquot representing up to 100 gleaf fresh weight equivalent (FWE) in 10 ml water was pumped onto the column andeluted stepwise with methanol/acetic acid (995:5), 100% methanol and water respec-tively. Absorbance was monitored at 280 nm. The fractions were tested for stimulationof oviposition and checked with p-anisidine phthalate to detect reducing sugars (Stahl1969, p. 857) and with ninhydrin (Stahl 1969, p. 889) or vanillin-potassium hydroxide(Stahl 1969, p. 904) to test for primary or secondary amino functions.

Figure 1. Extraction and partitioning scheme for the isolation of oviposition stim-ulants. All fractions were assayed simultaneously, and the assay was performed threetimes. Numbers in parentheses indicate the amount of oviposition as a percent of thetotal number of eggs deposited. Numbers were corrected for ‘background’, i.e. eggs de-posited on pads that received no extracts.

486



The active fraction from the polyamide column (water fraction) was condensed andused for paper chromatography on Whatman 3 MM sheets (46

×

57 cm). Developmentwas in the descending mode using tert-butanol: acetic acid:water-3:1:1. After drying,the sheets were sprayed on the sides with ninhydrin and heated at 110

°

C. The areacorresponding with the colored bands, as well as the areas in between, were extractedwith 50% ethanol and bioassayed.

The active fraction from the polyamide column was also run through a Dowex 50

×

8 cation exchange column (18 cm

×

1 cm diam; sample 10 g FWE). The sample wasadjusted to pH 2.5 with concentrated formic acid and eluted with 8N formic acid, wa-ter, and triethylamine (in 20% aqueous acetone, pH=10). After the column had beenstanding for three days, more activity could be eluted with triethylamine [Table 3, Et

3

N(2)]. The fractions were checked with ninhydrin and tested for activity.Hydrolysis was performed on the active polyamide fraction. The reaction was con-

ducted in a sealed vial at 110

°

C for 24 hrs, in 2N HC1. The products were condensedto dryness, redissolved and bioassayed. Seven such tests were performed.

Sources of Chemicals

L-glutamine, L-asparagine, L-glutamic acid, L-arginine, proline, L-tyrosine andtryptophane were obtained from Aldrich Chemical Co., Milwaukee, WI. L-alanine, as-partic acid,

γ

-amino butyric acid, citrulline, cysteine, glycine, histidine, DL-homocys-teine, isoleucine, leucine, lysine, DL-methionine, phenylalanine, hydroxy-L-proline,

Figure 2. Dose-response curve of oviposition (% of total number eggs produced perbioassay) versus the logarithm of the amount of extract used. All concentrations wererepresented once in a bioassay and the entire assay was performed four times. Re-gression analysis: y=1.18 + 11.86x, r2=0.91, P<0.01.


487

serine, DL-threonine, DL-valine and ninhydrin were obtained from Sigma ChemicalCo., St. Louis, MO. p-Anisidine phthalate was obtained from Pfaltz and Bauer, Inc.Waterbury, CT. Polyamide was obtained from Universal Scientific Inc. Atlanta, GA.Hexane, dichloromethane and ethylacetate (HPLC grade) were obtained from Bur-dick and Jackson, Muskegon, MI. Tertiary butanol was obtained from J. T. Baker Inc.,Philipsburg, NJ.

Statistical Analyses

Statistical analyses, consisting of one-way ANOVA, students t-test, and regressiontechniques, were performed where appropriate on a personal computer with theSOLO statistical system (BMDP Statistical Software, Inc., Los Angeles, CA). Meansseparations were by Fisher’s LSD (P=0.05).

R

ESULTS

AND

D

ISCUSSION

Preliminary experiments showed that increasing amounts of crude 95% ethanolextracts of fresh squash leaves applied to glasswool pads increased egg-laying. At ap-proximately 1 g FWE, i.e., the amount obtained from 1 g fresh leaf material, no fur-ther increase was obtained, and at higher dosages very erratic dose-response datawere obtained. A linear dose-response (r

2

= 0.91) was obtained when the percent of thetotal number of eggs laid was plotted against the logarithm of the extract concentra-tion (Fig. 2).

The partial isolation scheme (Fig. 1) shows that the oviposition stimulants re-mained in the aqueous fractions. Oviposition increased considerably when fractions oflow polarity were removed (Fig. 1) or when molecular filtration was performed (Table1). Although this may suggest the presence of inhibitors, the fractions with low polar-ity showed activity above ‘background’, i.e., when applied to glasswool pads, they re-ceived more eggs than non-treated pads. In addition to the molecular filtration downto a MWCO=1000, a filter with a MWCO=500 was used; in this case a significantamount of activity was lost. When the pH of the final aqueous fraction (Fig. 1) was ad-justed to either 10 or 2.5, no activity could be extracted by ethyl acetate, suggestingthat oviposition stimulants were highly polar molecules.

The final active aqueous fraction obtained through partitioning (Fig. 1) was sub-jected to low pressure column chromatography, using polyamide as the stationaryphase, and the fractions were tested for activity (Fig. 3). The first fraction (aqueous)contained virtually all of the activity. The first eluate from the polyamide column maycontain salts, organic acids, sugars (Winter & Hermann 1986) and some water soluble

T

ABLE

2. E

FFECT

OF

H

YDROLYSIS

OF

THE

A

QUEOUS

P

OLYAMIDE

(A

CTIVE

) F

RACTION

1

ON

O

VIPOSITION

BY

THE

P

ICKLEWORM

M

OTH

.

Treatment Oviposition (% of Total) Mean SEM

Non-hydrolyzed 50.3 25.1 37.2 35.6 49.0 49.2 51.3 42.5 a

2

3.8Hydrolyzed 35.4 61.8 46.5 46.8 37.1 34.8 40.2 43.2 a 3.6Blank (no application) 14.3 13.2 16.4 17.6 13.9 16.0 8.5 14.3 b 1.1

Total oviposition 796 1595 807 1585 2132 762 995

1

0.25 g FWE applied to oviposition sites.

2

Means followed by the same letter are not significantly different. Means separation by Fisher’s LSD (P = 0.05).

488



phenolic glycosides (Thieme 1964). Activity was also found in the very first part of theaqueous eluate, before any absorption at 280 nm was observed, excluding phenoliccompounds. Para-anisidine phthalate gave no reaction, indicating that no reducingsugars were present (Stahl 1969). Vanillin-potassium hydroxide or ninhydrin gavepositive reactions, indicating the presence of primary or secondary amino groups

T

ABLE

3. C

ATION

E

XCHANGE

C

HROMATOGRAPHY

OF

A

CTIVE

A

QUEOUS

F

RACTION

F

ROM

P

OLYAMIDE

C

HROMATOGRAPHY

.

Eluent Vol. (ml)NinhydrinReaction

Mean Oviposition(% of Total)

1

SEM

HCOOH (8N) 106 negative 28.52 b

3

2.30H

2

O 86 trace 13.54 a 1.68Et

3

N (1)

2

84 posit. (purple) 25.09 b 2.42Et

3N (2) 92 posit. (purple) 26.21 b 1.78Blank 6.63 a 0.43

1Six experiments were performed (n=6). A total of 700, 1345, 1320, 1198, 1610, and 1819 eggs were layed perexperiment.

2Et3N = triethylamine.3Means followed by the same letter are not significantly different (Fisher’s LSD, P = 0.05).

Figure 3. Polyamide column chromatography of active aqueous extract (see Fig. 1),monitored at 280 nm. Activity is expressed as percent of the activity in the pre-columnmatrix (active solution applied to column; activity = 100%). Numbers were correctedfor the blank (no application). Four independent assays were performed and datawere averaged.

Peterson & Elsey: Chemical Factors for Pickleworm Oviposition 489

(Stahl 1969). Hydrolysis of the active aqueous fraction from the polyamide column didnot change the activity significantly (Table 2). These results indicate that no activesmall peptides were present, or, if present, did not play a significant role.

The active fraction from the polyamide column was further separated on a cationexchange column. Activity was present in all fractions except the aqueous (Table 3),indicating acidic and basic compounds were involved in stimulation of oviposition.

Numerous paper chromatograms (PC) were developed from the active polyamidefraction. The developed papers were sprayed on the side with ninhydrin, and the en-tire chromatograms stained (hues of blue, purple and pink), except the front area.Twelve bands were cut, eluted and tested. Also tested were all the extracted regionsrecombined. An example of the PC and bioassay data, presented in Table 4, show thatvarious regions were active, and that the recombined sample was very active. SincePC and parallel bioassays showed that active regions, with the exception of one, couldbe stained with ninhydrin, primary or secondary amino functions, were involved. Fur-thermore, ion-exchange data showed that acidic and basic compounds were involved.These compounds could not be extracted from an aqueous solution at high or low pH,indicating amphoteric characteristics of the active compounds. These propertiesstrongly indicate amino acids. For this purpose common amino acids were tested. The

TABLE 4. PAPER CHROMATOGRAPHY1 OF ACTIVE FRACTION (AQUEOUS) OF POLYAMIDECOLUMN CHROMATOGRAPHY.

Oviposition (% Control)3

Band hRf2 Mean SEM

1 0-11.1 10.7 de4 3.92 11.1-19.5 10.1 de 3.73 19.5-26.5 6.4 e 4.24 26.5-34.7 48.6 a 32.95 34.7-40.5 17.6 cde 2.36 40.5-47.9 33.9 b 11.07 47.9-50.8 25.3 bc 4.48 50.8-57.7 22.0 bcd 6.49 57.7-63.8 11.6 de 3.110 63.8-68.3 24.6 bc 12.511 68.3-76.2 14.9 cde 1.612 76.2-100 52.8 a 18.3Recombined5 255 65.4Blank6 15 2.6

1Solvent system: tert-BuOH:HOAc:H2O-3:1:1. 4.0 g FWE of the aqueous fraction of the polyamide chromatog-raphy applied to paper sheets. Control: 0.25 g FWE of the polyamide fraction.

2All bands stained with ninhydrin, except number 12. hRf: relative traveling distance of compound, 0-com-pound did not move, 100-compound moved with solvent front.

3Data obtained from 4 chromatograms. Oviposition for the 12 bands and the recombined sample was ex-pressed as follows:

sample - blank (no applic.) × 100%control - blank

4Means followed by the same letter are not significantly different. Means separation by Fisher’s LSD (P =0.05).

50.5 g FWE of each chromatographic band was recombined.6No application of sample.


highest response was 8.3% of control (Table 5), all other responses were considerablylower, sometimes slightly negative. These responses were far below those obtained byextracts of the PC bands (Table 4), where six bands caused increases in excess of 20%of the control and the highest response was 52.8%. It therefore seems likely that someof the non-protein amino acids, which are ubiquitous in the cucurbit family (Noe &Fowden 1960, Fang et al. 1961, Dunnill & Fowden 1965) are responsible for stimula-tion of oviposition. Since these particular non-protein amino acids seem to be limitedto the cucurbits, a particular composition with respect to identity and relative concen-trations could explain pickleworm host plant specificity.

TABLE 5. INFLUENCE OF COMMON AUTHENTIC AMINO ACIDS1 ON OVIPOSITION BY THEPICKLEWORM MOTH.

Amino Acid Oviposition2

50 µg 200 µg

L-alanine 4.5 -2.5γ-amino butyric acid 1.7 1.9L-arginine -0.3 -3.2L-asparagine 4.8 2.3Aspartic acid 1.1 4.4Citrulline 4.6 1.8Cysteine 3.7 4.0L-glutamic acid 3.3 2.4L-glutamine 2.0 1.3Glycine 4.5 3.4Histidine 2.4 4.1DL-homocysteine 5.8 2.3Hydroxy-L-proline -0.4 1.7Isoleucine 0.3 -1.4Leucine 0.4 5.2Lysine 5.2 3.5DL-methionine 1.7 2.1Phenylalanine 0.4 4.9Proline 0.8 3.4Serine 3.9 4.6Threonine -0.8 3.9Tryptophane 8.3 3.0L-tyrosine 2.6 1.0DL-valine 3.5 -1.6

1Amino acids were tested in random order, three per test, at amounts of 50 µg and 200 µg per oviposition site.2Oviposition was expressed as follows:

sample - blank (no application) × 100%control - blank

where the extract used was 0.25 g FW squash leaf equivalent, obtained by extraction with 95% ethanol. Totaleggs layed per experiment ranged from 1341 to 3778.

Peterson & Elsey: Chemical Factors for Pickleworm Oviposition 491

Volatiles originating from whole leaves increased oviposition significantly (Table6). A similar increase was obtained when ‘whole leaf ’ volatiles were substituted byvolatiles originating from droplets produced by leaf trichome glands (Table 7). Subse-quent research (Peterson, et al. 1994) showed that these volatiles primarily cause in-creased attraction rather than stimulation of oviposition. Small, highly polar andnon-volatile compounds serve as oviposition stimulants. Most, if not all of these com-pounds carry a charge, are amphoteric, and most react positively for primary or sec-ondary amino functions. The active compounds were stable and hydrolysis did notdecrease activity.

ENDNOTE

Mention of a trademark or proprietary product does not constitute a guarantee orwarranty of the product by the U.S. Dep. Agric. and does not imply its approval to theexclusion of other products that also may be suitable.

REFERENCES CITED

CORLEY, W. L. 1973. Response of muskmelon botanical varieties to pickleworm infes-tation. HortScience 13:286-287.

TABLE 6. INFLUENCE OF ‘WHOLE LEAF’ VOLATILES1 ON OVIPOSITION ON ARTIFICIALSITES TREATED WITH SQUASH LEAF EXTRACTS.2

Treatment Oviposition (% Total) Mean SEM

Extr. without volatiles 38.4 37.9 45.4 42.8 34.4 39.8 a3 1.9Extr. with volatiles 59.3 54.7 53.6 46.6 49.8 52.8 b 2.2Blank (no application) 2.4 7.4 1.0 10.6 15.8 7.4 c 2.7

Total oviposition 4687 686 3415 1985 2210

1A small whole leaf sandwiched between glass fiber pads; bottom of leaf facing treated side.2Extracts free of volatiles; 0.25 FWE of aqueous polyamide fraction.3Means followed by the same letter are not significantly different. Means separation by Fisher’s LSD (P =

0.05).

TABLE 7. INFLUENCE OF LEAF GLANDULAR TRICHOME EXUDATES1 ON OVIPOSITION ONARTIFICIAL SITES TREATED WITH SQUASH LEAF EXTRACTS.2

Treatment Oviposition (% Total) Mean SEM

Ext. without volatiles 40.8 40.9 45.7 42.9 43.1 42.7 a3 0.89Ext. without volatiles,With trichome droplets

56.4 56.0 52.3 54.8 54.3 54.8 b 0.72

Blank (no application) 2.8 3.1 2.0 2.3 2.7 2.6 c 0.20

Total oviposition 5231 6840 9283 7230 4663

1Approximately 200 droplets (trichome exudates) applied between two layers of glass fiber pads.2Extracts free of volatiles; 0.25 g FWE of aqueous polyamide fraction.3Means followed by the same letter are not significantly different. Means separation by Fisher’s LSD (P =

0.05).


DAY, A., P. E. NUGENT, AND J. F. ROBINSON. 1961. Resistance of squash varieties tothe pickleworm and the value of resistance to insecticide control. J. Econ. En-tomol. 54: 1191-1197.

DILBECK, J. D., AND T. D. CANERDAY. 1968. Resistance of Cucurbita to the pickleworm.J. Econ. Entomol. 61:1410-1413.

DUNNILL, P. M., AND L. FOWDEN. 1965. The amino acids of seeds of the cucurbitaceae.Phytochemistry. 4:933-944.

ELSEY, K. D. 1981. Pickleworm: Survival, development, and oviposition on selectedhosts. Ann. Entomol. Soc. America 74:96-99.

ELSEY, K. D., AND T. L. MCFADDEN. 1981. Pickleworm: oviposition on an artificial sur-face treated with cucurbit extract. J. Econ. Entomol. 74:473-474.

ELSEY, K. D., AND E. V. WANN. 1982. Differences in infestation of pubescent and gla-brous forms of cucumber by pickleworms and melonworms. HortScience17:253-254.

ELSEY, K. D., T. L. MCFADDEN, AND R. B. CUTHBERT. 1984. Improved rearing systemfor pickleworm and melonworm (Lepidoptera: Pyralidae). J. Econ. Entomol.77:1070-1072.

ELSEY, K. D. 1985. Resistance mechanisms in Cucurbita moschata to pickleworm andmelonworm (Lepidoptera: Pyralidae). J. Econ. Entomol. 78:1048-1051.

ELSEY, K. D., J. A. KLUN, AND M. SCHWARTZ. 1989. Pickleworm sex pheromone: po-tential for use in cucumber pest management. J. Agric. Entomol. 6:275-282.

FANG, S-D., L-C LI, C-I NIU, AND K. F. TSENG. 1961. Chemical studies on Cucurbitamoschata Duch. I. The isolation and structural studies of cucurbitine, a newamino acid. Scientia Sinica Vol. X, No. 7, 845-851.

NOE, F. F., AND L. FOWDEN. 1960. β-Pyrazol-l-ylalanine, an amino acid from water-melon seeds (Citrullus vulgaris). Biochem. J. 77:543-546.

PETERSON, J. K., R. J. HORVAT, AND K. D. ELSEY. 1994. Squash leaf glandular tri-chome volatiles: Identification and influence on behavior of female picklewormmoth [Diaphania nitidalis (Stoll.)] (Lepidoptera: Pyralidae). J. Chem. Ecol.20:2099-2109.

PULLIAM, T. L. 1979. The effect of plant types and morphology of the cucumber, Cucu-mis sativus L., on infestation by the pickleworm, Diaphania nitidalis (Stoll.).M.S. Thesis, North Carolina State University.

QUISUMBING, A. R., AND R. L. LOWER. 1975. Screening cucumber cultivars for pickle-worm resistance. HortScience 10:147 (Abstr.).

STAHL, E. 1969. Thin-layer chromatography. Second Edition. Springer-Verlag, NewYork, Heidelberg, Berlin.

THIEME, H. 1964. Die Phenolglykoside der Salicaceen. 2. Mitteilung:Isolierung undNachweis. Pharmazie, 19:471-475.

WEHNER, T. C., K. D. ELSEY, AND G. G. KENNEDY. 1985. Screening for antibiosis topickleworm. HortScience 20:1117-1119.

WINTER, M., AND K. HERMANN. 1986. Bestimmung von Hydroxyzimtsäure- Derivatenin Obst und Gemüse durch HPLC nach Polyamid- Fraktionierung. Lebensmit-telchem. Gerichtl. Chem. 40:6-22.

Orozco-Santos et al.: Floating Row Cover & Transparent Mulch

493

FLOATING ROW COVER AND TRANSPARENT MULCH TO REDUCE INSECT POPULATIONS, VIRUS DISEASES AND

INCREASE YIELD IN CANTALOUPE

M

ARIO

O

ROZCO

-S

ANTOS

, O

CTAVIO

P

EREZ

-Z

AMORA

A

ND

O

SCAR

L

OPEZ

-A

RRIAGA

Instituto Nacional de Investigaciones Forestales y AgropecuariasCentro de Investigaciones del Pacifico Centro, Campo Experimental Tecoman,

Apartado postal 88Tecoman, Colima 28100, México

A

BSTRACT

The influence of floating row cover on cantaloupe, alone or combined with trans-parent polyethylene mulch, was evaluated to determine its effect on population den-sities of aphids, sweetpotato whitefly,

Bemisia

tabaci

(Gennadius), and the vegetableleafminer,

Lyriomyza

sativae

Blanchard, as well as virus incidence and yield of can-taloupe,

Cucumis

melo

L. Cv. Durango. The study was carried out in a dry tropic re-gion of Colima, Mexico. Aphids, sweetpotato whitefly and vegetable leafminer werecompletely excluded by floating row cover while the plots were covered. Transparentpolyethylene mulch reduced aphids and whitefly populations, but did not affect infes-tation by

L

.

sativae

. Floating row cover delayed the appearance of virus-diseasedplants for two weeks with respect to control (bare soil). Furthermore, the transparentmulch reduced virus incidence. The yield (weight) and number of cantaloupe melonsharvested were nearly four-fold higher in mulch plus row cover plots (when the coverwas removed during perfect flowering) compared with the control plots, while theyield with floating row cover alone was tripled. The yield from mulched plants aloneand covered plants (the cover removed during vegetative growth) was higher thanthat from plants grown on bare soil.

Key Words:

Cucumis melo

, aphids, sweetpotato whitefly, vegetable leafminer.

R

ESUMEN

Se evaluó la influencia de las cubiertas flotantes, solas o combinadas con una co-bertura de plástico transparente, sobre la densidad de población de áfidos y de moscablanca de la batata,

Bemisia tabaci

(Gennadius), y sobre la infestación del minador dela hoja

Lyriomyza

sativae

Blanchard, así como sobre la incidencia de virus y la pro-ducción dé melón “Cantaloupe”,

Cucumis

melo

L. Cv. Durango. La investigación serealizó en la región tropical seca del estado de Colima, México. Los áfidos, la moscablanca de la batata y el minador de la hoja fueron completamente excluídos por las cu-biertas flotantes, mientras las plantas estuvieron protegidas con la tela de polipropi-leno. El acolchado con plástico transparente redujo la población de áfidos y moscablanca, pero no tuvo efectos sobre la infestación por

L

.

sativae

. La cubierta flotante re-trasó durante dos semanas la presencia de plantas afectadas por virus con respecto altestigo (suelo desnudo). El plástico transparente también redujo la incidencia de vi-rus. El número de frutos y el rendimiento del melón en las parcelas con cubierta deplástico + cubierta flotante (la cubierta fue retirada durante la floración perfecta) fuecasi cuatro veces más alto en comparación a las parcelas con suelo desnudo, mientrasque en aquellas con cubierta flotante sola, el rendimiento se triplicó. La producción delas plantas con cubierta plástica sola o de las plantas con cubierta flotante (retiradadurante el crecimiento vegetativo) fue más alta que la de las plantas cutivadas sobre

suelo desnudo.

494



Cantaloupe melon,

Cucumis

melo

L., is the major cucurbit crop in the dry tropicareas of Colima State, Mexico, during fall and winter, with more than 1500 ha grownannually. Recently in this area, aphid-borne virus diseases have been a problem forcucurbit crops. In Mexico, a cucumovirus (cucumber mosaic) and potyviruses (water-melon mosaic-2, papaya ringspot-W and zucchini yellow mosaic) are the major virusdiseases affecting squash, watermelon, cucumber and cantaloupe (Delgadillo et al.1989). These viruses are transmitted efficiently by aphids in a nonpersistent manner(Nameth et al. 1986).

The sweetpotato whitefly,

Bemisia

tabaci

Gennadius, is an important pest of agri-cultural and horticultural crops throughout the world, causing damage directly byfeeding and indirectly through the excretion of honeydew (Byrne et al. 1990) and thetransmission of virus diseases (Cohen 1990). In the past decade, this insect has beenreported as a serious pest of various crops in Florida (Schuster & Price 1987), Arizona(Butler 1982) and California (Perring et al. 1991). Recently, it has been found in ex-tremely high populations in the agricultural areas of Arizona, California, Texas andnorthwestern Mexico (Gill 1992). Also, in the last two years, sweetpotato whitefly hasbecome the most important pest in melon-growing areas of Colima, Mexico (Farias etal. 1994). Other pests affecting melon crops are the vegetable leafminer,

Lyriomyzasativae

Blanchard, and some species of

Diabrotica

.Most of the management strategies against cucurbit virus diseases have been tar-

geted at the aphid-borne viruses. Several measures are used to influence the behaviorof the insect vectors in an attempt to interfere with virus transmission, including theuse of transparent polyethylene mulch (Brown et al. 1993, Lecoq & Pitrat 1983,Orozco et al.1994), oil sprays (Simons & Zitter 1980) and floating row covers (Natwick& Durazo 1985, Natwick & Laemmlen 1993, Orozco et al. 1994, Perring et al. 1989,Webb & Linda 1992). However, in dry tropic conditions, the use of oil sprays can causephytotoxic symptoms on the foliage and reduce the fruit yield of cantaloupe (Orozcoet al. 1994). The use of plastic mulches in agriculture continues to increase in hightechnology systems for horticultural crop production, and more recently, floating rowcovers of spunbonded polyester or polyethylene have been introduced to protect cucur-bits from insect-vectored viruses and their aphid and whitefly vectors (Natwick &Laemmlen 1993, Perring et al. 1989, Webb & Linda 1992) and from direct damagefrom pests, such as the vegetable leafminer (Natwick & Laemmlen 1993), melon-worm,

Diaphania hyalinata

(L.), pickleworm,

D

.

nitidalis

(Stoll) (Webb & Linda 1992)and cucumber beetles (Adams et al. 1990). Plastic mulch in combination with floatingrow covers show promise as a method for excluding insects, reducing virus diseasesand increasing yield of cucurbit crops (Orozco et al. 1994, Ramirez & Lopez 1991,Webb & Linda 1992). However, there is little information on the development of thecantaloupe crop grown under floating row cover in a semi-arid tropical region.

This study was carried out to evaluate the effect of transparent polyethylene mulchand floating row covers, with or without transparent mulch, on insect populations, viraldiseases and yield of melons grown in the field under semi-arid tropical conditions.

M

ATERIALS

AND

M

ETHODS

The field experiment was conducted at the Tecoman Experimental Station inColima, Mexico during the winter-spring of 1992-1993. On February 19, 1993, melon(

Cucumis

melo

L.) Cv. Durango was directly seeded at 23 cm intervals in the center of1.78 m wide beds, and plots were watered with a drip irrigation system to promotegermination. Each bed had an emitter tube with emission spacing of 0.5 m, and flowrate of 2.3 liter per h. Five treatments were evaluated: (1) transparent polyethylenemulch (TPM), (2) floating row cover (FRC, spunbonded polyethylene fiber: AGRIBON-


495

17, Bonlam S.A. de C.V. San Luis Potosi, Mexico) removed at the vegetative stage (20March, before staminated flowers appeared in the plants) or (3) the reproductivestage (28 March, when 50% of the plants had their first perfect flower); and (4) TPM+ FRC removed at the reproductive stage. Bare soil plots (5) were included as thecheck (control).

After seeding, FRC was set on the beds with both edges and ends of covers securedwith soil. Weeds became a problem under the covers, mainly in the floating row coverwith bare soil, and it was necessary to control grass with the herbicide Fluazifop-butil(Fusilade

TM

); broad-leaf weeds were removed manually. In TPM and bare soil plots,eleven insecticide sprays were applied during all the crop cycle, mainly for sweetpo-tato whitefly control, while in FRC plots, alone or combined with TPM removed at re-productive stage, only five sprays were necessary after the covers were removed. Theinsecticides used were endosulfan, methamidophos, diazinon, bifenthrin, delta-methrin, cyfluthrin, lambdacyhalothrin, and permethrin. Each treatment consisted offive 6.9 m-long beds replicated four times in a randomized complete block design.

Yellow pan plastic traps for aphids (28

×

22

×

12 cm) containing a mixture of waterand detergent were placed in the middle of one row in each plot, and the number ofaphids trapped were determined twice weekly to determine the effectiveness of eachtreatment in delaying aphid infestation. Aphids were stored in 70% alcohol and wereidentified to species using several aphid taxonomy keys (Basky 1993, Peña 1991, Pikeet al. 1990). Counts of sweetpotato whitefly adults were made on 16 plants (two leavesper plant) in early morning (7-8 h

A

.

M

.), before flight activity occurred. Leaves wereturned over gently and adults visually counted (Butler & Henneberry 1991). On 24March and 1 April (34 and 41 days after planting, respectively), the numbers of sweet-potato whitefly nymphs on five leaflets from two lower leaves were counted from eightplants with the aid of a 10x dissecting microscope. Also, on 19, 26 March, 2 and 12April, vegetable leafminer damage was assessed on the main guide shoot on 16 plantsper plot. A leaf was considered damaged if more than three miners per leaf werefound.

To evaluate virus incidence, all of the plants in each plot showing disease symp-toms were counted weekly. At harvest, plant tissue was analyzed by ELISA test(Clark & Adams 1977) to detect infection by cucumber mosaic virus (CMV), water-melon mosaic virus-2 (WMV-2), papaya ringspot virus-W (PRSV-W), zucchini yellowmosaic virus (ZYMV) and squash mosaic virus (SqMV).

Yield was assessed by picking all the fruits in two central beds of each experimen-tal plot and sorting them into marketable and culled categories. The harvest was doneevery day during two weeks, and marketable fruit (based on export regulations) wassized 9, 12, 15, 18 and 23 fruits per carton.

R

ESULTS

AND

D

ISCUSSION

Fourteen aphid species were caught in yellow water pan traps throughout the ex-perimental period. These species were (% of total):

Aphis

spiraecola

Patch (23%),

Uro-leucon

ambrosiae

(Thomas) (22%),

Myzus

persicae

(Sulzer) (13%),

A.

gossypii

(Glover)(12%),

Rhopalosiphum rufiabdominalis

(Sasaki) (11%),

R.

maidis

(Fitch) (4%),

A.craccivora

(Koch) (3%),

Schizaphis graminum

(Rondani) (3%),

A.

nerii

(Boyer deFonsc) (2%),

Macrosiphum

euphorbiae

(Thomas) (2%),

Lipaphis

erysimi

Kaltenbach(2%),

R

.

padi

(L.) (1%),

Toxoptera

aurantii

Boyer de Fonsc. (1%) and

A

.

fabae

Scopoli(1%).

No alate aphids were caught in yellow traps in the plots with FRC until the coverswere removed, while on the bare soil plots the highest aphid populations were caughtduring the first 50 days after sowing (Fig. 1). The use of TPM resulted in fewer aphids

496



per trap until 41 days after sowing. However, after 50 days, the numbers of trappedaphids were about the same for all the treatments.

Sweetpotato whitefly populations were extremely high, mainly in the bare soilcontrol plots, where 52, 35 and 32 adults per leaf were recorded 25, 32 and 38 days,respectively, after sowing (Fig. 2). Populations of whitefly nymphs and adults were ex-cluded from the melon plants while the polypropylene row cover was on the bed.Transparent polyethylene mulch reduced the number of whitefly adults per leaf withrespect to bare soil on some dates during the first 50 days after sowing. In plants withFRC, alone or combined with TPM removed at the reproductive stage, no whiteflynymphs were detected for 35 and 42 days after sowing (Table 1), while leaves from theplants with FRC removed at the vegetative stage had no nymphs at 35 days, and onlya few nymphs (4.5 per cm

2

) at 42 days. The TPM plots significantly reduced the num-ber of nymphs (about one half) compared to the control plots. In contrast, bare soilplots had significantly greater numbers of whitefly nymphs on both dates than theFRC and TPM treatments.

The appearance of virus disease symptoms were delayed one week on plants whenFRC was removed at vegetative stage, and symptoms were delayed 2-3 weeks whenFRC was removed at reproductive stage (Fig. 3). At harvest (67 days after sowing), thecontrol plots had the highest percentage (24%) of virus-infected plants followed byFRC on bare soil plots (16%). The TPM plots and the TPM + FRC plots showed thelowest (5-12%) number of virus-diseased plants. ELISA tests indicated that PRSV-Wwas the most prevalent virus in this study, however, ZYMV, CMV, WMV-2 and SqMVwere also detected. The PRSV-W, ZYMV, CMV and WMV-2 are transmitted in a non-persistent manner by some aphid species, while SqMV is transmitted by beetles(Nameth et al, 1986).

Leafminer damage was avoided for 35 and 42 days after sowing with FRC removedat the vegetative and reproductive stage, respectively, compared with bare soil (Fig.

Figure 1. The number of winged aphids collected in yellow water pan traps placedin cantaloupe plots with transparent mulch, floating row cover and bare soil. VS andRS = row cover removed at vegetative and reproductive stage, respectively.


497

4). During the first 35 days, control and TPM plots had the highest leafminer damage;however, by 52 days the percentage of damaged leaves was similar in all the treat-ments.

Yield of cantaloupe melons, expressed as mean number and weight of fruit was sig-nificantly higher for TPM + FRC plots than TPM and bare soil with or without FRC(Table 2). The yield of TPM + FRC (removed during reproductive stage) plots was

T

ABLE

1. E

FFECT

OF

T

RANSPARENT

M

ULCH

AND

F

LOATING

R

OW

C

OVER

ON

THE

N

UM-BER

OF

W

HITEFLY

N

YMPHS

ON

CANTALOUPE LEAVES. TECOMAN VALLEY,COLIMA, MEXICO. 1993.

Nymphs/cm2

Treatment26 March(35 DAS1)

2 April(42 DAS1)

Transparent polyethylene mulch (TPM) 2.7 b4 9.1 c4

Floating row cover (VS2) 0.0 a 4.5 bFloating row cover (RS3) 0.0 a 0.0 aTPM + Floating row cover (RS) 0.0 a 0.0 aBare soil (Control) 5.4 c 19.1 d

1DAS = Days after sowing.2VS = Removed at vegetative stage.3RS = Removed at reproductive stage.4 = Mean separation within columns by Duncan’s multiple test (P < 0.05).

Figure 2. Effect of transparent mulch and floating row cover on whitefly popula-tions in cantaloupe foliage. VS and RS = Row cover removed at vegetative and repro-ductive stage, respectively.


nearly four-fold higher than in control plots, and three-fold higher than with FRCalone. Moreover, the yield obtained in TPM plots and bare soil with FRC (removedduring vegetative growth) was higher than that on bare soil plots.

The results of this study demonstrate the beneficial effects of FRC alone or com-bined with reflective mulch for cantaloupe grown in dry tropical conditions. Floatingrow covers excluded beetles, leafminers, sweetpotato whitefly, and aphids. The exclu-sion of insect pests reduced the use of insecticides by 50%. In addition, the foliage ofplants was not colonized by whiteflies during the first 42 days with FRC removed atthe reproductive stage. Furthermore, TPM resulted in fewer winged aphids caught inyellow traps and fewer whitefly nymphs and adults on foliage with respect to control.These results with FRC are similar to previous research (Natwick & Durazo 1985,Natwick & Laemmlen 1993, Orozco et al. 1994, Perring et al. 1989, Webb & Linda1992). The repellent effect of the plastic mulch on aphids (Brown et al. 1993, Jones1991) and whitefly (Kelly et al. 1989) has been attributed to its high reflectance oflight (Kring & Schuster 1992).

Virus disease incidence was low in the control plots; this may be related to the lowaphid populations caught in yellow traps (maximum 65 aphids per trap per week). Inthis experiment, the major viruses found were PRSV-W, ZYMV, CMV and WMV-2which are transmitted by aphids (Nameth et al. 1986). Because of exclusion of aphidvectors in covered plots, and the fewer numbers of aphids alighting on the plantsgrown in TPM plots, TPM and FRC had a lower virus incidence compared with thecontrol. These data support the observations of Brown et al. 1993, Kring & Schuster1992, Natwick & Durazo 1985, Orozco et al. 1994, Perring et al. 1989, and Webb &Linda 1992.

The yield in weight of cantaloupe plants grown with TPM + FRC was higher thanthat of TPM mulch alone and bare soil. This can be attributed to more vigorousgrowth of plants in TPM plots alone or combined with FRC because of moisture con-

Figure 3. Effect of transparent mulch and floating row cover on incidence of can-taloupe plants with virus symptoms. VS and RS = row cover removed at vegetativeand reproductive stage, respectively.

Orozco-Santos et al.: Floating Row Cover & Transparent Mulch 499

servation and higher soil temperature. Preliminary studies have shown a yield in-crease of cantaloupe with the use of TPM (Orozco et al. 1994) and FRC (Orozco et al.1994, Perring et al. 1989, Webb & Linda 1992). Also, the effects of transparent mulchon soil temperature (Al-Assir et al. 1992) and moisture conservation (Tindall et al.1991) have been previously demonstrated.

The beneficial effects of polyethylene transparent mulch and floating row covermight be a practical and economical management tool for producing cantaloupe underthe dry tropical conditions of Colima, Mexico. However, additional on-farm trials are

TABLE 2. EFFECT OF TRANSPARENT MULCH AND FLOATING ROW COVER ON THE NUM-BER OF FRUIT AND YIELD OF CANTALOUPE. TECOMAN VALLEY, COLIMA, MEX-ICO. 1993.

No. of Fruit per6.9 m of Row1

Weight (kg) of Fruit per

6.9 m of Row1

Transparent polyethylene mulch (TPM) 33.0 c 28.37 cFloating row cover (VS2) 29.5 c 24.69 cFloating row cover (RS3) 54.2 b 47.04 bTPM + Floating row cover (RS) 68.6 a 62.51 aBare soil (Control) 19.2 d 16.08 d

1 = Mean separation within columns by Duncan’s multiple range tests (P < 0.05).2VS = Removed at vegetative stage.3RS = Removed at reproductive stage.

Figure 4. Effect of transparent mulch and floating row cover on the percentage ofcantaloupe leaves damaged by the vegetable leafminer. VS and RS = row cover re-moved at vegetative and reproductive stage, respectively.


needed to determine whether or not these methods can be practically integrated intocommercial cantaloupe production.

ACKNOWLEDGMENTS

This study was supported by funds from the Patronato para la Investigación y Ex-perimentación Agrícola del Estado de Colima, México (PIEAEC) and Instituto Nacio-nal de Investigaciones Forestales y Agropecuarias (INIFAP).

REFERENCES CITED

ADAMS, R. G., R. A. ASHLEY, AND M. J. BRENNAN. 1990. Row covers for excluding in-sect pests from broccoli and summer squash plantings. J. Econ. Entomol. 83:948-954.

AL-ASSIR, I. A., I. G. RUBEIZ, AND R. Y. KHOURY. 1992. Yield response of greenhousecantaloupe to clear and black plastic mulches. Biological Agriculture and Hor-ticulture 8:205-209.

BASKY, Z. 1993. Identification key for alate aphids caught in yellow pan traps. ActaPhytopathologica et Entomologica Hungarica, 28: 71-121.

BROWN, J. E., J. M. DANGLER, F. M. WOODS, K. M. TILT, M. D. HENSHAW, W. A.GRIFFEY, AND M. S. WEST. 1993. Delay in mosaic virus onset and aphid vectorreduction in summer squash grown on reflective mulches. HortScience 28:895-896.

BUTLER, G. D., JR. 1982. Sweetpotato whitefly, Bemisia tabaci, a new pest of cotton inArizona. Ariz. Agr. Expt. Sta. P-56: 49-50.

BUTLER, G. R., AND T. J. HENNEBERRY. 1991. Effect of oil sprays on sweetpotato white-fly and phytotoxicity on watermelons, squash and cucumbers. SouthwesternEntomologist 16:63-72.

BYRNE, D. N., T. S. BELLOWS JR., AND M. P. PARRELLA. 1990. Whiteflies in agricul-tural systems, pp. 227-261 in D. Gerling [ed.] Whiteflies: their bionomics, peststatus and management. Intercept. Great Britain.

CLARK, M. F., AND A. N. ADAMS. 1977. Characteristics of the microplate method of en-zyme-linked immunosorbant assay for the detection of plant viruses. JournalGen. Virol. 34:475-483.

COHEN, S. 1990. Epidemiology of whitefly-transmitted viruses, pp. 211-225 in D. Ger-ling. [ed.] Whiteflies: their bionomics, pest status and management Intercept.Great Britain.

DELGADILLO, S. F., T. J. A. GARZON, AND P. A. VEGA. 1989. Cucurbit viruses in Mexico:A survey. Rev. Mex. de Fitopatología 7:136-139.

FARIAS-L., J., M. OROZCO-S., AND S. GUZMAN-G. 1994. Effect of Imidacloprid onsweetpotato whitefly (Homoptera: Aleyrodidae) populations on cantaloupemelon. XXIX Cong. Nal. de Entomología. Annual meeting southwesternbranch-ESA. Monterrey, N.L., México. p.188-189.

GILL, R. J. 1992. A review of the sweetpotato whitefly in southern California. Pan-Pa-cific Entomologist 68:144-152.

JONES, R. A. C. 1991. Reflective mulch decreases the spread of two non-persistentlyaphid transmitted viruses to lupin (Lupinus angustifolius). Ann. appl. Biol.118:79-85.

KELLY, J. W., P. H. ADLER, D. R. DECOTEAU, AND S. LAWRENCE. 1989. Colored reflec-tive surfaces to control whitefly on poinsettia. HortScience 24:1045.

KRING, J. B., AND D. J. SCHUSTER. 1992. Management of insects on pepper and tomatowith UV-reflective mulches. Florida Entomol. 75:119-129.

LECOQ, H., AND M. PITRAT. 1983. Field experiments on the integrated control ofaphid-borne viruses in muskmelon, p. 169-176 in R. T. Plumb and J. M. Thresh[eds.] Plant virus epidemiology. Blackwell Scientific Publication. Oxford, Eng-land.

Orozco-Santos et al.: Floating Row Cover & Transparent Mulch 501

NAMETH, S. T., J. A. DODDS, A. O. PAULUS, AND F. F. LAEMMLEN. 1986. Cucurbit vi-ruses of California: An ever-changing problem. Plant Disease 70:8-12.

NATWICK, E. T., AND A. DURAZO. 1985. Polyester covers protect vegetables from white-flies and virus disease. California Agric. 39:21-22.

NATWICK, E. T., AND F. F. LAEMMLEN. 1993. Protection from phytophagous insectsand virus vectors in honeydew melons using row covers. Florida Entomol.76:120-126.

OROZCO-S., M., O. LOPEZ-A., O. PEREZ-Z., AND F. DELGADILLO-S. 1994. Effect of trans-parent mulch, floating row cover and oil sprays on insect populations, virus dis-eases and yield of cantaloupe. Biological Agriculture and Horticulture 10:229-234.

PEÑA, M. R. 1991. Manual de prácticas del curso: Afidos como vectores de virus. Cen-tro de Fitopatología. Colegio de Postgraduados. Montecillo, México. 148 p.

PERRING, T. M., A. COOPER., D. J. KAZMER, C. SHIELDS, AND J. SHIELDS. 1991. Newstrain of sweetpotato whitefly invades California vegetables. California Agric.45:10-12.

PERRING, T. M., R. N. ROYALTY, AND C. A. FARRAR. 1989. Floating row covers for theexclusion of virus vectors and the effect on disease incidence and yield of can-taloupe. J. Econ. Entomol. 82:1709-1715.

PIKE, K. S., L. BOYDSTON, AND D. ALLISON. 1990. Alate aphid vivipare associated withsmall grains in North America: A key and morphometric characterization. J.Kansas Entomol. Soc. 63:559-602.

RAMIREZ, V. J., AND V. C. LOPEZ. 1991. Efecto de cubiertas flotantes y acolchado deplástico sobre el control de virosis e incremento de rendimientos en pepino ymelón. Memorias del XVIII Cong. Nal. de Fitopatología. Puebla, México. p. 205.

SCHUSTER, D. J., AND J. F. PRICE. 1987. The western flower thrips and the sweetpo-tato whitefly: New pests threatening Florida tomato production. Univ. of Fla.,IFAS, Bradenton GCREC Res. Rep. BRA1987-16.

SIMONS, J. N., AND T. A. ZITTER. 1980. Use of oils to control aphid-borne viruses. PlantDisease 64:542-546.

TINDALL, J. A., R. B. BEVERLY, AND D. E. RADCLIFFE. 1991. Mulch effect on soil prop-erties and tomato growth using micro-irrigation. Agron. J. 83:1028-1034.

WEBB, S. E., AND S. B. LINDA. 1992. Evaluation of spunbonded polyethylene row cov-ers as a method of excluding insects and viruses affecting fall-grown squash inFlorida. J. Econ. Entomol. 85:2344-2352.

502



PROBABILITY OF DETECTING CARIBBEAN FRUIT FLY (DIPTERA: TEPHRITIDAE) INFESTATIONS BY FRUIT

DISSECTION

W

ALTER

P. G

OULD

USDA-ARS, Subtropical Horticulture Research Station,Miami, FL

A

BSTRACT

Quarantine inspectors search for fruit fly infestations in incoming shipments byvisual inspection and by dissecting or cutting a sample of fruit in each shipment. Thereliability of the latter procedure for detecting fruit fly larvae is questionable and,therefore, a test was conducted to determine its effectiveness. Infested grapefruit,mangoes, guavas, and carambolas were cut open to determine the efficacy of cuttingfruit in detecting larval infestations of Caribbean fruit flies,

Anastrepha suspensa

(Loew). From 1 to 36% of the larvae were detected by dissection, but 17.9 to 83.5% ofthe infested fruit were detected. The percentage of Caribbean fruit fly larvae presentin an infested fruit that can be detected by cutting varies with the type of fruit in-fested. The overall percentage of infested fruit detected also varies in the same man-ner. Carambolas were the easiest fruit in which to find larvae and green guavas themost difficult. Overall, only 9.5% of the larvae in field-infested guavas and 16.9% ofthose in cage-infested fruits were detected. Between 17.9% (green guavas) and 83.5%(carambolas) of the infested fruit was detected. There was considerable variation inthe number of larvae found by different inspectors.

Key Words: Caribbean fruit fly,

Anastrepha suspensa

, larval detection.

R

ESUMEN

Los inspectores cuarentenarios detectan infestaciones de moscas de las frutas bus-cando frutas dañadas y cortando una muestra de ellas en un determinado carga-mento. Se disectaron toronjas, mangos, guayabas, y carambolas infestadas paradeterminar la eficiencia del corte de frutas en la deteccción de la infestación larval de

Anastrepha suspensa

(Loew). Del 1 al 36% de las larvas fueron detectados mediantela disección; sin embargo, este método permitió encontrar del 17.9 al 83.5% de las fru-tas infestadas. El porcentaje de larvas de moscas de las frutas del Caribe detectadocortando frutas infestadas varió con los diferentes tipos de frutas. El porcentaje defrutas infestadas varió de la misma manera. La fruta en la cual fue más fácil encon-trar larvas fue la carambola y la más difícil la guayaba verde. En resumen, solamentefue detectado el 9.5% de las larvas en las guayabas, y el 16.9% en las frutas que fueroninfestadas en jaulas. Fue detectado entre el 17.9% (guayabas verdes) y el 83.5% (ca-rambolas) de las frutas infestadas. Hay considerable variación en el número de larvas

encontradas por diferentes inspectores.

Prevention of the spread of undesirable pests is a worldwide problem and quaran-tines have been established for many pest organisms. The family Tephritidae includesmany species of fruit flies whose larvae attack valuable cultivated fruits and vegeta-bles. Two dozen important species and hundreds of species of minor importance existaround the world (Robinson & Hooper 1989). Most fruit flies lay their eggs under theskins of fruits and vegetables. The larvae burrow and feed inside the fruit or vegetable

Gould: Fruit Fly Detection by Fruit Cutting

503

making it unpalatable and accelerating decay. Many countries have plant protectionagencies whose mission is to exclude these unwelcome pests (Shannon 1994). Themain line of defense is the vigilance of inspectors who check incoming fruit and vege-table shipments. The main method of detecting fruit flies in a shipment of a commod-ity is to sample a number of fruits or vegetables and examine them for possible signsof infestation, a procedure that often includes cutting open some of the fruits and ex-amining them for eggs and/or larvae (Anon. 1993).

Cutting fruits and vegetables to detect insect infestation is time consuming andonly a fraction of a shipment can be inspected in this way, but there are no feasible al-ternatives. Holding the commodity until any larvae emerge is not practical because itwould hold up the shipment for too long and destroy its value. Other methods, suchas detecting sound of feeding larvae, have not yet proven practical (Calkins & Webb1988, Hansen et al. 1988, Hansen et al. 1992, Harrison et al. 1993).

Many factors can affect the probability of finding a larva, such as the ability of theinspector, the stage of the insect present, and the type of fruit. In this study, I deter-mined the probability of detecting Caribbean fruit fly,

Anastrepha suspensa

(Loew),larvae given three variables. I looked at four types of fruit, different stages of ripe-ness, and variations in the efficiency of the inspectors.

M

ATERIALS

AND

M

ETHODS

Four different types of fruit were used. Guavas were obtained from J. R. Brooksand Son, a local packing house, and ripe guavas were harvested from trees on theUSDA station in Miami. Mangoes and carambolas were also obtained from J. R.Brooks and Son. Grapefruits were obtained from Bernard Egan & Co., a packinghouse in central Florida. Guavas received from the packing house were separated intoripe (yellow) and immature (green). The guavas had a significant field infestation.Grapefruits, mangoes, and carambolas did not have a detectable infestation in thefield so they were exposed to approximately 100,000 Caribbean fruit flies in an out-door cage for 1 to 3 days (carambolas and mangoes), or 7 days (grapefruit). Field-in-fested guavas were cut open 1 day after being received from the packing house orbeing picked and could contain all stages of eggs and larvae. Mangoes were held 5 to13 days after infestation before being cut open, and grapefruit and carambolas wereheld for 7 days after infestation before cutting. These holding periods allowed devel-opment of larvae to the third instar and precluded presence of eggs and early instars.

The fruits were randomly divided into a control and a treatment group. The num-ber of fruits used per replicate depended on the number available, between 15 and 150per replicate, and the experiment was replicated three to five times for each type offruit.

To compare effect of fruit type and ripeness, cutting was done by one inspector us-ing the following guidelines: Each fruit was cut with a 15-cm utility knife into 1-cmthick slices (from 5 to 10 depending upon fruit size). Mangoes were sliced from eachside until the seed was reached, then turned 90

°

and sliced again until the seed wasreached. Each slice was examined for the presence of Caribbean fruit fly larvae, andthe total number observed was recorded. Sliced fruits with larvae and unsliced controlfruits were placed in 1-liter plastic buckets containing vermiculite and held for sev-eral weeks. The vermiculite was then sifted for larvae and pupae.

Another experiment was conducted to examine the effect of the inspector on theprobability of detection. Five additional inspectors were recruited for the test. Allwere either active employees of the Florida Department of Agriculture & ConsumerServices, Division of Plant Industry (DPI) or ex-DPI members who had routinely cutfruits looking for larvae. The five DPI or ex-DPI inspectors and the original inspector

504



each cut 10 or 20 infested grapefruit (fly cage infested). The five DPI inspectors wereallowed to cut fruit any way they chose. The original inspector followed the previousguidelines.

Percentage of larvae detected by cutting and percentage of infested fruits detectedby cutting were transformed using the angular transformation (arcsin) and bothtransformed and untransformed data analyzed using unbalanced ANOVA by GLM(SAS Institute 1988). Untransformed percentages are presented in the tables for com-parison. If the ANOVA F-test was significant at the 0.05 level, the Waller-DuncanK-ratio t-test (SAS Institute 1988) was used to separate means. T-test and Proc Regwere also used (SAS Institute 1988).

R

ESULTS

AND

D

ISCUSSION

There was no significant difference between the number of larvae emerging fromcontrol and the cut fruits (paired t-test, P=.998,

α

=0.05). Therefore, very few larvaewere physically damaged by the cutting, and tearing the fruit open did not affect thelarvae’s ability to mature and leave the fruit.

The infestation levels in the fruits varied (Table 1). Green guavas from Brooks hadthe lowest infestation level, ripe guavas from Brooks slightly higher, and ripe guavasfrom the USDA station, much higher levels. The infestations from the cage varied aswell. In most of the replications, the infestations were low (one or fewer larvae perfruit); however, there were several with moderate infestations and several with ex-tremely high infestations.

There was considerable variation between replications in the percentage of larvaedetected by cutting for all fruit types (Table 2). Green guavas overall had the lowestpercentage of larvae detected (1.1%). Carambolas had the highest (about 36%). Only9.5% of the larvae in field-infested guavas were detected by cutting. Only 16.9% of thelarvae in all of the cage-infested fruits (carambolas, grapefruits, mangoes) togetherwere detected by cutting.

From 64 to 98% of larvae escaped detection. However, since many larvae were of-ten found in one fruit, there was a higher probability of an infested fruit being de-

T

ABLE

1. F

RUIT

I

NFESTATION

L

EVELS

OF

A.

SUSPENSA

(L

ARVAE

PER

F

RUIT

).

Field-Infested Guavas Replication Mean

1 2 3 4 5

Green-Brooks 6.000 0.167 0 0.200 0 1.273Ripe-Brooks 8.662 0.967 0.240 0.200 — 2.517Ripe-USDA station 39.120 38.920 13.320 — — 30.453

Cage-Infested Fruits Replication Mean

1 2 3 4

Grapefruits 0.633 3.840 0.680 58.580 15.933Mangoes 225.750 0.100 0.750 1.600 57.050Carambolas 20.220 11.000 0.320 — 10.513


505

tected than Table 2 would indicate (Table 3). As many as 83.5% of carambolas withlarvae were detected. This is probably because carambolas are a fairly translucentfruit and larvae are more easily observed. Green guavas were the most difficult inwhich to detect larvae; only 17.9% of the infested fruit were detected. This may be be-cause they were inspected one day after we received them, thus the fruit flies could bein the egg stage or very early instar and very difficult to see. This fruit is also toughand difficult for the inspector to cut up. Analysis of variance found that in the percent-age of infested fruit detected, there was a significant difference only between the car-ambolas and all of the other fruit, except ripe guavas from Brooks. None of the otherpairwise comparisons was significant.

The effect of size of infestation and the percentage of larvae found did not show ahigh correlation, but generally the more larvae that were present, the lower the per-centage found by the inspector. This may be due to a fatigue factor in those extremeinfestations where thousands of larvae were present. Taking log ten of the number oflarvae present makes the distribution of the percentage of larvae detected look likethe normal distribution. The best regression had an r

2

of 0.11; with an outlier deleted,it improved only to 0.29.

T

ABLE

2. P

ERCENTAGE

OF

A.

SUSPENSA

L

ARVAE

D

ETECTED

BY

C

UTTING

.

Fruit type Replication Mean

1

1 2 3 4

Carambolas 9.0 35.8 62.5 — 35.8aRipe Guavas-Brooks 29.6 13.5 25.6 — 22.9abGrapefruits 15.8 7.8 11.8 2.8 9.5bcMangoes 1.9 0.0 20.0 0.0 5.5bcRipe Guavas-USDA station 4.7 0.0 9.0 — 4.6bcGreen Guavas-Brooks 1.1 2.2 0.0 — 1.1c

1

Means within a column followed by the same letter are not significantly different by ANOVA followed byWaller-Duncan K-ratio t-test.

T

ABLE

3. P

ERCENTAGE

OF

A.

SUSPENSA

I

NFESTED

F

RUITS

D

ETECTED

BY

C

UTTING

.

Fruit Type Replication Mean

1

1 2 3 4 5

Carambolas 60.5 90.0 1 — — 83.5aRipe Guavas-Brooks 77.8 66.7 60.0 0.0 — 51.1abRipe Guavas-USDA station 66.7 36.0 48.0 36.0 — 46.7bGrapefruits 23.3 22.2 58.0 — — 34.5bMangoes 85.0 0.0 28.6 0.0 — 28.4bGreen Guavas-Brooks 7.2 22.3 0.0 60.0 0.0 17.9b

1

Means within a column followed by the same letter are not significantly different by ANOVA followed byWaller-Duncan K-ratio t-test on arcsin transformed data. Untransformed means are presented.

506



In the inspector comparison, the percentage of larvae found varied from 8% to 49%(Table 4). Most of the difference could be attributed to the methods each inspectorused. Inspector 1 was obviously hurrying and found the fewest larvae. Inspector 2shredded the fruit into tiny pieces, took much longer than the others, and found morelarvae. The other three inspectors’ results were very similar. Because the inspectorsknew their results were being compared and studied, they probably found more lar-vae than they would have otherwise. The main reason for performing this test was todetermine the relationship between the method and the inspector so we could assessthe results of the first part of this study compared to ‘real world’ inspectors. We con-cluded that the data obtained with the inspector in the first part of this study (usinga consistent cutting method) were reasonably comparable to those of the DPI inspec-tors.

In summary, the percentage of Caribbean fruit fly larvae present in an infestedfruit that can be detected by cutting varied with the type of fruit infested. The overallpercentage of infested fruit detected also varied in the same manner. Carambolaswere the easiest fruit in which to find larvae and green guavas the most difficult.Overall, only 9.5% of the larvae in guavas and 16.9% of those in cage-infested fruitswere detected. Between 17.9 and 83.5% of the infested fruit was detected. There wasconsiderable variation among inspectors based on the method used to inspect thefruit.

A

CKNOWLEDGMENTS

I thank Craig Campbell, J. R. Brooks and Son, Inc., and Bernard Egan and Co. forcontributing fruit used in this study. I also thank D. Chalot, W. Francillon and L.Lodyga, of the Florida Department of Agriculture and Consumer Services, Division ofPlant Industry for their assistance. I thank W. Montgomery, R. Pantaleon, and D.Storch of USDA-ARS for their assistance. I thank G. Hallman and M. Hennessey ofUSDA-ARS for their critical review of this manuscript, and also G. Hallman and E.Schnell of USDA-ARS for translation of the abstract to Spanish.

R

EFERENCES

C

ITED

A

NONYMOUS

. 1993. Animal Plant Health Inspection Service. Plant Import Manual:Nonpropagative. U.S. Govt. Printing Office. Washington, D.C.

C

ALKINS

, C. O.,

AND

J. C. W

EBB

. 1988. Temporal and seasonal differences in move-ment of the Caribbean fruit fly larvae in grapefruit and the relationship to de-tection by acoustics. Florida Entomol. 71: 409-416.

T

ABLE

4. P

ERCENTAGE

OF

A.

SUSPENSA

L

ARVAE

D

ETECTED

BY

C

UTTING

BY

D

IFFERENT

I

NSPECTORS

.

Inspector 1 08.4Inspector 2 49.1Inspector 3 29.5Inspector 4 17.8Inspector 5 17.7Inspector 6 16.3

Mean 23.1St. Dev. 14.4


507

H

ANSEN

, J. D., C. L. E

MERSON

,

AND

D. A. S

IGNOROTTI

. 1992. Visual detection of sweet-potato weevil by non-invasive methods. Florida Entomol. 75: 369-375.

H

ANSEN

, J. D., J. C. W

EBB

, J. W. A

RMSTRONG

,

AND

S. A. B

ROWN

. 1988. Acoustical de-tection of oriental fruit fly (Diptera: Tephritidae) larvae in papaya. J. Econ. En-tomol. 81: 963-965.

H

ARRISON

, R. D., W. A. G

ARDNER

, W. E. T

OLLNER

,

AND

D. J. K

INARD

. 1993. X-ray com-puted tomography studies of the burrowing behavior of fourth-instar pecanweevil (Coleoptera: Curculionidae). J. Econ. Entomol. 86: 1714-1719.

R

OBINSON

, A. S.,

AND

G. H

OOPER

, (eds.) 1989. Fruit Flies, Vol. 3A World Crop Pests.Elsevier, Amsterdam. 372 pp.

SAS I

NSTITUTE

. 1988. SAS/STAT User’s Guide. Release 6.03 Ed. SAS Institute, Cary,NC.

S

HANNON

, M. J. 1994. APHIS, pp. 1-10

in

J. L. Sharp and G. J. Hallman [eds.] Quar-antine treatments for pests of food plants. Westview Press, Boulder, Colorado.

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

290 pp.

Smith & Zungoli: Termites and Rigid Board Insulation

507

RIGID BOARD INSULATION IN SOUTH CAROLINA: ITS IMPACT ON DAMAGE, INSPECTION AND CONTROL OF

TERMITES (ISOPTERA: RHINOTERMITIDAE)

B. C. S

MITH

AND

P. A. Z

UNGOLI

Urban Entomology Research LaboratoryClemson University

Clemson, SC 29634 0365

A

BSTRACT

A total of 225 pest control companies was randomly selected from the 315 mem-bers of the South Carolina Pest Control Association to determine their opinion on theimpact of rigid board construction on termite control. Of those surveyed, 77% con-ducted inspections and/or treatments for subterranean termites (Isoptera: Rhinoter-mitidae,

Reticulitermes

and

Coptotermes spp.

) in South Carolina. The focus of surveyquestions was on demographics, occurrence of rigid board insulation, prevalence ofrigid board insulation damage and structural damage, and treatment of structureswhen rigid board insulation was present. Results indicated termite infestations asso-ciated with rigid board insulation are not uncommon; 34% of the companies reportedthe presence of rigid board insulation on structures that have been treated or in-spected for termites. Companies chose not to treat structures with rigid board insula-tion 43% of the time. Twenty-five percent (25%) of the companies reported damage torigid board insulation due to insects other than termites. In addition, 12% of the re-spondents have been sued because of termite damage hidden by rigid board insula-tion. The primary recommendation to solve these problems is to remove a small gapof insulation, a vision strip, just above the soil level along the perimeter of a structure.

Key Words: Rigid board insulation, polystyrene, polyisocyanurate,

Reticulitermes

spp.,

Coptotermes

spp.

R

ESUMEN

Una muestra de 225 compañías de control de plagas fue aleatoriamente seleccio-nada de entre los 315 miembos de la Asociación de Control de Plagas de Carolina del

508



Sur, para una encuesta en Febrero de 1994. De las compañías encuestadas que res-pondieron, el 77% se dedica a inspecciones o tratamientos para termites subterráneos(Isoptera: Rhinotermitidae,

Reticulitermes

y

Coptotermes

spp.) en Cariolina del Sur.El foco de las preguntas de la encuesta fue la demografía, aparición y prevalencia dedaños en estructuras rígidas de aislamiento térmico, el daño estructural, y el trata-miento de las estructuras con aislamiento térmico rígido. Los resultados indicaronque las infestaciones de termites asociadas con estructuras rígidas de aislamiento tér-mico son comunes; el 34% de las compañías reportó la presencia de aislamiento tér-mico rígido en las estructuras que habían sido tratadas o inspeccionadas. Lascompañías decidieron no tratar estructuras con aislamiento térmico rígido en el 43%de los casos. El 25% de las compañías reportó daños a estructuras rígidas de aisla-miento térmico debido a otros insectos. En adición, el 12% de los que respondieron laencuesta había sido demandado debido a daños de termites ocultos por las estructurasrígidas de aislamiento térmico. La recomendación primaria para resolver estos pro-blemas es quitar un pequeño pedazo del aislamiento, una banda de visión, justamente

por encima del nivel del suelo a lo largo del perímetro de la estructura.

In the last few years, there has been an increasing awareness of the use of rigidboard insulation to increase the energy efficiency of structures. Great concern hasbeen expressed by Pest Control Operators (PCOs) with regard to their ability to prop-erly inspect and treat these structures for subterranean termites (

Reticulitermes

spp.and

Coptotermes

spp.), and other insects to the extent that the National Pest ControlAssociation has issued guidelines for its members (Kramer 1993). Problems with ter-mite treatments in the presence of rigid board insulation have also recently gained at-tention in the literature (Guyette 1994).

Rigid board insulation, primarily foam board insulation, includes materials suchas polystyrene (styrofoam) and polyisocyanurate. Rigid board insulation is used as aperimeter insulation around structures; the insulation is usually placed against thefoundation walls on the exterior of the masonry structure hidden from view by skincoats such as wood, aluminum or vinyl siding, or stucco. When rigid board insulationis in direct contact with the ground, it is essentially the same as wood-to-ground con-tact. The presence of these materials creates a situation which is conducive to termiteinfestations because of continuous hidden pathways directly into the wooden mem-bers of a structure. This makes it extremely difficult for the PCO to visually inspectand treat for termites. In addition, these materials are associated with an increase inmoisture levels, which allows for highly favorable conditions to harbor termites.

Rigid board is being used as insulation due to low costs and good energy savingqualities. In 1985, approximately 5% of the building stock contained rigid board insu-lation (Christian 1990). By 1992, 50% of new construction contained some rigid boardinsulation (Kramer 1993). Good thermal characteristics, as well as highstrength-to-weight ratios, have increased the use of these materials (Strzepek 1990).However, the strength and hardness of the rigid board may also be adversely affectedby termite excretions (Building Research Station 1966). Hickin (1971) reported thatplastic materials do not serve as a source of nutrition for termites, and damage ofthese materials is from termites physically removing the rigid board from their path.Plastics and other materials are not passed through the digestive system, rather thebitten-off fragments are removed or incorporated into tunnels. Polystyrene is subjectto severe termite damage (Gay & Wetherly 1969). Tests showed that exposed ends andedges are more prone to attack than flat or curved surfaces. Cracks, rough edges,holes, and other deviations may promote termite activity on plastic materials (Becker


509

1972). The Formosan subterranean termite,

Coptotermes formosanus

Shiraki, dam-aged soft styrene resins in building materials (Shimizu et al. 1970).

The purpose of this survey was to demonstrate problems associated with the pres-ence of rigid board insulation in South Carolina, primarily with termites, but alsoother insects. Surveys were sent to owners, managers, PCOs, and other persons asso-ciated with pest control companies to gain their perspective on the extent to which theproblem exists. Recommendations to solve these problems, compiled from survey re-sponses, will be discussed.

M

ATERIALS

AND

M

ETHODS

A sample of 225 individuals was chosen from the 315 pest control companies be-longing to the South Carolina Pest Control Association (SCPCA); a total of 775 com-panies are operating in South Carolina. Survey forms were mailed in February, 1994,in a timed and organized mailing strategy and numbered to facilitate identification ofreturned surveys. The survey requested information for 1993 only. One week follow-ing the initial mailing, a follow-up letter was sent as a reminder to complete and to re-turn survey forms. Three weeks following the initial mailing, a repeat survey formwas mailed to all non-respondents. At eight weeks, another survey form was mailedto all non-respondents. All analyses of the data were performed using SAS (SAS In-stitute 1985). Results are primarily descriptive statistics.

The content of questions consisted of four types: demographics, occurrence of rigidboard insulation, prevalence of rigid board insulation damage and structural damage,and treatment of structures when rigid board insulation was present.

R

ESULTS

AND

D

ISCUSSION

Demographics

Of 225 survey forms, 187 were returned (83%); of those, 173 (77%) represented re-spondents who conduct termite inspections and/or treatments. Respondents werestratified by three regions within the state; Piedmont (the upper northwest portion),Central (middle one-third portion), and Coastal (lower southeastern portion) (Fig. 1).These regions were selected based on climatic differences within the state. Of thoserespondents who performed termite inspections and/or treatments, 17% were in thePiedmont, 38% in the Central, and 40% in the Coastal regions, with 5% in combina-tions of the three regions.

Most respondents (73%) were owners of the pest control companies surveyed, sug-gesting that information obtained was reliable. The number of termite treatmentsand inspections performed, in addition to the number of technicians employed, can befound in Table 1. Most companies were relatively small with less than 5 termite tech-nicians employed. However, this does not imply the operations were small, as evi-denced from the number of inspections and treatments performed in a given year:over three-quarters (82%) of all the termite treatments were performed by companieswith 5 or fewer technicians.

Occurrence of Rigid Board Insulation

The documented presence of rigid board insulation on structures treated or in-spected for termites was reported by 34% of the companies. Of these companies, 17%reported rigid board insulation on structures in the Piedmont, 39% in the Central,and 39% in the Coastal regions, with 5% in combinations of the three regions. These

510



values might not necessarily indicate the amount of new housing construction in eachregion, but certainly might reflect the increased termite pressures farther south.

As many as 81% of companies reported up to 10% of structures they treated and/or inspected for termites had rigid board insulation, and 2% of the companies reportedgreater than 75% of the structures had rigid board insulation. Rigid board insulationwas reported to be installed above grade on structures by 41% of the companies, andbelow grade on structures by 59% of the companies.

Prevalence of Rigid Board Insulation Damage and Structural Damage

Eighty-one percent of the companies reported termite damage to rigid board insu-lation installed above grade in up to 10% of the structures they inspected.Seventy-two percent of the companies reported termite damage to rigid board insula-tion installed below grade in up to 10% of the structures they inspected, and 9% of thecompanies reported termite damage to rigid board insulation installed below grade ingreater than 75% of the structures (Table 3).

Termite damage to the wooden structural members in the presence of rigid boardinsulation installed both above and below grade was reported by the occurrence onstructures and percent time seen by companies (Table 4).

Excessive termite damage to rigid board insulation was seen by 10% of the compa-nies, whereas moderate and slight damage to insulation was seen by 40% and 50% ofthe companies, respectively. Excessive termite damage to the wooden members of a

Figure 1. Regions of South Carolina.


511

structure was seen by 30% of the companies; moderate and slight damage was seenby 40% and 30% of the companies, respectively.

The amounts of termite damage to rigid board insulation and wooden structuralmembers, at various locations on a structure, was reported by mean percent damage(Table 5). The amount of damage in wooden members is positively correlated to theamount of damage seen in rigid board insulation in most locations throughout a struc-ture (Table 6).

When the following foundation types were encountered, termite damage to rigidboard insulation was present in the following amounts: monolithic slab (33%), sup-ported slab (9%), floating slab (19%), basement (7%), crawlspace (23%), and any com-bination of one or more of the previous types (9%) (Table 7).

Twenty-five percent of the companies reported damage to rigid board insulationcaused by insects other than termites. Carpenter ants were responsible for 87% of thedamage, while other insects such as wasps (Sphecidae) and dermestids (Dermestidae)were to blame for the remaining 13% damage. These figures indicate that rigid boardinsulation is subject to damage on a wider scope than previously thought.

T

ABLE

1. P

ERCENTAGE

OF

T

REATMENTS

, I

NSPECTIONS

,

AND

T

ECHNICIANS

R

EPORTEDBY

R

ESPONDENTS

AS

A

SSESSED

BY

A

M

AIL

S

URVEY

OF

M

EMBER

C

OMPANIESOF

THE

S

OUTH

C

AROLINA

P

EST

C

ONTROL

A

SSOCIATION

, A

PRIL

1994, N=225.

Treatments Inspections Technicians

Number % Number % Number %

1-50 35 1-50 5 1-5 8251-100 15 51-100 10 6-10 10

101-150 10 101-150 7 11-15 3151-200 5 151-200 10 16-20 0201-250 5 201-250 7 21-25 0251-300 5 251-300 8 >25 5

>300 25 >300 53

T

ABLE

2. P

ERCENTAGE

OF

T

IME

R

IGID

B

OARD

I

NSULATION

D

AMAGE

D

UE

TO

T

ERMITESWAS

S

EEN

(A

BOVE

AND

B

ELOW

G

RADE

)

BY

R

ESPONDENTS

,

BY

F

REQUENCY

OF

O

CCURRENCE

ON

S

TRUCTURES

AS

A

SSESSED

BY

A

M

AIL

S

URVEY

OF

M

EMBER

C

OMPANIES

OF

THE

S

OUTH

C

AROLINA

P

EST

C

ONTROL

A

SSOCIATION

, A

PRIL

1994, N=225.

Above grade Below grade

Occurrence (%) % Time Seen Occurrence (%) % Time Seen

1-10 81 1-10 7211-25 15 11-25 10.526-50 4 26-50 3.551-75 0 51-75 5

>75 0 >75 9

512



Treatment When Rigid Board Insulation Was Present

Pest control companies chose to treat structures with rigid board insulation 57%of the time, compared to 43% that did not. Of those respondents that chose to treattermite-infested structures, 24% treated as they would any other structure and pro-vided a guarantee, 33% treated but did not provide a guarantee, 31% removed insu-lation below grade (Table 7), and another 13% did something other than those optionsmentioned. Responses for the latter category were: trenched out below the insulation,treated, and backfilled; cut insulation off above grade and then sealed bottom of insu-lation; recommended that a qualified builder replace insulation and check structurefor soundness; pretreated but did not know rigid board insulation was later installed;only treated if insulation was not below grade; modified guarantee such that companywas not responsible for damage as a result of hidden entry through insulation; treatedas result of having won a bid.

Of those respondents that chose not to treat structures, 34% believed the insula-tion created a situation conducive to termite infestation that was not treatable or cor-rectable, 45% would not guarantee the treatment (Table 7), another 21% didsomething other than those options mentioned. Responses to the latter category in-

T

ABLE

3. P

ERCENTAGE

OF

T

IME

S

TRUCTURAL

D

AMAGE

D

UE

TO

T

ERMITES

WAS SEEN(ABOVE AND BELOW GRADE) BY RESPONDENTS, BY FREQUENCY OF OCCUR-RENCE ON STRUCTURES AS ASSESSED BY A MAIL SURVEY OF MEMBER COM-PANIES OF THE SOUTH CAROLINA PEST CONTROL ASSOCIATION, APRIL 1994,N=225.

Above grade Below grade

Occurrence (%) % Time Seen Occurrence (%) % Time Seen

1-10 76 1-10 7411-25 12 11-25 1526-50 8 26-50 451-75 0 51-75 0

>75 4 >75 7

TABLE 4. MEAN PERCENT INSULATION AND STRUCTURAL DAMAGE DUE TO TERMITESAS ASSESSED BY A MAIL SURVEY OF MEMBER COMPANIES OF THE SOUTHCAROLINA PEST CONTROL ASSOCIATION, APRIL 1994, N=225.

Insulation Structural

Location % Std. Dev. Location % Std. Dev.

Between grade level and footings

42.25 26.21 Between grade level and footings

9.20 4.82

Just below grade 9.50 5.23 Just below grade 12.75 5.99At grade level 14.85 7.08 At grade level 20.13 18.11Between grade level and sill plate

22.15 20.16 Between grade level and sill plate

28.61 23.65

Above the sill 11.25 6.35 Above the sill 29.31 14.21

Smith & Zungoli: Termites and Rigid Board Insulation 513

cluded: customer did not want insulation removed; gave job to another company; rec-ognized situation due to previous problems experienced with this type of insulation.

Trends show that PCOs generally do one of two things once rigid board is detectedon a structure to be treated or inspected. They remove perimeter insulation just abovethe soil level in order to visually inspect for termites, or refuse to guarantee the treat-ment or treat the structure at all. Since only 25% of companies treated structures asthey would any other, this suggests that the PCO is aware of the problem, but educa-tion is still necessary to inform those who do not know.

This survey will be a vital part of the process to inform PCOs. Although the prob-lem is evident in South Carolina, it is not unique to this state alone. With the ever in-creasing government regulations on new housing, the problem will continue to grow.The real problem lies in the fact that rigid board is placed in a continuous fashionwhich does not allow for a proper inspection to take place. Insulation may be, and usu-ally is, placed in contact with the soil and is often installed below grade, even ex-tended down to the footings of a structure. The Model Energy Code (MEC) states thefollowing for slab-on-grade floors, i.e. monolithic slab construction (abundantlypresent in South Carolina): “In climates below 6,000 annual Fahrenheit heating de-gree days (HDD), the insulation shall extend downward from the top of the slab for aminimum distance of 24 inches . . . greater than 6,000 annual Fahrenheit heating de-gree days (HDD), the insulation shall extend downward from the top of the slab for aminimum of 48 inches . . .” (Council of American Building Officials 1992). The MEC isessentially the standard regulation set and used for most building codes throughout

TABLE 5. CORRELATION BETWEEN PERCENT OCCURRENCE OF INSULATION AND STRUC-TURAL DAMAGE DUE TO TERMITES, USING PEARSON CORRELATION COEFFI-CIENTS (α=0.05) AS ASSESSED BY A MAIL SURVEY OF MEMBER COMPANIES OFTHE SOUTH CAROLINA PEST CONTROL ASSOCIATION, APRIL 1994, N=225.

Location Correlation Coefficient p-Value

Between grade level and footings 0.2443 0.5598Just below grade 0.7657 0.0037At grade level 0.4725 0.1030Between grade level and sill plate 0.6832 0.0006Above the sill plate 0.5244 0.0658

TABLE 6. PERCENT INSULATION DAMAGE DUE TO TERMITES, BY FOUNDATION TYPE ASASSESSED BY A MAIL SURVEY OF MEMBER COMPANIES OF THE SOUTH CARO-LINA PEST CONTROL ASSOCIATION, APRIL 1994, N=225.

Foundation % Std. Dev.

Monolithic slab 33.33 40.81Supported slab 9.17 1.82Floating slab 18.45 19.20Basement 7.05 1.14Crawlspace 22.62 19.49Combination of above types 9.38 2.18


the country. Unfortunately, enforcement of codes in South Carolina is done on a vol-untary basis only, within each county.

In either scenario, the government mandates that rigid board insulation be placed,not only in contact with the soil, but as deep as over 1 m below grade level. We believethat the physical removal of the rigid board just above grade level (a vision strip of 15cm, for example) around the structure’s perimeter would allow the PCO to detect ter-mite activity, including the usual first signs - shelter tubes. Admittedly, there will bea thermal break (thermal short-circuit) caused by removing rigid board insulation,but we feel that the amount of material the termites are removing (the thermal lossas a result of termites) as they damage rigid board and structural members wouldwarrant measures deemed necessary to correct the problem. In addition to the mon-etary loss of rigid board, the monetary loss of structural wooden members certainly isgreat, but precise figures are unavailable. The benefits of removing a small gap of in-sulation above grade level along the perimeter of structures would certainly outweighthe costs in the long run; additional insulation can be placed elsewhere in the struc-ture to recoup the thermal loss from this adjustment.

Some rigid board manufacturers are investigating rigid board with the addition ofinert chemical fillers such as disodium octaborate tetrahydrate (Kramer 1993). Al-though this could potentially exclude termites from damaging the materials, it wouldnot necessarily deter termites from tunneling behind the materials where gaps in con-struction are present. Hidden termite entry once again would ensue.

We believe one final note is rather disturbing. Twelve percent of the respondentshave been involved in litigation due to termite damage hidden by rigid board insula-tion. The majority of these companies, 57%, were from the Coastal region. The com-panies that belong to the SCPCA are among the best in South Carolina. Pastexperience with many of these companies demonstrates they are dedicated to theirwork and, therefore, less likely to be sued. This last number alone serves to illustratethe point that there is, indeed, a problem requiring attention at many levels, not justfrom the PCO.

ACKNOWLEDGMENTS

We wish to express thanks to DowElanco and the South Carolina Pest Control As-sociation for funding this project. The help of all those Pest Control Operators in

TABLE 7. TREATMENT OPTIONS AND THE PERCENTAGE OF COMPANIES THAT CHOSE TOTREAT STRUCTURES VERSUS NOT TREAT STRUCTURES AS ASSESSED BY AMAIL SURVEY OF MEMBER COMPANIES OF THE SOUTH CAROLINA PEST CON-TROL ASSOCIATION, APRIL 1994, N=225.

Chose to treat % Chose not to treat %

Treated as they would any other structure and provided a guarantee

24 Insulation created a situation conducive to termite infestation that was not treatable or correctable

34

Treated but did not provide a guarantee 33

Could not guarantee the treat-ment 45

Removed insulation below grade 31

Did something other than above options 21

Did something other than above options

13

Smith & Zungoli: Termites and Rigid Board Insulation 515

South Carolina who participated in this survey is greatly appreciated. Without theirassistance this work would not have been possible. Gratitude goes out to KevinHathorne and Terry Pizzuto for helping prepare the survey forms to be mailed. In ad-dition, thanks to Dr. Hoke Hill for statistical guidance. This is technical contributionNo. 4006 of the South Carolina Agricultural Experiment Station.

REFERENCES CITED

BECKER, G. 1972. Problems of Testing Materials with Termites, pp. 249-255 in A. H.Walters and E. H. Hueck-Van der Plas [eds]. International BiodeteriorationSymposium. Vol. 2. Applied Science Publishers Ltd., London.

BUILDING RESEARCH STATION (Watford, UK), Overseas Division. 1966. Plastics inBuilding. Overseas Building Notes. 108: 1-9.

CHRISTIAN, J. E. 1990. The Most Needed Building Foundations Research Products,pp. 655-662 in Insulation Materials, Testing, and Applications. American Soci-ety for Testing and Materials, Philadelphia. 759 pp.

COUNCIL OF AMERICAN BUILDING OFFICIALS. 1992. Model Energy Code. Council ofAmerican Building Officials, Leesburg, VA. 87 pp.

GAY, F. J., AND A. H. WETHERLY. 1969. Termite Resistance of Plastics V. CSIRO, Can-berra.

GUYETTE, J. E. 1994. Termites Targeting Foam Insulation. Pest Control. 62: 49, 50,52.

HICKIN, N. E. 1971. Termites: a world problem. Hutchinson & Co. Ltd., London. 233pp.

KRAMER, R. D. 1993. Foam Insulation and Termites. Service Letter ESPC 055075,Number 1284. National Pest Control Association, Inc., Dunn Loring, VA. 3 pp.

SAS INSTITUTE. 1985. SAS User’s Guide: Statistics. SAS Institute, Cary, NC. 956 pp.SHIMIZU, K., Y. NAKASHIMA, AND A. SAKANOSHITA. 1970. Studies on termite damage

to a synthetic high polymer. Bulletin of the Faculty of Agriculture, MiyazakiUniversity. 17: 281-285. Author has only read English summary.

STRZEPEK, W. R. 1990. Overview of Physical Properties of Cellular Thermal Insula-tions, pp. 121-140 in D. L. McElroy and J. F. Kimpflen [eds]. Insulation Mate-rials, Testing, and Applications. American Society for Testing and Materials,Philadelphia. 759 pp.

516



DORMANT SEASON APPLICATION OF

STEINERNEMA CARPOCAPSAE

(RHABDITIDA: STEINERNEMATIDAE) AND

HETERORHABDITIS

SP. (RHABDITIDA: HETERORHABDITIDAE) ON ALMOND FOR CONTROL OF

OVERWINTERING

AMYELOIS TRANSITELLA

AND

ANARSIA LINEATELLA

(LEPIDOPTERA: GELECHIIDAE)

F

ERNANDO

A

GUDELO

-S

ILVA

1

, F

RANK

G. Z

ALOM

2

, A

LBERT

H

OM

1

,

AND

L

ONNIE

H

ENDRICKS

3

Biosys, 1057 E. Meadow CirclePalo Alto, CA 94303

1

Current address: FAS Technologies. 1550 Walnut St. #5. Berkeley, CA 94709

2

Department of Entomology, University of California, Davis, CA 95616

3

Cooperative Extension, 2145 W. Wardrobe, Merced, CA 95340

A

BSTRACT

Overwintering larval populations of

Amyelois transitella

in mummy almonds and

Anarsia lineatella

in hibernaculae on almond trees in California orchards were re-duced by infectives of

Steinernema carpocapsae

strain All and a cold tolerant

Heter-orhabditis

species (HL81) applied in the dormant season. The effect of this reductionin terms of preventing economic damage the following year is not known. Althoughthe population reduction was statistically significant, it is unlikely the increase inmortality achieved would prove to be an economically viable alternative to currentpractices.

Key Words: Almond,


,

Heterorhabditis

sp.,

Amyelois tran-sitella

,

Anarsia lineatella

R

ESUMEN

Poblaciones hibernanates de larvas de


en “almendras momi-ficadas”, y poblaciones hibernanates de

Anarsia lineatella

en hibernáculos de almen-dros en California, fueron reducidas por estados infectivos de dos nemátodosentomopatógenos,


cepa A11 y una especie de

Heterorhabdi-tis

(HL81) tolerante a baja temperatura, aplicados durante la estación de dormancia.El efecto de esta reducción en términos de impedir el daño económico al año siguienteno es bien conocido. Aunque la reducción en la población fue estadísticamente signifi-cativa, se duda que el incremento de la mortalidad logrado sea una alternativa econó-

micamente viable para las prácticas presentes de control de estos insectos.

For many years entomopathogenic nematodes have been considered to be poten-tial biological mortality agents of many economically important insect pests (Glaseret al. 1940, Forschler & Gardner 1991). Because their nutritional requirements werepoorly known for some time, it was difficult to produce adequate quantities of thenematodes for large scale field experiments. Advances in the understanding of the nu-tritional requirements (Buecher & Hansen 1977) and the development of mass cultur-

Agudelo et al.: Nematodes Reduce Overwintering Larvae

517

ing of these nematodes (Bedding 1984, Friedman 1990) has made it possible toproduce them in sufficient quantities for such research. Increased emphasis on fieldevaluation will result in a better assessment of these organisms as practical insectcontrol agents.

Entomopathogenic nematodes appear to have the greatest potential as controlagents when directed against immature insect stages occupying secluded niches, suchas split almonds (Lindegren et al. 1978, Lindegren et al. 1987), tree galleries (Deseo& Miller 1985), leaf sheaths and cabbage heads (Welch & Briand 1961), artichokecrowns (Bari & Kaya 1984) and soil crevices (Wright et al. 1987, Shanks & Agudelo-Silva 1990).

The use of entomopathogenic nematodes against two important insect pests of

Prunus

spp. that occupy secluded niches as larvae during the dormant season has notbeen fully investigated. These insects are the navel orangeworm,


(Walker) (Lepidoptera: Pyralidae) and the peach twig borer

Anarsia lineatella

(Zeller), Lepidoptera: Gelechiidae). Together, these insects are estimated to cause di-rect damage of more than $10 million to California almond growers alone, not includ-ing treatment costs.

A. transitella

is the most important insect pest of almonds.Larvae of the two summer generations invade the nuts following almond hullsplit andfeed on the meats. Almond handlers accept up to 4% damaged meats, but they awardbonuses to growers for lower damage (Klonsky et al. 1990). After harvest, some nutsremain on the trees where they continue to attract

A. transitella.

The nuts dry and be-come “mummies” which often contain

A. transitella

larvae. The overwintering larvaebecome adults the following spring and are the source of infestation in the subsequentcrop. Cultural controls for

A. transitella

include removing these “mummies” and har-vesting nuts as early as possible in the season to avoid peak moth flights. Almost halfof the orchards (~ 100,000 hectares in a given year) receive in-season organophos-phate sprays for

A. transitella

(Klonsky et al. 1990). While there is evidence that ap-plications of beneficial nematodes can infect

A. transitella

larvae infesting newly splitalmonds in the summer (Agudelo-Silva et al. 1987, Lindegren et al. 1978), control ofoverwintering larvae has not been attempted.

A. lineatella

is an important pest of both almonds and stone fruits in California,and the summer generation causes direct damage to nut meats and fruit. The larvaeoverwinter in hibernaculae built in crotches of twigs or branches. The larvae emergeduring February and early March (Zalom et al. 1992) and migrate to shoot tips wherethey burrow into them to complete larval development. Organophosphate insecticidesare routinely applied during the dormant season for

A. lineatella

control, and thesehave been the recommended alternative to disruptive in-season sprays (Rice & Jones1988, Zalom et al. 1991). Recently, the application of dormant sprays has been criti-cized for environmental reasons, and suitable alternatives or mitigating measures arebeing pursued (e.g. Barnett et al. 1993).

Here we report results of field and laboratory trials employing two entomopatho-genic nematode species applied during the dormant season for control of overwinter-ing

A. transitella

and

A. lineatella

larvae in almonds. In the dormant season, pestpopulations are relatively low and concentrated on the mummy nuts.

M

ATERIALS

AND

M

ETHODS

Nematodes


strain All and a cold tolerant

Heterorhabditis

species(currently undescribed but hereafter referred to as HL81) were used in the tests.

Het-

518



erorhabditis

sp. strain HL81 (isolated in Holland) has been shown to parasitize in-sects at 9

°

C and 10

°

C(Simmons & Van der Schaf 1986). Infective juveniles wereartificially mass reared (Biosys unpubl.) and stored in moist foam at 5

°

C(

S. carpocap-sae

strain All) or at 10

°

C (

Heterorhabditis

sp. strain HL81) until needed. Before eachtest application, the nematodes were examined under magnification to confirm thatthe survival rate was greater than 90 percent.

A. transitella

Trials were conducted in Le Grand, Merced Co., and Durham, Butte Co., Califor-nia, during February and December, 1987, respectively. The Le Grand test was con-ducted to determine if mortality of overwintering

A. transitella

could be increased bya dormant season

S. carpocapsae

application. Treatments were applied on 23 Febru-ary in a commercial orchard (var. “Le Grand”) which was heavily infested with over-wintering

A. transitella

(33% mummies infested in a pretreatment survey). Sixapplications rates were evaluated: 0, 0.62, 1.25, 2.5, 5.0, 10.0 and 20.0 million nema-todes per tree. Each experimental unit was a tree and treatments were replicated fivetimes. A totally randomized design was utilized. Nematodes were applied suspendedin water (10.2 liters per tree) using a handgun sprayer (Farmtec, Oakland, CA). Im-mediately after the application, ten mummies were collected from each treated tree,individually placed in plastic bags in ice chests containing ice and brought to the lab-oratory to be examined for presence of nematodes. Nematodes were extracted fromthe mummies following the method developed by Lindegren et al (1987). One week af-ter the application, 50 mummies were randomly collected from each of the trees,cracked by hand and examined for

A. transitella.

Larvae were tallied as alive or dead.Dead larvae were dissected and examined microscopically to ascertain the presence ofnematodes.

The Durham test was conducted to compare the ability of both

S. carpocapsae

and

Heterorhabditis

sp. strain HL81 to infect

A. transitella

larvae at low temperature.Treatments were applied on 15 December in a commercial orchard (var. “Nonpareil”,“Ne-plus Ultra,” and “Mission”; 2:1:1 ratio).

Treatments evaluated were

S. carpocapsae

,

Heterorhabditis

sp. strain HL81 andan untreated control. Mummies were randomly collected from trees and divided into15 groups each of which contained 130 mummies. The mummies were treated suchthat there was certainty that all the mummies received nematodes. Each group ofmummies was placed on a piece of flat plastic laid out on the ground in the orchardand then arranged on the plastic touching each other, in a single layer. Twenty mum-mies were taken from each group and examined for

A. transitella

larvae. Infestationof these mummies was determined to be 59.5%. Each group was randomly assignedto one of the treatments, and the treatments were applied using a previously cali-brated spray bottle. Nematodes were suspended in water and applied at a rate of 51nematodes per mummy (5,610 nematodes suspended in water were applied to eachgroup of 110 mummies). Immediately after application (less than 15 minutes), 10mummies were randomly chosen from each group, placed in individual bags andstored in ice chests to be brought to the laboratory for nematode extraction (Lindegrenet al.1987). The remaining mummies (100 per group) were placed in individual wiremesh cages which were suspended from branches in the almond trees. Two weeks af-ter the nematodes were applied, the mummies from each of the cages were placed inice chests containing ice and brought to the laboratory to be examined for the presenceof larvae. Dead larvae were dissected and microscopically examined to ascertain thepresence of nematodes.

Agudelo et al.: Nematodes Reduce Overwintering Larvae

519

A. lineatella

This trial was conducted in a 2-year old non-bearing almond orchard near Cortez,Merced Co., CA, to determine the efficacy of a dormant application of

S. carpocapsae

on overwintering

A. lineatella

larvae. The treated trees were infested with

A. lin-eatella

overwintering in hibernaculae on the branches. Treatments were applied on20 January, 1987, with a handgun sprayer and consisted of

S. carpocapsae

at rates of1.5 and 0.5 million per tree, a conventional dormant spray of diazinon (16.0 liter AIper ha) plus Volck supreme oil (48 liter per ha) and water as a check. The treatmentswere arranged in a completely randomized design with twenty single tree replicates,each treatment tree separated by an untreated buffer tree. No attempt was made tofind dead larvae after the application of the treatments because the hibernacula aredifficult to locate in significant quantity. It was decided that a more appropriate andpractical evaluation of the efficacy of the nematodes on mortality of overwintering lar-vae would be to count shoot strikes in the spring. Shoot strikes can be used to evaluatelarval control (Summers et al. 1959) since the overwintering larvae which emergefeed on new shoots and bore into them. The shoot tips which are killed as a result ofthis feeding are relatively easy to identify. Efficacy was evaluated on 27 March, 1987,by counting the total number of shoot tips damaged by

A. lineatella

larvae followingemergence of the overwintering generation.

R

ESULTS

AND

D

ISCUSSION

A. transitella

In the Le Grand test, the nematode applications did not statistically increase themortality of overwintering larvae. Although the mortality rates for the 0.62 and 5.00million

S. carpocapsae

per tree application seemed to increase larval mortality, therewas no significant difference (

P

>0.05) between rates of application when data weresubjected to one way ANOVA. Similarly, no significant relationship was found (r

2

=0.014;

P

>0.05) when dose/mortality data was subjected to regression analysis (Table1). No nematodes were found inside the dead larvae.

There was a positive relationship between the number of nematodes applied pertree and the number of nematodes delivered to the mummies (

F

= 66.3;

P

<0.001) (Ta-ble 1). The relationship between dose rate and number of mummies receiving nema-todes was also positive, indicating that better coverage is obtained as the number ofnematodes applied per tree increases. These are important results as they demon-strate the potential of delivering nematodes to the site of the overwintering larvae us-ing commercially available spraying equipment. Larval mortality in this trial wasmuch lower than that observed by Lindegren et al. (1978) and Agudelo-Silva et al.(1987) for applications at hullsplit (summer). Since there were no differences in larvalmortality between treatments and no nematodes were found in the dead larvae, mostlikely the nematodes did not infect

them. This may have been due, at least partially,to temperatures during the time of this study. The low temperature probably im-paired the ability of nematodes to move on the mummies to reach the larvae insidethem, and even if a nematode ultimately reached a larva, the nematode and its sym-biont bacteria may not have lethally infected it. Although the temperature reached18

°

C for a few hours during the seven days after the nematodes were applied, most ofthe time the temperature was too low to allow nematode infection and development.During the seven days following application, temperatures did not exceed 10

°

C for60% of the time (101 hours). Development and reproduction of

S. carpocapsae

occursbetween 15

°

C and 27

°

C (Kaya 1977).

520



In the Durham test, it was hoped that Heterorhabditis sp. strain HL81, a cold-tol-erant nematode strain, would prove more effective controlling A. transitella during itsoverwintering period. Acceptable delivery of nematodes to the mummies was achievedwith 90 and 82 percent of the mummies for S. carpocapsae and Heterorhabditis sp. re-spectively releasing nematodes in the extraction procedure (Lindegren et al 1987)(Table 2). Larval mortality of mummies treated with either nematode species was 2.2times greater than mortality observed from mummies that were treated with wateralone. This difference was statistically significant (P < 0.05) and strongly suggeststhat applications of entomopathogenic nematodes can increase mortality of A. tran-sitella in mummy nuts during the dormant period. No nematodes were found insidethe dead larvae. Unfortunately, A. transitella survival was over 88% for both species.The fact that no nematodes were found inside the dead larvae is not surprising. Mostlikely, nematodes invaded larvae and killed them, but died before completing devel-opment and reproduction. The dead nematodes would have decayed before the deadA. transitella larvae were examined. The periods of low temperatures (< 10°C) that oc-curred between the time of nematode application and the time when the dead larvaewere examined (when infection by the nematodes must have taken place) were notconducive for the successful growth of the nematode and its symbiont bacteria (Moly-neux, 1986). Successful growth of the symbiont is necessary for nematode develop-ment (Poinar & Thomas 1966). The significant (P < 0.05) difference between thenumber of dead larvae found in the mummies treated with nematodes and thosetreated with water only suggests that the nematodes were responsible for the differ-ences in mortality. It is possible that more cold-tolerant nematode strains than thoseapplied in this trial could increase mortality. However, the increase in larval mortal-ity that could be attributed to nematode treatment in this trial, although importantbiologically, would not be acceptable relative to conventional controls currently avail-able.

A. lineatella

The difference in the number of A. lineatella shoot strikes between the almondtrees treated with S. carpocapsae during the dormant season (Table 3) at 1.5 million

TABLE 1. STEINERNEMA CARPOCAPSAE STRAIN ALL JUVENILES APPLIED TO ALMONDTREES FOR CONTROL OF AMYELOIS TRANSITELLA LARVAE OVERWINTERING INMUMMY ALMONDS, 23 FEBRUARY, 1987 (N=5). LARVAL MORTALITY WASEVALUATED 7 DAYS AFTER TREATMENT.

Mean Nematodes2 per Mummy % Mummies

with Nematodes

Number Nematodes per Tree (× 106)

Percent Dead1 Larvae ± SD

Mean %2

± SD Range

0.00 2.5 (3.6) 0.0 (0) — 00.62 4.0 (6.9) 27.6 (35.6) 12-108 601.25 2.7 (6.0) 21.2 (10.0) 12-84 602.50 2.6 (2.4) 13.2 (9.8) 12-204 485.00 5.3 (11.0) 55.6 (34.6) 12-204 88

10.00 2.0 (2.6) 68.6 (61.1) 12-240 8820.00 2.1 (3.3) 232.0 (138.0) 36-588 92

1Fifty mummy nuts per replicate.2Ten mummy nuts per replicate.

Agudelo et al.: Nematodes Reduce Overwintering Larvae 521

and 0.5 million per tree and the water check was statistically significant (P < 0.05),but the level of control (about 24%) was significantly (P < 0.05) less than the level ofcontrol (~ 93%) achieved with the conventional dormant treatment of diazinon plus oil(F=34.27; DF=3). These data suggest that S. carpocapsae have the ability to penetrateoverwintering hibernaculae, and kill the larvae inside them, reducing their overwin-tering populations and subsequent damage to trees. The hourly temperatures duringsome of the days when the larvae were exposed to the nematodes was high enough (upto 23°C) to allow nematode movement and larval infection. However, the temperaturewas low (less than 10°C) for several days during the time that the larvae were exposedto the nematodes, and this probably diminished the growth, reproduction and infec-tivity of the nematodes (Kaya 1977).

The ability of entomopathogenic nematodes to reduce overwintering A. transitellaand A. lineatella larval populations is not resolved. It was promising that larval mor-tality was achieved with existing nematode strains under field conditions, and thatnematodes could be reliably delivered to mummy almonds in commercial orchards (LeGrand test). However, larval mortality of both A. transitella and A. lineatella, whilesignificant, was below commercially acceptable levels. It can be hypothesized that ifa nematode were available that would remain active in larvae for extended periods oftime below 10°C, the level of larval mortality achieved could be greater. It is also pos-

TABLE 2. STEINERNEMA CARPOCAPSAE STRAIN ALL AND HETERORHABDITIS SP. STRAIN(HL81) APPLIED AT A RATE OF 51 NEMATODES PER MUMMY FOR CONTROL OFAMYELOIS TRANSITELLA LARVAE, 15 DECEMBER 1987.

Treatment

Mean Percent Dead Larvae1

± SD

Mean Number Nematodes2 per Mummy

± SD

Range of Nematodes per

Mummy

Percent Mummies with

Nematodes

S. carpocapsae 11.8 (4.1)a 26.8 (25.02) 0-126 90Heterorhabditis sp. 11.6 (2.3)a 19.4 (26.51) 0-162 82Water check 3.6 (2.8)b 0.0 (0.0) — —

1Means followed by the same letter are not significantly different (P > 0.05) when compared by Duncan’s(1995) new multiple range test.

2Ten mummies nuts per replicate.

TABLE 3. STEINERNEMA CARPOCAPSAE STRAIN ALL APPLIED TO ALMOND TREES FORCONTROL OF ANARSIA LINEATELLA LARVAE OVERWINTERING IN HIBERNACU-LAE, 20 JANUARY, 1987 (N=20). EFFECT OF TREATMENTS MEASURED ASNUMBER OF LARVAL SHOOT STRIKES PER TREE ON 27 MARCH, 1987.

Treatment RateMean Shoot Strikes1

per Tree ± SD

Water check — 28.4 (10.6)cS. carpocapsae 1.5 × 107 per tree 22.3 (9.8)bS. carpocapsae 0.5 × 107 per tree 21.4 (9.3)bDiazinon + oil 16 l AI per Ha. + 48 L per Ha. 2.1 (2.7)a

1Means followed by the same letter are not significantly different (P > 0.05) when compared by Duncan’s(1955) new multiple range test.


sible that treatments for both pest species could be delayed until early bloom whenwarmer conditions would be present, yet overwintering larvae would be present asthe treatment target. In order to be commercially feasible at present, the level of lar-val mortality achieved must reach 70 to 90 percent, and the cost of applying nema-todes must be competitive with conventional in-season sprays for A. transitella, ($24to $40 per ha) and dormant sprays for A. lineatella ($20 to $56 per ha).

ACKNOWLEDGMENTS

This study was supported in part by the Almond Board of California. We especiallythank B. Wilk and C. Kitayama, Scientific Methods, Chico, CA, for their valuable as-sistance.

REFERENCES CITED

AGUDELO-SILVA, F., J. E. LINDEGREN, AND K. A. VALERO. 1987. Persistence of Neoa-plectana carpocapsae (Kapow selection) infectives in almonds under field con-ditions. Florida Entomol. 70: 288-291.

BARI, M. A., AND H. H. KAYA. 1984. Evaluation of the entomogenous nematode Neoa-plectana carpocapsae (Steinernema feltiae) Weiser (Rhabditida: Steinernema-tidae) and the bacterium Bacillus thuringiensis for suppression of theartichoke plume moth (Lepidoptera: Pterophoridae). J. Econ. Entomol. 1: 225-229.

BARNETT, W. W., J. EDSTROM, R. COVIELLO, AND F. G. ZALOM. 1993. Insect pathogen“Bt” controls peach twig borer on fruits and almonds. Calif. Agric, 47: 4-6.

BEDDING, R. A. 1984. Large scale production, storage and transport of insect-parasiticnematodes Neoaplectana spp, and Heterorhabditis spp. Ann. Appl. Biol. 101:117-120.

BUECHER, E. J., AND E. L. HANSEN. 1977. Mass culture of axenic nematodes using con-tinuous aeration. J. Nematol. 3: 199-200.

DESEO, K. V., AND L. A. MILLER. 1985 Efficacy of entomogeneous nematodes Stein-ernema spp., against clearwing moths, Synanthedon spp., in north Italian ap-ple orchards. Nematologica 31: 100-108.

DUNCAN, D. B. 1955. Multiple range and multiple F tests. Biometrics 11: 4-6.FORSCHLER, B. T., AND W. A. GARDNER. 1991. Field efficacy and persistence of ento-

mopathogenic nematodes in the management of white grubs (Coleoptera: Scar-abeidae) in turf and pasture. J. Econ. Entomol. 84: 1454-1459.

FRIEDMAN, M. J. 1990. Commercial production and development, pp. 153-172 in R.Gaugler and H. K. Kaya, [eds.] Entomopathogenic Nematodes in BiologicalControl. CRC Press. Boca Raton.

GLASER, R. W., E. E. MCCOY, AND H. B. GIRTH. 1940. The biology and economic im-portance of a nematode parasitic in insects. J. Parasitol. 26: 479-495.

KAYA, H. K. 1977. Development of the DD-136 strain of Neoaplectana carpocapsae atconstant temperatures. J. Nematol. 9: 346-349.

KLONSKY, K., F. G. ZALOM, AND W. W. BARNET. 1990. California’s almond IPM pro-gram. California Agric. 44: 21-24.

LINDEGREN, J. E., C. E. CURTIS, AND G. O. POINAR. 1978. Parasitic nematodes seeksout navel orangeworm in almond orchards. California Agric. 32: 10-11.

LINDEGREN, J. E., F. AGUDELO-SILVA, K. A. VALERO, AND C. E. CURTIS. 1987. Compar-ative small-scale field application of Steinernema feltiae for navel orangewormcontrol. J. Nematol. 19: 503-504.

MOLYNEAUX, A. S. 1986. Heterorhabditis spp. and Steinernema (=Neoaplectana) spp:temperature, and aspects of behavior and infectivity. Exp. Parasitol. 62: 169-180.

POINAR, G. O. JR., AND G. M. THOMAS. 1966. Significance of Achromobacter nemato-philus Poinar and Thomas (Achromobacter nematophilus: Eubacteriales) in the

Agudelo et al.: Nematodes Reduce Overwintering Larvae 523

development of the nematode DD-136 (Neoaplectana: Steinernematidae). Par-asitol. 56: 385-390.

RICE, R. E., AND R. A. JONES. 1988. Timing post-bloom sprays for peach twig borer(Lepidoptera; Gelechiidae) and San Jose Scale (Homoptera: Diaspididae). J.Econ. Entomol. 81: 293-299.

SHANKS, C. H., AND F. AGUDELO-SILVA. 1990. Field pathogenicity of heterorhabiditidand steinernematid nematodes (Nematoda) infecting black vine weevil larvae(Coleoptera: Curculionidae) in cranberry bogs. J. Econ. Entomol. 83: 107-110.

SIMMONS, W. R., AND D. A. VAN DER SCHAFF. 1986. Infectivity of three Heterorhabdi-tis isolates for Otiorhynchus sulcatus at different temperatures, pp. 285-289 inR. A. Samson, J. M. Vlak, and D. Peters., [eds.]. Fundamental and Applied As-pects of Invertebrate Pathology Foundation of the Fourth International Collo-quium of Invertebrate Pathology. Wageningen. The Netherlands.

SUMMERS, F. M., W. W. BARNETT, R. E. RICE, AND C. V. WEEKLY. 1995. Control of peartwig borer on almonds and peaches. California J. Econ. Entomol. 52(4): 637-639.

WELCH, H. E., AND L. J. BRIAND. 1961. Field experiment on the use of a nematode forthe control of vegetable crop insects. Proc. Entomol. Soc. Ontario. 91: 197-202.

WRIGHT, R. J., F. AGUDELO-SILVA, AND R. GEORGIS. 1987. Soil applications of Stein-ermatid and Heterorhabditid nematodes for control of Colorado Potato Beetle,Leptinotarsa decemlineata (Say). J. Nematol. 19: 201-206.

ZALOM, F. G., R. A. VAN STEENWYK, W. J. BENTLEY, R. COVIELLO, R. E. RICE, W. W.BARNETT, M. A. HOY, M. M. BARNES, B. A. TEVIOTDALE, W. D. GUBLER, AND M.V. MCKENRY. 1991. Almond pest management guidelines. Univ. of CaliforniaPMG Publ. 1. 22pp.

ZALOM, F. G., W. W. BARNETT, R. E. RICE, AND C. V. WEAKLEY. 1992. Factors associ-ated with flight patterns of the peach twig borer (Lepidoptera: Geleichiidae) ob-

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

served using pheromone traps. J. Econ. Entomol. 85: 1904-1909.

Landolt: Sweet Baits for Moths

523

ATTRACTION OF

MOCIS LATIPES

(LEPIDOPTERA: NOCTUIDAE) TO SWEET BAITS IN TRAPS

P

ETER

J. L

ANDOLT

U.S. Department of Agriculture, Agricultural Research ServiceP. O. Box 14565, Gainesville, FL 32604

A

BSTRACT

Traps baited with solutions of molasses or jaggery (unrefined palm sugar) cap-tured significant numbers of the moth

Mocis latipes

Guenee, indicating their attrac-tion to these baits. Numbers of moths captured were affected by bait concentrationand by bait age. Greatest moth captures were obtained with 20% molasses in wateror 5, 10 or 20% jaggery in water. Molasses and jaggery baits aged in the laboratory forup to three days before field testing were more attractive than freshly-made baits to

M. latipes

moths. This is the first demonstration of

M. latipes

attraction to sugar-based baits.

Key Words: Insecta, moth, sugar, fermentation, microbial.

R

ESUMEN

Las trampas cebadas con soluciones de melaza o azúcar de palma sin refinar cap-turaron cantidades significativas de la polilla

Mocis latipes

Guenee, indicando su

524



atracción a esos cebos. El número de polillas capturadas fue afectado por la concen-tración y edad de los cebos. Las mayores capturas de polillas fueron obtenidas con 20%de melaza en agua o 5, 10 o 20% de miel de palma en agua. La atraccción de los cebosde melaza y azúcar de plama envejecidos hasta tres días en el laboratorio antes deprobarse en el campo fue superior a la de los cebos frescos. Esta es la primera demos-

tración de la atracción de

M. latipes

hacia cebos azucarados.

Sweet baits (sugar baits) are widely used by collectors of Lepidoptera to attractmoths and butterflies to a site where they may be captured, as described by Holland(1903) and Sargent (1976). The ingredients used, although varied, include sugar andsugar-rich materials, as well as fruits and alcoholic beverages. It is thought that theattraction and feeding of moths on artificial sugar baits is an indication of their ten-dency to feed on natural sources of sugars, such as rotting fruit, tree sap, insect hon-eydew, and flower nectars (Norris 1935). There are no direct comparisons offormulations of sweet baits or quantitative efforts to optimize bait effectiveness in at-tracting Lepidoptera, although they appear to lure a wide variety of moths and but-terflies (Sargent 1976, Norris 1935 and references therein).

Although such baits are commonly used to collect non-pest species for taxonomicor life history studies, there are indications that they may be attractive to a numberof important pest species of moths as well. Norris (1935) supposed that nearly all tem-perate Noctuidae (which would necessarily include many pests) are attracted to sugarbaits, but did not provide documentation. Frost (1928) captured over 48,000 noctuidmoths in 2 years of trapping tests in peach orchards using pails of molasses or syrupsolutions, but did not report the identity of the species captured. Molasses and sugarsyrups have been evaluated as a bait placed in pails for attracting and killing

Gra-pholita molesta

(Busck) moths (Frost 1926, 1928, 1929) and

Cydia pomonella

(L.)moths (Eyer 1931) in fruit orchards. Similarly, pans of poisoned vinegar and molasseswere reported to attract and kill corn earworm moths,

Helicoverpa zea

(Boddie)(Glover 1855, cited by Ditman & Cory 1933). Similar materials may provide sourcesof useful attractants for monitoring pest populations or developing attracticidal ap-proaches to suppress pest populations.

This paper reports the attractiveness of selected sweet baits to the moth,

Mocislatipes

Guenee, as evidenced by captures in baited traps. This species is a pest of pas-ture grasses, sugar cane and sorghum in the southeastern United States andthroughout much of the neotropics (Ogunwolo & Habeck 1975; Strayer 1975). A sexpheromone of this species is known to be attractive to males (Landolt & Heath 1989),but there are no methods available for attracting and trapping females. Their attrac-tion to any type of food bait would provide the basis for development of new lures formonitoring and, possibly, controlling populations. The capture of

M.

latipes

in trapscontaining sweetened food baits was previously observed by the author in tests toevaluate food-baited traps for tephritid fruit flies. The objectives of this study were todetermine if these moths could be trapped with sugar-based baits and to evaluate ef-fects of concentration and age of selected baits on numbers of moths attracted to, andcaptured in, a trap. Information on the optimum concentration and age of baits wouldbe of use in future attempts to isolate and identify attractive volatile chemicals ema-nating from these baits. Also, the determination of an aging effect on bait effective-ness would support previous suggestions that microbial fermentation is a crucialcomponent of a good sugar bait for Lepidoptera (Norris 1935).


525

M

ATERIALS

AND

M

ETHODS

Four trapping tests were conducted to determine the attractiveness of baits to themoths. These were: 1) an initial comparison of moths captured with five differentbaits and a control, 2) an assessment of the effect of bait age on moths captured, 3) anassessment of effects of bait concentration on moths captured, and 4) a comparison ofthe effectiveness of two optimized baits.

Invaginated glass traps, McPhail traps (Newell 1936), were used for all tests.These traps, used for monitoring tephritid fruit flies, hold about 0.5 liters of fluid andhave a 6.5-cm diam opening at the trap bottom for insects to enter. All baits were pre-pared and tested in 200-ml amounts per trap. Traps were hung on wire loops mountedon wooden stakes, with the trap opening above the tallest vegetation (about 50 cmabove ground). This height was selected because

M. latipes

were observed in earlymorning flight just above the top of the grass and weed canopy. A randomized com-plete block design was used for the first three tests, with blocks at four different sites.Traps within a block were 10 m apart. All trap sites were in grassy areas along roadsbordering crops (soybean, peanut, sugar cane).

All captured moths were sorted and counted by species and sex. However, numberssuitable for statistical analyses were obtained only for

M. latipes

, which was abun-dant throughout the study period (late August to early November).

Bait Comparison

The first experiment was a preliminary comparison of six different baits. Thetreatment selection was based partly on unreported observations of moths in trapsbaited for fruit flies. Bait treatments were 1) de-ionized water; 2) fruit pectin (Sure-Jell, Kraft General Foods, White Plains, NY); 3) brewer’s yeast (Fleischmann’s Yeast,Specialty Brands, San Francisco, CA) and sucrose; 4) honey; 5) molasses (Grandma’sMolasses, Mott’s, Stanford, CT); and 6) jaggery or unrefined palm sugar (Indian Kol-hapur Jaggery, House of Spices Inc., Jackson Heights, N.Y.). For treatments 2, 4, 5,and 6, 20 g of material was mixed in 200 ml of de-ionized water for each trap. Fortreatment three, 20 g of purified cane sugar and one packet of yeast (7 g) were addedto 200 ml of water. All baits were prepared on the day they were used in traps. The ex-periment was set up as three blocks of traps; each block was a set of 6 traps, each trapcontaining a different bait. Traps were removed, emptied, cleaned, and rebaited after48 hours. A total of 11 block replicates was accumulated over four different baitingtimes (three block sites by four baiting times, with one block missing due to trapbreakage). Treatments were randomized within each block.

Effects of Bait Age

The second experiment compared numbers of moths trapped in either 10% molas-ses or 10% jaggery solutions that were aged before placement in the field. Multiplebait samples (200 ml) were prepared 1, 2, 3, 5, and 7 days before the beginning of atrapping experiment and held in the laboratory at 23

±

0.5

°

C in 500-ml glass jars cov-ered with paper toweling. Bait solutions were placed in traps in the field in the after-noon, and traps were removed and emptied the following morning. Trap blockscontaining one of each of the five treatment ages were set up at four different sites.For each material this test was conducted on three different days providing 12 blockreplicates. Treatments were randomized within all blocks.

526



Effects of Dose

The third experiment compared the effectiveness of a range of concentrations ofmolasses and jaggery on numbers of moths captured. Concentrations tested were 2.5,5, 10, 20, and 40 g of material per 200 ml water per trap. Baits were prepared two daysbefore traps were baited and stored in the laboratory in 500-ml glass jars covered withpaper toweling. Traps were baited in the afternoon and removed and emptied the fol-lowing morning. Trap blocks (five traps) were set up at four sites, with all treatmentsrandomized within each block. For each material, treatment comparisons were con-ducted on three different days, providing 12 block replicates (four sites, three days).

Comparison of Molasses and Jaggery

Solutions of 10% jaggery and 20% molasses were compared directly in McPhailtraps for their effectiveness as trap baits for

M. latipes

. Baits were prepared two daysbefore placement in traps in the field and stored in the laboratory in 500-ml glass jarscovered with paper toweling. Traps containing either molasses or jaggery were pairedin the field; 10 pairs of traps were placed in fields in the afternoon, then removed andemptied the following morning.

Data for captures of

M. latipes

in traps from multiple comparison tests (tests 1, 2,and 3) were transformed to percentages of block totals, followed by an arcsin trans-formation before using an analysis of variance (Steel & Torrie 1960). Significance ofdifferences between treatment mean percents was determined using Duncan’s multi-ple range test (Duncan 1955) when ANOVA indicated a significant F value. Means for

Figure 1. Mean (± SE) percentages of block totals for numbers of M. latipes mothscaptured in traps baited with water, or aqueous solutions of 10% pectin, 10% honey,10% sugar and 3.5% brewer’s yeast, 10% molasses, or 10% jaggery, with a total of 177trapped. Percentages sharing the same letter are not significantly different by Dun-can’s multiple range test at P > 0.05.


527

data comparing the attractiveness of molasses and jaggery were tested for signifi-cance, without transformation, using Student’s t-test (Steel & Torrie 1960). For all ex-periments, numbers of males and females of

M. latipes

were pooled for analyses ofdata.

R

ESULTS

In the first experiment comparing different bait materials, significantly more

M.latipes

were captured in traps baited with yeast and sugar, molasses, and jaggerythan in traps baited with water, pectin, or honey (F = 12.5, df = 5,60,

P

< 10

-6

) (Fig. 1).Greatest numbers of

M. latipes

were captured in traps containing 10% jaggery, com-pared to molasses, yeast and sugar, honey, pectin, or water. (Fig. 1). Significantlymore moths were caught in traps baited with 10% molasses than were captured intraps baited with honey (Fig. 1). None were captured in traps baited with pectin orwith water.

For the second experiment, bait age significantly affected captures of

M. latipes

intraps baited with either molasses (F = 3.3, df = 4,45,

P

=0.02) or jaggery (F = 2.6, df =4,49,

P

= 0.05) (Fig. 2). Captures increased significantly from the first to the third daywith both materials, followed by an apparent reduction in effectiveness, although thiswas not significant at the 5% level.

In the third experiment, captures of

M. latipes

in traps baited with either molassesor jaggery were significantly affected by concentration (F = 15.1, df = 4,50,

P

< 10

-6

)

Figure 2. Mean (± SE) percentages of block totals for numbers of M. latipes mothscaptured in traps baited with either 10% molasses (open bars) or 10% jaggery (cross-hatched bars) baits of different ages, with a total of 653 trapped. Within a bait cate-gory (molasses or jaggery) mean percentages sharing a letter are not significantlydifferent by Duncan’s multiple range test at P > 0.05.

528



(Fig. 3). Numbers of moths in traps baited with molasses increased with each increasein concentration and were significantly greater at 20% compared to all other testedconcentrations. Numbers of moths captured in traps baited with jaggery increasedwith concentration increases up to 5%, with no differences in trap catches among 5,10, and 20% solutions.

For the fourth experiment, traps baited with three-day-old 10% jaggery capturedsignificantly more

M. latipes

than traps baited with three-day-old 20% molasses (t =2.97, df = 9,

P

= 0.016, Mean

±

SE = 8.8

±

1.9 moths per trap per night for jaggery;Mean

±

SE = 4.9

±

1.1 moths per trap per night for molasses).Although counts of male and female

M. latipes

were combined for statistical com-parisons of bait effectiveness, total numbers of males and females trapped indicate anoverall sex ratio of near 1 to 1, with 1,027 males and 1,039 females captured in all ofthe traps baited with molasses and jaggery for experiments 1 through 4 (Table 1).Other pest species of noctuids were also captured in these tests (Table 1), including

Anticarsia gemmatalis

Hubner,

Mocis disseverans

(Walker), and

Spodoptera fru-giperda

(J. E. Smith). Three species of

Leucania

were captured but were not deter-mined to species.

D

ISCUSSION

The results of the experiments reported here clearly indicate that

M. latipes

areattracted to sweet baits, such as molasses or jaggery. Because the sex ratio for

M. lati-pes

trapped with either of these two materials was near one to one, it may be possible

Figure 3. Mean (± SE) percentages of block totals for numbers of M. latipes mothscaptured in traps baited with either three-day-old dosages of molasses or three-day-old dosages of jaggery, with a total of 1,099 trapped. Within a bait category, mean per-centages that share a letter are not significantly different by Duncan’s multiple rangetest at P > 0.05.


529

to develop attractants based on these baits for monitoring the presence of females inaffected crops.

Numbers of other noctuid moths trapped indicate there is some potential to de-velop attractants and traps based on sugar baits for other pest species of moths. It isassumed that the relatively lower numbers of these other species trapped is due totheir lower populations at sites during the trapping period. The location and seasonfor these tests were selected to maximize the presence of

M. latipes

. It will be neces-sary to target populations of other pest species of interest to more adequately deter-mine their attraction to these baits.

It is unclear what ingredients are critical to the attractiveness of the tested baitsfor this and other species of moths. Sargent (1976) indicated that sugar (presumablysucrose, as refined cane or beet sugar) is the only ingredient required, but also addedbeer to attract

Catocala

species. Holland (1903) described using a solution of sugar,beer and rum. Experiments to control species of

Grapholita

and

H. zea

were con-ducted with molasses or sugar syrups (Frost 1926, 1928, 1929; Eyer 1931; Ditman &Cory 1933). The results reported here for

M. latipes

indicate that honey or a mixtureof sugar and brewer’s yeast were not effective and that additional factors present inthe molasses and jaggery are important to moth attraction.

Attractiveness of sugar baits to moths is assumed to be due in part to fermentationprocesses necessary for the production of attractants (Norris 1935; Sargent 1976).This assumption is supported here by the significant increase in numbers of mothscaptured in three-day-old traps. If this effect is due to the proliferation of microbes inthe baits, the differences in response among treatments observed in the first experi-ment could, to some extent, reflect their suitability as a broth or culture medium fora particular type of microbial agent.

T

ABLE

1. S

PECIES

OF

N

OCTUID

M

OTHS

C

APTURED

IN

T

RAPS

B

AITED

WITH

U

NREFINED

P

ALM

S

UGAR

(J

AGGERY

)

OR

M

OLASSES

IN

S

EPT

./O

CT

. 1993. G

AINESVILLE

,F

LORIDA

.

Species Jaggery Molasses

Females Males Females Males

Mocis latipes (Guenee) 554 528 485 499Anticarsia gemmatalis Hubner 60 106 90 177Mocis disseverans (Walker) 26 36 36 43Anomis erosa Hubner 30 7 15 4Pseudaletia unipuncta (Haworth) 8 7 5 3Spodoptera frugiperda (J. E. Smith) 16 8 14 12Spodoptera exigua (Hubner) 3 0 2 0Spodoptera ornithogalli (Guenee) 1 6 5 2Spodoptera latifascia Walker 0 0 2 0Leucania sp. 15 17 16 17Agrotis subterranea F. 3 1 3 0Agrotis ipsilon (Hufnagel) 1 0 1 0Anicla infecta Ochsenheimer 12 8 5 8Helicoverpa zea (Boddie) 4 0 1 1


Additional research is needed to isolate and identify attractive odorants emanat-ing from these baits when placed in traps. The effectiveness of this system is limitedby the aging effects on bait attractiveness, by the small capacity of the trap for cap-tured moths, and by the fragile nature of the trap design. Ideally, a formulated lureof identified food attractants could be used to bait a dry trap design.

ACKNOWLEDGMENTS

Technical assistance was provided by K. M. Davis. Traps were generously providedby T. Howler (USDA, APHIS, Gainesville, FL) and J. Sivinski (USDA, ARS, Gaines-ville, FL.). Jaggery was recommended as a sweet bait for insects by S. Kulkarny,Gainesville, FL, and fruit pectin was suggested by R. Martinez, USDA, APHIS, Gua-temala City, Guatemala. This work was supported by grant US-2217-92 from theUnited States-Israel Binational Agriculture Research and Development Fund and bythe Cooperative State Research Service, USDA, Agreement No. 90-37250-5356.

REFERENCES CITED

DITMAN, L. P., AND E. N. CORY. 1933. The response of corn earworm moths to varioussugar solutions. J. Econ. Entomol. 26: 109-115.

DUNCAN, D. B. 1955. Multiple range and multiple F tests. Biometrics 11: 1-41.EYER, J. R. 1931. A four year study of codling moth baits in New Mexico. J. Econ. En-

tomol. 24: 998-1001.FROST, S. W. 1926. Bait pails as a possible control for the oriental fruit moth. J. Econ.

Entomol. 19: 441-450.FROST, S. W. 1928. Continued studies of baits for oriental fruit moth. J. Econ. Ento-

mol. 21: 339-348.FROST, S. W. 1929. Fourth contribution to a study of baits, with special reference to

the oriental fruit moth. J. Econ. Entomol. 22: 101-108.HOLLAND, W. J. 1903. The moth book. A guide to the moths of North America. Dover

Publications Inc., New York. 479 pp.LANDOLT, P. J., AND R. R. HEATH. 1989. Lure composition, component ratio, and dose

for trapping male Mocis latipes (Lepidoptera: Noctuidae) with synthetic sexpheromone. J. Econ. Entomol. 82: 307-309.

NEWELL, W. 1936. Progress report on the Key West (Florida) fruit fly eradicationproject. J. Econ. Entomol. 29: 116-120.

NORRIS, M. J. 1935. The feeding habits of the adult Lepidoptera Heteroneura. Trans.Roy. Entomol. Soc. London 85: 61-90.

OGUNWOLU, E. O., AND D. H. HABECK. 1975. Comparative life histories of three Mocisspp. in Florida (Lepidoptera: Noctuidae). Florida Entomol. 58: 97-103.

SARGENT, T. D. 1976. Legion of night. The underwing moths. Univ. MassachusettsPress, Amherst, MA, 222 pp.

STEEL, R. G. D., AND J. H. TORRIE. 1960. Principles and procedures of statistics.McGraw Hill Book Co., New York. 481 pp.

STRAYER, J. 1975. Sugarcane insect control. Florida Coop. Ext. Serv., Plant ProtectionPointers, Ext. Entomol. Rep No. 40. 6 pp.

Briano et al.: Fire Ant Diseases in Argentina

531

PROTOZOAN AND FUNGAL DISEASES IN

SOLENOPSIS RICHTERI

AND

S. QUINQUECUSPIS

(HYMENOPTERA: FORMICIDAE) IN BUENOS AIRES PROVINCE, ARGENTINA

J. B

RIANO

,

1

D. J

OUVENAZ

,

2

D. W

OJCIK

,

2

H. C

ORDO

1

AND

R. P

ATTERSON

2

1

USDA-ARS South American Biological Control Laboratory, Bolivar 1559 (1686) Hurlingham, Buenos Aires province, Argentina

2

USDA-ARS Medical and Veterinary Entomology Research Laboratory,P.O. Box 14565, Gainesville, FL, 32604, USA

A

BSTRACT

The diversity and abundance of protozoa and fungi infecting colonies of the fireants

Solenopsis richteri

Forel and

S. quinquecuspis

Forel were surveyed in BuenosAires province, Argentina. A total of 185 roadside sites was selected, and 1,836 colo-nies were sampled and examined under phase-contrast microscopy. Pathogens werefound at 32% of the sites and in 10% of the colonies. The microsporidium

Thelohaniasolenopsae

Knell, Allen & Hazard was the most common microorganism; it waspresent at 25% of the sites and 8% of the colonies. In some sites within the surveyedregion, the percentage of infected colonies with

T. solenopsae

ranged from 40 to 80%.Other pathogens present were the microsporidium

Vairimorpha invictae

Jouvenaz &Ellis and the fungus

Myrmecomyces annellisae

Jouvenaz & Kimbrough. A field sitewas selected for future ecological studies.

Key Words: Microsporidia, fungi, natural enemies, biological control, imported fireants, myrmecology.

R

ESUMEN

La diversidad y abundancia de protozoarios y hongos infectando colonias de las“hormigas coloradas”

Solenopsis richteri

Forel y

S. quinquecuspis

Forel fueron rele-vadas en la Provincia de Buenos Aires, Argentina. Un total de 185 sitios en banquinafueron seleccionados y 1.836 colonias fueron muestreadas y examinadas con micros-copía de contraste de fase. Se encontraron patógenos en 32% de los sitios y 10% de lascolonias. El microsporidio

Thelohania solenopsae

Knell, Allen y Hazard fue el más co-mun de los microorganismos encontrados, estuvo presente en el 25% de los sitios y en8% de las colonias. En algunos lugares de la región relevada, el porcentage de coloniasinfectadas con

T. solenopsae

fue del 40 al 80%. Otros patógenos presentes fueron el mi-crosporidio


Jouvenaz y Ellis y el hongo

Myrmecomyces annelli-sae

Jouvenaz y Kimbrough. Fue seleccionado un sitio de campo para futuros estudios

ecológicos.

The black imported fire ant,

Solenopsis richteri

Forel, and the red imported fireant,

S. invicta

Buren, were accidentally introduced into the United States from SouthAmerica (Lofgren 1986b).

S. richteri

has been displaced by

S. invicta

and now occursin a relatively small area of northeastern Mississippi and northwestern Alabama.

S.invicta

has spread dramatically and has become one of the most serious medical andeconomic pests in 11 southeastern States and Puerto Rico (Adams 1986, Lofgren1986a & 1986b, Mackay et al. 1992, Vinson & Mackay 1990, Vinson & Sorensen 1986).

532



However, their present status as different species should be investigated because, atleast in the United States, these two species cross and produce viable hybrids.

The black and red imported fire ants were introduced into the United States freeof their major natural enemies present in their native land (Jouvenaz et al. 1977).Consequently, a classical biological control approach could be possible with a complexof pathogens, parasites, and/or predators being introduced into the North Americanfire ant populations.

In South America, several pathogens are known from

S. invicta

,

S. richteri

andother members of the

S. saevissima

Smith complex; some of them were surveyed forin Brazil, Uruguay and Argentina (Allen & Buren 1974, Allen & Silveira Guido 1974,Jouvenaz 1983, 1986, Jouvenaz et al. 1980 & 1981, Wojcik et al. 1987).

Since 1988, part of the USDA research on biological control of fire ants has beenestablished at the USDA-ARS South American Biological Control Laboratory in Hurl-ingham, Buenos Aires Province, Argentina. Because the target species,

S. invicta

,does not occur within that area, the primary goal has been to search, select, and eval-uate the potential for natural enemies of native

Solenopsis

spp. as biological agentsfor control of

S. invicta

in the United States.The objectives of this work were to survey the diversity and abundance of patho-

gens in the area of the USDA laboratory and to select a convenient field study site toinitiate ecological studies.

M

ATERIALS

AND

M

ETHODS

The northeastern region of Buenos Aires province was surveyed from March toSeptember 1988. The area covered approximately 40,000 km

2

. A total of 185 collectingsites was selected systematically every 10 to 50 km (depending on time available)along the roadsides of the major highways of the region. At each site, usually the first10 colonies found on the roadside were sampled, which resulted in a total of 1,836 fireant colonies being sampled. According to Trager (1991), the only fire ant species inthat area are

S. richteri

and

S. quinquecuspis

. Although not all the samples wereidentified, we estimate that at least 80% of them were

S. richteri

.The colonies were sampled by inserting a 7-ml vial into the mounds. The inner

walls of the vials were dusted with talc to prevent escape of the ants. When severaldozen ants had fallen into the vials, they were removed, capped and placed on ice ina cooler for transportation to the laboratory. Once in the laboratory, the ants werekilled by freezing. Then each sample was placed in a glass tissue grinder with about2-4 ml of water and ground up for about 30 s. One drop of the aqueous extract was ex-amined under phase-contrast microscopy (400

×

) for the presence of spores of protozoaand fungi. The sensitivity of this procedure for detecting spores in low numbers hasbeen successfully tested in previous research (Jouvenaz et al. 1977).

The main criteria in the selection of the field site for future ecological studies were:the incidence of diseases in the local fire ant populations, the type of habitat, and theproximity to the USDA laboratory. Arbitrarily, the limits of the region surveyed wereestablished at 300 km from the USDA laboratory.

R

ESULTS

AND

D

ISCUSSION

The region surveyed and the approximate location of most of the pathogen-positivesampling sites are shown in Fig. 1. Of the total of 185 sites, protozoa and fungi werefound at 59 (32%) of the sites (Table 1). The most common pathogen was the protozoan

Thelohania solenopsae

Knell, Allen, & Hazard (1977) (Microsporida: Thelohaniidae),


533

it was present at 46 (25%) of the sites. The fungus

Myrmecomyces annellisae

Jouvenaz& Kimbrough (1991) (Deuteromycotina: Hyphomycetes) was found at 14 (8%) of thesites and the protozoan


Jouvenaz & Ellis (1986) (Microsporida:

Figure 1. Region surveyed in Buenos Aires province and location of most of thepathogen-positive sites. Shaded areas show the main sampling areas.

534



Burenellidae) was found at 9 (5%) of the sites. At some sites, more than one pathogenwas found infecting different colonies, but never the same colony. However, dual in-fections with

T. solenopsae

and

V. invictae

were observed within extremely stressedlaboratory colonies (J. B., unpublished data).

According to the incidence of diseases, the region surveyed was arbitrarily dividedinto 4 main areas (Fig. 1): Saladillo (livestock grazing area), Isla Talavera (swampyarea), Pergamino (annual crops area), and Bragado (mixture of annual crops and live-stock grazing area). The area of Saladillo (180 km SW of Buenos Aires) had the high-est incidence of pathogens with 65% of the sites infected (Table 1). The most commonpathogen was

T. solenopsae

; it was present at 47% of the sites. The fungus

M. annel-lisae

was found at 14% of the sites, and

V. invictae

at 12%. The area of Isla Talavera(100 km NW of Buenos Aires) showed the second highest incidence of diseases with50% of the sites infected;

T. solenopsae

was found at 33% and

V. invictae

at 20%. In thearea of Bragado (224 km W of Buenos Aires) 22% of the sites were infected, 18% with

T. solenopsae

and 11% with

M. annellisae

. The area of Pergamino (250 km NW of Bue-nos Aires) had 21% of the sites infected with

T. solenopsae

. No other disease was foundin this area. In other areas of the province, only 16% of the sites had diseases;

T. so-lenopsae

was present in 14% and

V. invictae

in less than 3%. (Table 1).Of the 1,836 fire ant colonies sampled, 182 (10%) were infected, 8% with

T. sole-nopsae

, 1% with

V. invictae

and 1% with

M. annellisae

(Table 2). The highest percent-age of infected colonies (33%) was found in the area of Isla Talavera, then inPergamino (17%), Saladillo (12%), and Bragado (9%). In all these areas,

T. solenopsae

was the most common pathogen (Table 2). In the other areas of the region, only 4% ofthe colonies were infected.

At several collecting sites,

T. solenopsae

was found infecting a high proportion ofcolonies. For example: 80% at Rt. 8, km 254 (Pergamino); 66% at Rt. 8, km 247 (Per-gamino); 50% at Rt. 205, km 180 (Saladillo); 47% at Rt. 12, km 104 (Isla Talavera);and 40% at Rt. 5, km 224 (Bragado). At one collecting site,

V. invictae

infected 60% ofthe colonies (Rt. 12, km 91, Isla Talavera). The prevalence of

T. solenopsae

and

V. in-victae

in those sites represents the highest infection rates with these pathogens in fireants ever reported for South America (See references below). However, the total per-centages of colonies infected with these two microsporidia in the region surveyed (8and 1%, respectively) were similar to those found in

S. invicta

in Mato Grosso andMato Grosso do Sul, Brazil (2.2 to 11.4% and 2 to 4% of the colonies respectively) (Jou-

T

ABLE

1. P

ATHOGEN

-

POSITIVE

S

AMPLING

S

ITES

W

ITHIN

A

REA

S

URVEYED

IN

B

UENOS

A

IRES

P

ROVINCE

.

Area

Number of Sampling

Sites

Number (%) of Sampling Sites with Pathogens

T. solenopsae V. invicta M. annellisae

Total

1

Saladillo 49 23 (47) 6 (12) 7 (14) 32 (65)Isla Talavera 6 2 (33) 1 (20) 0 3 (50)Bragado 27 5 (18) 0 3 (11) 6 (22)Pergamino 24 5 (21) 0 0 5 (21)Others 79 11 (14) 2 (<3) 0 13 (16)

Total 185 46 (25) 9 (5) 14 (8) 59 (32)

1

Sites had >1 pathogen, so totals are lower than the sum of 3 columns.


535

venaz et al. 1980, Jouvenaz 1986, Jouvenaz & Ellis 1986, Wojcik 1986, Wojcik et al.1987). In two surveys conducted in 1987 in the provinces of Buenos Aires, Santa Fe,and Entre Rios, Argentina, 10.5% of the fire ant colonies were infected with

T. sole-nopsae

(D. J. & D. W., unpublished data).The relatively high prevalence of fire ant pathogens in Argentina and Brazil con-

trasts with the lack of specific pathogens in the North American populations of theblack and red imported fire ants (Jouvenaz et al. 1977). Surveys in areas infested withboth species of imported fire ants in the United States revealed few colonies infectedwith pathogens (Lofgren et al. 1975). Jouvenaz et al. (1977) reported considerablevariation in the rate of infection of

S. invicta

with

M. annellisae

in Florida, Alabamaand Georgia. This is probably the only (specific?) microorganism imported into theUnited States in association with the imported fire ants.

The prevalence of pathogens reported in this paper was probably underestimatedbecause the sampling of the fire ant colonies included only adult workers. Possiblyearly infective stages were present in immature fire ants and missed. In addition,other pathogens known to occur in fire ants in South America, such as bacteria and vi-ruses (Avery et al. 1977, Jouvenaz 1983) were not detected with the light microscope.The prevalence of these other pathogens deserves further investigation.

We conclude that, within the region surveyed, the microsporidium

T. solenopsae

was the most common pathogen of indigenous fire ant populations. In some sites, it in-fected a high proportion of the colonies. Therefore, it should be evaluated as a poten-tial biological control agent for the imported fire ants in the United States. Althoughthe areas of Isla Talavera and Pergamino showed higher number of colonies infectedwith T. solenopsae, the area of Saladillo was selected as the field study site because:(1) it showed the highest number of sites infected with T. solenopsae, (2) it was theonly area where the three pathogens were present, (3) it represented the most appro-priate habitat for future long-term ecological studies, and (4) it was near the USDAlaboratory and frequent field work would be simplified.

ACKNOWLEDGMENT

We would like to thank David Williams (USDA-ARS Medical and Veterinary En-tomology Research Laboratory, Gainesville, FL, USA), Philip Koehler (University ofFlorida, Gainesville, FL, USA) and Daniel Gandolfo (USDA-ARS South American Bi-ological Control Laboratory, Hurlingham, Buenos Aires, Argentina) for reviewing thismanuscript.

TABLE 2. PATHOGEN-POSITIVE COLONIES WITHIN AREA SURVEYED IN BUENOS AIRESPROVINCE.

Area

Number of Sampled Colonies

Number (%) of Colonies with Pathogens

T. solenopsae V. invicta M. annellisae Total

Isla Talavera 90 24 (27) 6 (7) 0 30 (33)Pergamino 175 29 (17) 0 0 29 (17)Saladillo 613 49 (8) 12 (2) 14 (2) 75 (12)Bragado 205 16 (8) 0 3 (<2) 19 (9)Others 753 27 (<4) 2 (<1) 0 29 (4)

Total 1,836 145 (8) 20 (1) 17 (1) 182 (10)


REFERENCES CITED

ADAMS, C. T. 1986. Agricultural and medical impact of the imported fire ants, pp 48-57 in C. S. Lofgren and R. K. Vander Meer [eds.], Fire ants and leaf-cuttingants: biology and management. Westview Press, Boulder, CO. 434 pp.

ALLEN, G. E., AND W. F. BUREN. 1974. Microsporidian and fungal diseases of Solenop-sis invicta Buren in Brazil. J. New York Entomol. Soc. 82: 125-30.

ALLEN, G. E., AND A. SILVEIRA-GUIDO. 1974. Occurrence of microsporidia in Solenop-sis richteri and Solenopsis sp. in Uruguay and Argentina. Florida Entomol. 57:327-9.

AVERY, S. W., D. P. JOUVENAZ, W. A. BANKS, AND D. W. ANTHONY. 1977. Virus-likeparticles in a fire ant, Solenopsis sp., (Hymenoptera: Formicidae) from Brazil.Florida Entomol. 60: 17-20.

JOUVENAZ, D. P. 1983. Natural enemies of fire ants. Florida Entomol. 66: 111-121.JOUVENAZ, D. P. 1986. Diseases of Fire Ants: Problems and Opportunities, pp. 327-

338 in C. S. Lofgren and R. K. Vander Meer [eds.], Fire ants and leaf-cuttingants: biology and management. Westview Press, Boulder, CO. 434 pp.

JOUVENAZ, D. P., AND E. A. ELLIS. 1986. Vairimorpha invictae n. sp. (Microspora: Mi-crosporida), a parasite of the red imported fire ant, Solenopsis invicta Buren(Hymenoptera: Formicidae). J. Protozool. 33: 457-461.

JOUVENAZ, D. P., AND J. W. KIMBROUGH. 1991. Myrmecomyces annellisae gen. n., sp.n. (Deuteromycotina: Hyphomycetes), an endoparasitic fungus of fire ants, So-lenopsis spp. (Hymenoptera: Formicidae). Mycolog. Res. 95: 1395-1401.

JOUVENAZ, D. P., G. E. ALLEN, W. A. BANKS, AND D. P. WOJCIK. 1977. A survey forpathogens of fire ants, Solenopsis spp., in the southern United States. FloridaEntomol. 60: 275-279.

JOUVENAZ, D. P., W. A. BANKS, AND J. D. ATWOOD. 1980. Incidence of pathogens in fireants, Solenopsis spp. in Brazil. Florida Entomol. 63: 345-346.

JOUVENAZ, D. P., C. S. LOFGREN, AND W. A. BANKS. 1981. Biological control of im-ported fire ants: a review of current knowledge. Bull. Entomol. Soc. America 27:203-208.

KNELL, J. D., G. E. ALLEN, AND E. I. HAZARD. 1977. Light and electron microscopestudy of Thelohania solenopsae n. sp. (Microsporida: Protozoa) in the red im-ported fire ant, Solenopsis invicta. J. Invertebr. Pathol. 29: 192-200.

LOFGREN, C. S. 1986a. The economic importance and control of imported fire ants inthe United States, pp. 227-256 in S. B. Vinson [ed.], Economic impact and con-trol of social insects. Praeger, N. Y. 421 pp.

LOFGREN, C. S. 1986b. History of imported fire ants in the United States, pp. 36-49 inC. S. Lofgren and R. K. Vander Meer [eds.], Fire ants and leaf-cutting ants: bi-ology and management. Westview Press, Boulder, CO. 434 pp.

LOFGREN, C. S., W. A. BANKS, AND B. M. GLANCEY. 1975. Biology and control of im-ported fire ants. Annu. Rev. Entomol. 20: 1-30.

MACKAY, W. P., S. B. VINSON, J. IRVING, S. MAJDI, AND C. MESSER. 1992. Effect of elec-trical fields on the red imported fire ant (Hymenoptera: Formicidae). Environ.Entomol. 21: 866-870.

TRAGER, J. C. 1991. A revision of the fire ants, Solenopsis geminata group (Hy-menoptera: Formicidae: Myrmicinae). J. New York Entomol. Soc. 99: 141-198.

VINSON, S. B., AND W. P. MACKAY. 1990. Effects of the fire ant Solenopsis invicta onelectrical circuits and equipment, pp. 496-503 in R. K. Vander Meer, K. Jaffeand A. Cedeño [eds.], Applied myrmecology: a world perspective. WestviewPress, Boulder, CO. 741 pp.

VINSON, S. B., AND A. A. SORENSEN. 1986. Imported fire ants: Life history and impact.Texas A & M University, College Station, Texas and Texas Department of Agri-culture, Austin, Texas. 28 pp.

WOJCIK, D. P. 1986. Observations on the biology and ecology of fire ants in Brazil, pp.88-103 in C. S. Lofgren and R. K. Vander Meer [eds.], Fire ants and leaf-cuttingants: biology and management. Westview Press, Boulder, CO. 434 pp.

Briano et al.: Fire Ant Diseases in Argentina 537

WOJCIK, D. P., D. P. JOUVENAZ, W. A. BANKS, AND A. C. PEREIRA. 1987. Biological con-trol agents of fire ants in Brazil, pp. 627-628 in J. Eder and H. Rembold [eds.],Chemistry and biology of social insects. Verlag J. Perperny, Munchen. 757 pp.

538



NATURAL ENEMIES OF

BEMISIA TABACI

(HOMOPTERA: ALEYRODIDAE) IN CUBA

A

NTONIO

C

ASTINEIRAS

Tropical Research and Education CenterUniversity of Florida, IFAS

18905 SW 280th St.Homestead, Florida 33031

The sweet potato whitefly,

Bemisia tabaci

(Guennadius), reached pest status inCuba during the growing season of 1989-90, seriously damaging tomato, eggplant,squash, cucumber and other vegetable crops. Taxonomic status of the genus

Bemisia

in Cuba is currently under study. As the correct identity of the species damagingcrops is yet to be determined, the name

B. tabaci

will be used in this note.Several parasitoids (Aphelinidae, Platygasteridae, Encyrtidae), predators

(Chrysopidae, Anthocoridae, Coccinellidae, Phytoseiidae, Stigmaeidae) and the ento-mopathogenic fungus

Paecilomyces farinosus

(Dickson & Fries) Brown & Smith havebeen reported as natural enemies of

B. tabaci

from different countries (Cock 1993).Polaszek et al. (1992) noted parasitoids collected from the Caribbean region, but thereare no records of biological control agents of

B. tabaci

from Cuba.Surveys for natural enemies were conducted in the most important agricultural

areas located at the Western, Central and Eastern regions of Cuba, and on the Isle ofPines, from January to April 1990, 1991 and 1992. The climate in Cuba during this pe-riod is characterized by temperatures of 23-25

°

C, 80-85% relative humidity and lessthan 5 mm rainfall. Sampled plots (1 ha) were selected from eggplant, tomato, pepper,cabbage, squash, cucumber, and sweet potato fields not heavily sprayed with chemicalinsecticides, or only treated with


(Berl.). Samples consisted ofone mature leaf, taken at random from each 50 plants on an X-shaped design, once ineach growing season. A total of 5,000 leaves were collected from 100 fields every year.More than 10,000 whitefly nymphs per year were collected. All leaf parts with white-flies were cleaned of other insects, cut and kept in vials until adult parasitoidsemerged. Whiteflies were considered parasitized only when a parasitoid emergedfrom a nymph. When fungal parasites were found, only dead nymphs surrounded bymycelium were considered infected. The fungus was isolated, identified, and inocu-lated on healthy nymphs in the laboratory to corroborate its pathogenic effect. Imma-ture predators (except spiders) were fed with whitefly larvae until development wascompleted. Material from the surveys was identified at the Institute of Plant Protec-tion (Ministry of Agriculture) and the National Museum of Natural History (Academyof Sciences), Havana, Cuba.

Four parasitoids, an entomopathogenic fungus, one hyperparasitoid, and fivepredators were found (Table 1).

Encarsia luteola

Howard,

E. nigricephala

Dozier and

E. quaintancei

Howard.—

E. luteola

and

E. quaintancei

were the parasitoids most frequently found in Cubaduring the growing seasons of 1989-92. They were present on all seven crop specieson which whiteflies were collected. Less abundant than the former two species,

E. ni-gricephala

was only found in the Western region of the country. According to Polas-zek et al. (1992),

E. nigricephala

,

E. luteola

and

E. quaintancei

are well representedin the Caribbean Basin, but the three species had not been documented from Cubabefore.

Eretmocerus

sp. (Aphelinidae), and

Signiphora

sp. (Signiphoridae).—Parasitoidsof the genus

Eretmocerus

are frequently associated with sweet potato whitefly (Polas-

Scientific Notes

539

zek et al. 1992). Some species, such as

E. haldemani

Howard (Gerling 1967) or

E.mundus

Mercet (Kapadia & Puri 1990), are major constituents of the parasitoid com-plex of

B. tabaci

in different countries. In Cuba,

Eretmocerus

sp. was only found once,on peppers, in the Central and Eastern regions in 1991.

Signiphora

sp. is a hyperpar-asitoid and it was collected from samples taken from tomato, cabbage, squash, cucum-ber and sweet potato from the Western and Central agricultural areas of Cuba.

Paecilomyces fumosoroseus

(Wize).—

P. fumosoroseus

is considered a potential bio-logical control agent of the sweet potato whitefly (Osborne et al. 1990). First epizooticsof

P. fumosoroseus

were found in April 1990 in the South Havana area after a periodof heavy rain. The same situation was observed in 1991 and 1992 when the weatherwas very humid. Osborne et al. (1990) found over 90% mortality of fourth instarnymphs treated with the fungus in the laboratory. In Cuba, natural epizootics killed70-80% of nymphs in the field. The fungus also appeared spontaneously on

B. tabaci

in some of the greenhouses in the Institute of Plant Protection in Havana.

Theridula gonygaster

Simon and

Theridula

sp.—These spiders of the family The-ridiidae build small webs on the lower surface of the leaves where they catch adultsof the sweetpotato whitefly. They were observed very frequently in heavily infestedcucumber, squash, sweet potato and tomato fields in the Western and Central regionsof the country.

T

ABLE

1. N

ATURAL

E

NEMIES

OF

B

EMISIA

T

ABACI

IN

C

UBA

, E

FFECT

ON

THE

H

OST

,

AND

D

ISTRIBUTION

.

Natural EnemiesParasitism or

Predation

Distribution

Western Central Eastern Isle of Pines

Encarsia luteola

Howard20-30% X X X X

E. nigricephala

Dozier 15-20% X

E. quaintancei

Howard 25-35% X X X X

Eretmocerus

sp. Not evaluated X X

Signiphora

sp. Not evaluated X X

Paecilomycesfumosorosus

(Wize)

70-80% X

Theridulagonygaster

Simon

5-10 adults/web X X

Theridula

sp. 5-10 adults/web X

Delphastuspallidus

LeC.

Not evaluated X X X X

Chrysopa exterior

Navas

Not evaluated X X X X

Cyrtopeltis varians

(Dist.)

Not evaluated X X X

540



Delphalstus pallidus

LeC. (Coccinellidae),

Chrysopa exterior

Navas (

Chrysopidae

)and

Cyrtopeltis varians

(Dist.) (Miridae).—Despite their wide distribution in Cubathese predators were not frequently associated with

B. tabaci.

Immatures found oninfested sweet potato and cucurbit leaves in the field completed their development inthe laboratory when they were fed on sweetpotato whitefly larvae. In California,

D.pusillus

is under study for use in management programs for

B. tabaci

(Heinz et al.1994).

In general, parasitoids and predators did not seem to have a strong effect as lim-iting factors of the sweet potato whitefly populations. Apparently crop species did notaffect parasitism, but a greater diversity of natural enemies was found in cucurbits,tomato and sweet potato. The fungus

P. fumosoroseus

showed a high percentage ofparasitism, but only during very humid periods, making it a good candidate for bio-control in greenhouses.

This is Journal Experimental Series No R-04117.

S

UMMARY

Surveys for natural enemies of the sweet potato whitefly,

Bemisia tabaci

(Guenn.),were conducted in Cuba from January to April 1990, 1991 and 1992. Four parasitoids(

Encarsia luteola

Howard,

E. nigricephala

Dozier,

E. quaintancei

Howard and

Eret-mocerus

sp.), one hyperparasitoid (

Signiphora

sp.), an entomopathogenic fungus(

Paecilomyces fumosoroseus

(Wize)) and five predators (

Theridula gonygaster

Simon,

Theridula

sp.

Delphastus pallidus

LeC.,

Chrysopa exterior

Navas, and

Cyrtopeltisvarians

(Dist.)) were detected.

R

EFERENCES

C

ITED

C

OCK

, J. M. 1993. (Ed.)

Bemisia tabaci

-an update 1986-1992 on the cotton whiteflywith an annotated bibliography. International Institute of Biological Control.,Ascot, Berks, UK. 78 pp.

G

ERLING

, D. 1967. Bionomics of the whitefly-parasite complex associated with cottonin Southern California (Homoptera:Aleurodidae; Hymenoptera:Aphelinidae).Ann. Entomol. Soc. America 60:1306-1321.

H

EINZ

, K. M., J. R. B

RAZZLE

, C. H. P

ICKETT

, E. T. N

ATWICK

, J. M. N

ELSON

,

AND

M. P.P

ARELLA

. 1994. Predatory beetle may suppress silverleaf whitefly. California.Agric. 48:35-40.

K

APADIA

, M. N.,

AND

S. N. P

URI

. 1990. Development, relative proportions and emer-gence of Encarsia transvena (Timberlake) and Eretmocerus mundus Mercet,important parasitoids of Bemisia tabaci (Guennadius). Entomol. 15:235-239.

OSBORNE, L. S., G. K. STOREY, C. W. MCCOY, AND J F. WALTEW. 1990. Potential forcontrolling the sweet potato whitefly, Bemisia tabaci, with the fungus, Paecilo-myces fumosoroseus. Proceedings and abstracts, Vth International Colloquiumon Invertebrate Pathology and Microbial Control, Adelaide, Australia, 20-24August 1990. Adelaide, Australia, pp. 386-390.

POLASZEK, A., G. A. EVANS, AND F. D. BENNETT. 1992. Encarsia parasitoids of Bemisiatabaci (Hymenoptera:Aphelinidae, Homoptera: Aleyrodidae): a preliminaryguide to identification. Bull. Entomol. Res. 82:375-392.

Scientific Notes

541

EXTRACTION OF HOUSE FLY (DIPTERA: MUSCIDAE) LARVAE FROM POULTRY MANURE USING BERLESE/TULLGREN

FUNNELS

D

ONALD

R. B

ARNARD

Medical and Veterinary Entomology Research LaboratoryAgriculture Research Service

U.S. Department of AgricultureGainesville, FL 32604 U.S.A.

Accurate determination of the numbers of late instar house fly (

Musca domestica

L.) larvae in samples of poultry manure is the first step in developing density estima-tors for populations of pupae and emerging adult flies. These density estimators canbe used to anticipate fly abatement needs on agricultural facilities and to improve theefficiency of releases of pupal parasitoids in augmentation and inundation-based bio-logical control programs for

M. domestica

. Presently, there is no practical method fordetermining the density of house fly larvae in a sample of poultry manure. Estimatesof this factor (Geden & Stoffolano 1987; Stafford & Bay 1987) have been deducedmainly from extraction data using Berlese/Tullgren funnels (Brydon & Fuller 1966).However, the accuracy of these devices for separating house fly larvae from samplesof poultry manure is not well understood.

This study was made to assess four parameters whose influence on the mean andvariance of numbers of third instar house fly larvae extracted from samples of poultrymanure placed in Berlese/Tullgren funnels is unknown. These parameters are: (1) thesize of the manure sample, (2) the number of larvae in the sample, (3) the cumulativeextraction rate of larvae from the manure sample over time, including the relation be-tween extraction rate and temperature change within the manure sample, and (4) theeffect of enclosure of larvae and manure in sample containers (for

≤

4 h) on the accu-racy of larval extraction.

Berlese/Tullgren funnels were constructed according to the design of Brydon &Fuller (1966). Poultry manure for the experiments was collected for 24 h in trays sus-pended beneath commercial laying hens. The manure was stored at -20

°

C then al-lowed to thaw to ambient temperature 8 h before use.

A 4

×

4 factorial design was used to determine the effect of larval density and ma-nure volume on the accuracy of larval extraction. Each treatment combination wasreplicated four times (

n

= 64) and comprised one of four volumes of manure (100, 200,300, and 400 cc) and one of four levels of larval density (50, 100, 300, 500). Two hoursbefore a test began, known numbers of 72-hour-old early third instar larvae reared at26

°

C on the Gainesville house fly diet (Hogsette 1992) were place on, and allowed topenetrate, each manure sample. Individual samples were placed onto a single layer ofcheesecloth on top of the 5 mm mesh screen (Brydon & Fuller 1966) in each funnel andshaped to a thickness of

≈

40 mm. Larvae exiting the samples, after funnel coverswere positioned and the lights (100 watts) turned on, dropped into 70% ethanol in wa-ter in 0.25-liter glass jars at the bottom of each funnel. Jars were replaced at 12 h in-tervals until no larvae were collected for 2 consecutive intervals.

To determine the cumulative number of larvae extracted from a manure sampleover time, nine samples of 100 larvae in 400 cc of manure were prepared as describedabove and placed in funnels at 0 h. Collection jars were removed from funnels at 6, 24,30, and 48 h. Temperature change at the top, middle, and bottom positions in threemanure samples was monitored continuously, and reported hourly, using an Omni-data

EL-820 data logger (Omnidata, P.O. Box 3489, Logan, UT 84321) and ES-060temperature probes. The experiment was repeated four times (

n

= 36).

542



A 2

×

4 factorial design was used to determine the effect of

≤

4 h of enclosure of lar-vae and manure in sample containers on the accuracy of larval extraction. Treatmentfactors were open/closed sample container and the length of time the container wasopen or closed (1, 2, 3, or 4 h). At 1 h intervals, beginning 4 h before all samples wereplaced in the funnels (0 h) and ending at 1 h, six samples of 100 larvae in 400 cc of ma-nure were placed into sealable plastic containers (10

×

10

×

15 cm) and held at ambi-ent temperature (22-26

°

C). Air-tight lids were placed on three of the containers andthese containers labelled as closed. The remaining three unsealed containers were la-belled as open. All samples were placed in funnels at 0 h and the funnels operated, asbefore, for 48 h.

In all tests, the percent extraction of larvae for each funnel was calculated as a ra-tio of the numbers of larvae in the collection jar to the numbers of larvae in the ma-nure sample, multiplied

×

100. Data for percent extraction (transformed to arcsin)were analyzed using analysis of variance procedures (SAS Institute, Inc. 1988).Means separation was performed using Tukey’s studentized range test at

P

= 0.05.The relationship between cumulative extraction rate (%CR) and temperature changein the manure sample at top (t), middle (m), and bottom (b) positions was determinedusing the regression model: %CR = t + t

2

+ m + m

2

+ b + b

2

(SAS Institute, Inc. 1988).Percent extraction of third instar house fly larvae from samples of poultry manure

in Berlese/Tullgren funnels was not influenced by the size of the manure sample or bythe numbers of larvae in the sample (Table 1). The overall percent extraction of larvaein this study (

n

= 118) was 98.5

±

1.9.The length of time Berlese/Tullgren funnels were operated influenced the percent

extraction of fly larvae (

F

1,35

= 39.69,

P

= 0.01). Significantly fewer (57.7%) larvae wererecovered after 6 h of operation than after 24 (90.2%), 30 (96.6%) or 48 h (96.6%). Cu-mulative percent extraction of larvae was related to the linear and quadratic effectsof manure temperature change (

F

2,143

= 54.68; R

2

= 0.437) at the top position. Therewas no significant change in manure temperature at any position after 30 h of funneloperation.

The enclosure of fly larvae and manure in sample containers for up to 3 h did notinfluence overall percent extraction of larvae compared with the data for larvae inopen containers for the same length of time (x

open

= 98.8

±

1.8%; x

closed

= 99.2

±

0.7%).The results at 4 h (x

open

= 99.3

±

0.05%; x

closed

= 64.0

±

55.4%), however, were equivocal.Of the three containers closed for 4 h, one yielded 0 larvae while percent extraction forthe remaining two was 96%. The results at 4 h, for closed containers, while not statis-tically different from the earlier times, indicate that enclosing larvae and manure ina sample container for more than 3 h at 22-26

°

C (as might be necessary during trans-port of samples) will result in variable extraction responses.

T

ABLE

1. M

EAN

P

ERCENT

E

XTRACTION

(

±

SEM)

OF

T

HIRD

I

NSTAR

H

OUSE

F

LY

L

ARVAEFROM

P

OULTRY

M

ANURE

IN

B

ERLESE

/T

ULLGREN

F

UNNELS

.

Manure Volume

Number of Larvae

50 100 300 500

100 cc 99.5 (1.0) 88.1 (18.7) 95.8 (3.9) 99.3 (0.9)200 cc 95.5 (4.1) 100.0 (0.0) 99.3 (0.5) 98.8 (0.5)300 cc 100.0 (0.0) 99.8 (0.5) 98.3 (2.2) 99.5 (1.0)400 cc 99.3 (2.0) 98.3 (1.7) 98.5 (1.3) 98.1 (0.8)

Scientific Notes

543

S

UMMARY

Berlese/Tullgren funnels provide an accurate estimate of the density of third in-star house fly larvae in samples of poultry manure. Percent extraction of larvae didnot vary with sample size, when the range of sample sizes was between 100 and 400cc of poultry manure, and was unaffected by larval densities from 50 to 500 per sam-ple. Maximum extraction of house fly larvae requires

≥

30 h of funnel operation. Theaccuracy of extraction decreases when manure and larvae are enclosed together in asample container for more than 3 h prior to placement of samples in funnels.

R

EFERENCES

C

ITED

B

RYDON

, H. W.,

AND

R. G. F

ULLER

. 1966. A portable apparatus for separating fly lar-vae from poultry droppings. J. Econ. Entomol. 59: 448-452.

G

EDEN

, C. J.,

AND

J. G. S

TOFFOLANO

. 1987. Dispersion patterns of arthropods associ-ated with poultry manure in enclosed houses in Massachusetts: spatial distri-bution and effects of manure moisture and accumulation time. J. Entomol. Sci.23: 136-148.

H

OGSETTE

, J. A. 1992. New diets for production of house flies and stable flies (Diptera:Muscidae) in the laboratory. J. Econ. Entomol. 85: 2291-2294.

SAS I

NSTITUTE

, I

NC

. 1988. SAS/STAT user’s guide, release 6.03. SAS Institute, Cary,NC.

S

TAFFORD

III, K. C.,

AND

D. E. B

AY

. 1987. Dispersion pattern and association of housefly

Musca domestica

(Diptera: Muscidae) larvae and both sexes of

Macrochelesmuscaedomesticae

(Acari: Macrochelidae) in response to poultry manure mois-

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

ture, temperature, and accumulation. Environ. Entomol. 16: 159-164.

Scientific Notes

543

PARASITOIDS AND PREDATORS ASSOCIATED WITH

SYNTOMEIDA EPILAIS

(LEPIDOPTERA: ARCTIIDAE) ON OLEANDER

H

EATHER

J. M

C

A

USLANE

1

AND

F

RED

D. B

ENNETT

1,2

1

Department of Entomology & Nematology, University of Florida, P.O. Box 110620, Gainesville, FL 32611-0620

2

Current Address: Crofton, Baldhoon Road, Laxey, IM4 7NA,Isle of Man, United Kingdom

Oleander,

Nerium oleander

L., is an apocynaceous ornamental shrub native to theMediterranean region. It is planted widely in the southern United States, from Flor-ida to California, throughout the Caribbean and into Mexico (Reinert 1980). In Flor-ida, oleander is ubiquitous in central and southern regions, often planted inmunicipal parks, recreational areas, highway right-of-ways, around public buildingsand in homeowner’s yards.

One of the major insect pests of oleander in Florida and the Caribbean is the larvaof the arctiid moth,

Syntomeida epilais

(Walker) (the “polka-dot wasp moth”). Olean-der caterpillars are predictable and perennial pests, often defoliating oleander bushesduring multiple generations per year. This species appears to be a native of the Neo-tropics. It is indigenous throughout Florida, the Keys and the Dry Tortugas (Kimball

544



1965), and it is present in Mexico, the Caribbean region and northern countries ofSouth America (Bratley 1932).

Larvae and adults of

S. epilais

contain cardiac glycosides obtained from their hostplant (Rothschild et al. 1973) and are distasteful to avian predators (Jones 1934), asare other cardiac glycoside-containing insects such as the monarch butterfly,

Danausplexippus

(L.) (Brower et al. 1967). Parasitoids recorded from

S. epilais

include the ta-chinid flies

Chetogena (=Euphorocera) floridensis

(Townsend),

Lespesia aletiae

(Riley)(Patton 1958) and

Lespesia

sp. (Fernald 1934). No published records of predationhave been found.

Collections of immature stages of

S. epilais

were made in several areas of Floridaas part of an effort to understand the contribution of natural enemies to populationregulation of the oleander caterpillar. In addition to several small collections, largenumbers of immatures were collected on four occasions (Table 1). Larvae were fed cutoleander foliage while prepupae and pupae were placed in cylindrical plastic contain-ers (15-cm diam

×

25-cm height) for emergence of adults.On Davis Island on 4 August 1991, numerous larvae were collected on oleander

and from the leaf litter below oleander. Pupae were collected from aggregations in de-pressions in the trunk of an oak tree. Individuals of an unidentified

Brachymeria

spe-cies were observed flying around the plants and pupation sites. These wasps were alsoobserved in August 1994, in Clearwater, although no collections were made. Waspsflew quickly and buzzed loudly, similar to houseflies.

Brachymeria incerta

(Cresson)and two species of tachinid (

C. floridensis

and

L. aletiae

) emerged from

S. epilais

col-lected on Davis Island. This is a new host record for

B. incerta

which has been listedboth as a primary parasitoid attacking pupae of many Lepidoptera and as a secondaryparasitoid attacking

Carcelia lagoae

(Townsend) and other Lepidoptera-parasitizingtachinids (Burks 1960).

Brachymeria incerta

was confirmed as a primary parasitoidof

S. epilais

by presenting mated females with pupae of

S. epilais

that had beenreared from second instars in the laboratory and, therefore, were unlikely to containlarvae of larval/pupal parasitizing tachinids. Adult

B. incerta

emerged from these pu-pae approximately 1 month later.

T

ABLE

1. D

ATES

AND

L

OCATIONS

OF

C

OLLECTIONS

FOR

S.

EPILAIS

N

ATURAL

E

NEMIESFROM

O

LEANDER

IN

F

LORIDA

.

Collection DateCollection Location

(Collected by)

S. epilais

ImmatureStates Collected

Natural EnemiesEncountered

4-Aug-91 Davis Island, Hillsborough Co.(F. D. Bennett)

Larvae, prepupae, pupae

Brachymeria incerta

, tachinids

11-Aug-93 Gainesville,Alachua Co.

(H. J. McAuslane)

Largae, prepupae, pupae

Brachymeria incerta

, fire ants

20-Jun-94 Homestead,Dade Co.

(H. J. McAuslane)

Larvae Pathogens, tachinids

6-Jan-95 Gainesville,Alachua Co.

(H. J. McAuslane)

Larvae, prepupae, pupae

Tachinids, pteromalid hyperparasite of tachinid, pathogens

Scientific Notes

545

In Gainesville, on 11 July 1993, predation by an immature pentatomid

Podisusmaculiventris

(Say) was observed on a third-instar

S. epilais

feeding on oleander. Thenymph was reared to adulthood on third-instar oleander caterpillars. This pentato-mid is a generalist feeder but has not been recorded from

S. epilais

according toMcPherson (1982). Many

B. incerta

were reared from the pupae collected from an ag-gregated pupation site in Gainesville on 11 August 1993. The geographic range of

B.incerta

, listed as central and southern Florida, West Indies, Guyana and Brazil(Burks 1960), can be extended to north Florida. The pupal mass was also heavily in-fested by red imported fire ants,

Solenopsis invicta

Buren, which were observed feed-ing actively on

S. epilais

pupae.Two tachinids emerged from a collection of larvae in Homestead on 20 June 1994.

However, of the 300+ larvae collected, more than 80% of them succumbed to a patho-gen. The symptomology proceeded from the excretion of large, liquid feces, to the des-iccation of the abdomen, to death. However, the specimens were not examinedmicroscopically to determine the nature of the pathogen.

Of the approximately 100 larvae collected in Gainesville on 8 January 1995, 13went on to pupate and produce moths while 57 produced

L. aletiae

. In addition, threeadults of the hyperparasitic perilampid

Perilampus hyalinus

Say were reared frompuparia of

L. aletiae

. While

P. hyalinus

has been collected from

Lespesia frenchii

(Will-iston),

L. euchaetiae

(Webber) and

L. melalophae

(Allen) (Krombein et al. 1979), it hasnot been recorded from this species. The remainder of the larvae suffered mortalitydue to a pathogen that produced symptoms characteristic of a viral infection (degen-eration of the larval body to a black liquid).

Despite the casual nature of these observations, they indicate that a number of po-tentially significant natural enemies and pathogens of

S. epilais

exist in Florida.However, despite these sources of mortality, oleander caterpillar continues to defoli-ate oleander regularly in southern portions of the state. Despite the toxic nature of

S.epilais

larvae and pupae, the predators and parasitoids associated with this speciesare generalist feeders. In a similar situation,

Euploea core corinna

(W. S. Macleay), anative nymphalid in Australia that has exploited oleander as a novel host, is attackedby two species of tachinid (

Paradrino laevicula

Mesnil and

Winthemia neowinthemi-oides

[Townsend]) and

Brachymeria lasus

(Walker) which also are generalist feeders(Rahman & Zalucki 1986).

Thanks to F. Mead, G. Steck and L. Stange (Florida Dept. of Agriculture and Con-sumer Services, Division of Plant Industry, Gainesville, FL) for identification of thepentatomid, and tachinid and hymenopteran parasitoids, respectively. Thanks also toD. Habeck and H. Frank (Dept. of Entomology & Nematology, University of Florida)for critical reviews of an earlier version of this manuscript. This is Florida Agricul-tural Experiment Station Journal Series No. R-04448.

S

UMMARY

The oleander caterpillar,

Syntomeida epilais

, is attacked in Florida by various gen-eralist parasitoids including the tachinids

Lespesia aletia

and

Chetogena (=Euphoro-cera) floridensis

and the chalcidid

Brachymeria incerta

(new host record). Fire ants,

Solenopsis invicta

, a predatory pentatomid,

Podisus maculiventris

, and pathogensalso attack

S. epilais.

R

EFERENCES

C

ITED

B

RATLEY

, H. E. 1932. The oleander caterpillar,

Syntomeida epilais

, Walker. FloridaEntomol. 15: 55-64.

546



B

ROWER

, L. P., J. V. Z. B

ROWER

,

AND

J. M. C

ORVINO

. 1967. Plant poisons in a terres-trial food chain. Proc. Natl. Acad. Sci. 57: 893-898.

B

URKS

, B. D. 1960. A revision of the genus

Brachymeria

Westwood in America northof Mexico. Trans. American Entomol. Soc. 86: 225-273.

F

ERNALD

, H. T. 1934. [Oleander - Polka dot wasp moth (

Syntomeida epilais

Walk.)].Bull. Insect Pest Survey 14: 23-24.

J

ONES

, F. M. 1934. Further experiments on the coloration and relative acceptabilityof insects to birds. Trans. Roy. Entomol. Soc. Lond. 82: 443-453.

K

IMBALL

, C. P. 1965. Lepidoptera of Florida. Div. of Plant Ind., Florida Dept. of Agr.,Gainesville, Florida.

K

ROMBEIN

, K. V., P. D. H

URD

, J

R

., D. R. S

MITH

,

AND

B. D. B

URKS

. 1979. Catalog of Hy-menoptera in America north of Mexico. Vol. 1. Symphyta and Apocrita (Para-sitica). Smithsonian Institution Press, Washington, D.C.

MCPHERSON, J. E. 1982. The Pentatomoidea (Hemiptera) of northeastern NorthAmerica with emphasis on the fauna of Illinois. Southern Illinois UniversityPress, Carbondale, IL.

PATTON, C. N. 1958. A catalogue of the Larvaevoridae of Florida. Florida Entomol. 41:29-39, 77-89.

RAHMAN, H. U., AND M. P. ZALUCKI. 1986. Parasitoid records for Euploea core corinna(W. S. Macleay) (Lepidoptera: Nymphalidae) in south-eastern Queensland.Aust. Entomol. Mag. 12: 109-111.

REINERT, J. A. 1980. Control of the oleander caterpillar on oleander. Proc. FloridaState Hort. Soc. 93: 168-169.

ROTHSCHILD, M., J. VON EUW, AND T. REICHSTEIN. 1973. Cardiac glycosides (heartpoisons) in the polka-dot moth Syntomeida epilais Walk. (Ctenuchidae: Lep.)with some observations on the toxic qualities of Amata (=Syntomis) phegea

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

(L.). Proc. Roy. Soc. London Ser. B 183: 227-247.

546



PARASITES OF THE PEPPER FLOWER-BUD MOTH(LEPIDOPTERA: GELECHIIDAE) IN FLORIDA

F

RED

D. B

ENNETT

Entomology and Nematology Department, University of Florida,Gainesville, FL 32611-0620, USA

Current address: Crofton, Baldhoon Road, Laxey, Isle of Man IM4 7NA

The pepper flower-bud moth,

Symmetrischema capsica

(Bradley & Polovny, (Lep-idoptera: Gelechiidae) described from specimens reared from

Capsicum annuum

L. inMontserrat, West Indies, occurs widely in the Caribbean. It has been an importantpest of

Capsicum

spp. in this region for at least 65 years (Wilson 1923). It occurs instates in the USA bordering the Gulf of Mexico and also has been collected in Mexico(Bradley & Polovny 1965). In 1988 this species was reared from an uncultivated

Cap-sicum

sp. in Honduras (unpubl. data). Initially placed in the genus

Gnorimoschema

by Bradley & Polovny (1965), the species was transferred to the genus

Symme-trischema

by Polovny (1967). Earlier it had been confused with, and identified as,

Gnorimoschema gudmanella

(Walsingham); many of the earlier references to its sta-tus and records of its natural enemies appear in the literature under that name. Inthe Caribbean, Parasram (1973) and des Vignes (1978) reported flower-bud drop of 40to 100% as the result of attack by

S. capsicum

.This insect was first reported from Florida in 1944 and from Texas in 1945 (An-

nand 1945). Schuster (1960) reported 70-100% bud damage, seasonally, in the Rio

Scientific Notes

547

Grande Valley, Texas. Currently, in Florida, pepper bud-moth is not a pest of commer-cial peppers; possibly it is controlled by the pesticides applied against pepper weeviland beet armyworm (D. J. Schuster pers. comm.).

Des Vignes (1978 & 1981) reported on studies of parasites of

S. capsica

which heundertook in Trinidad, whereas, the only report of parasites in the USA appears to bethat given by Schuster (1960) (see Table 1).

In May 1985, while surveying for natural enemies of

Parlatoria ziziphi

(Lucas) inthe “Little Haiti” section of Miami, I collected flower buds of

Capsicum minima

L. in-fested by

S. capsica

(identity confirmed by John Heppner) and reared two species ofparasites. I made further collections on subsequent surveys from June 1986 to July1987 and assessed the level of attack and the occurrence of parasites. The data ob-tained from these collections are given in Table 2. Only the braconid

Pseudapantelesdignus

(Mues.) (formerly

Apanteles

dignus

) and the eulophid

Euderus

sp. (possibly

E.purpureus

Yoshimoto) were reared.

P. dignus

has been reared from this host in theCaribbean and from several other gelechiid moths in the USA; among the latter arerecords from Florida (Marsh 1979). Because of the uncertainty of its specific identity,it is not possible to give the distribution of

E

. ?

purpureus

. Specimens identified as

Euderus

sp. are known to attack pepper bud-moth in Puerto Rico and Trinidad (desVignes 1978, 1981). Records of parasites of

S. capsica

, including those from the cur-rent survey, are given in Table 1. In Florida, only two parasites have been reared from

S. capsica

, in contrast to Trinidad where des Vignes (1981), during a two-year study,recorded 6 species. It is probable that a more extensive survey in Florida would con-tribute new parasite records.

I greatly appreciate the determinations of the parasitoid species by P. M. Marsh(Braconidae) and M. E. Schauff (Eulophidae), and of the pepper flower-bud moth byJ. Heppner.

T

ABLE

1. P

ARASITES

RECORDED

FROM

S

YMMETRISCHEMA

CAPSICA

.

Family Genus and Species Stage Distr. Ref.

1

Encyrtidae

Apsilophrys capsicum

(Burks)

egg/lar. TrinidadP. Rico

2

Braconidae

Agathis

sp. larval Trinidad 2

Bracon

sp. larval Trinidad 2

Chelonus phthorimaeae

Gahanegg/lar. Texas 3

Chelonus

sp. egg/lar. P. Rico 2

Orgilis capsicolae

Mues. larval Texas 3

Pseudapanteles dignus

(Mues.)

larval TrinidadP. RicoFlorida

124

Bethylidae

Perisierola

sp. larval Trinidad 2Eulophidae

Euderus

sp. larval TrinidadP. Rico

12

Euderus

?

purpureus

Yosh. larval Florida 4Pteromalidae Genus & sp. indet. larval Montserat 1

1

Ref. = Reference: 1 = Parasram 1973; 2 = des Vignes 1978 & 1981; 3 = Schuster 1960; 4 = current study.

548



T

AB

LE

2.

P

AR

AS

ITIS

M

OF

S.

CA

PS

ICA

IN

P

EP

PE

R

F

LO

WE

R

-B

UD

S

C

OL

LE

CT

ED

AT

“L

ITT

LE

H

AIT

I

”, M

IAM

I

, J

UN

E

198

6-J

UL

Y

198

7.

Dat

eB

uds

Col

lect

edB

uds

Att

acke

dB

uds

wit

h

Lar

vae

Un

para

siti

zed

Lar

vae

Par

asit

ized

by

% P

aras

itiz

ed

Pse

ud

apan

tele

sE

ud

eru

s

86.0

6.28

4645

3321

—12

36.4

86.0

7.21

5432

1818

——

—86

.09.

2453

1807

07—

——

87.0

1.27

2008

0303

——

—87

.04.

2763

5323

132

843

.587

.05.

2857

5544

351

820

.587

.06.

2248

4523

114

852

.187

.07.

2845

2805

05—

——

Tota

l11

37

3627

.6

Scientific Notes

549

University of Florida, Florida Agricultural Experimental Station Journal SeriesNo. R-04104.

S

UMMARY

Parasites of the pepper flower-bud moth


(Bradley &Polovny),´ (Lepidoptera: Gelechiidae) are reported from Florida for the first time.Combined parasitism by the braconid

Pseudapanteles dignus

(Mues.), and the eu-lophid

Euderus

?

purpureus

Yoshimoto as high as 52.1% (average 28.6%) was re-corded.

R

EFERENCES

C

ITED

A

NNAND

, P. N. 1945. Report of the chief of the bureau of entomology and plant quar-antine USDA. 1944-1945: 30

B

RADLEY

, J. D.,

AND

D. P

OLOVNY

.´ 1965. The identity of the pepper flower-bud moth.Bull. Entomol. Res. 56: 57-65.

D

ES

V

IGNES

, W. G. 1978. Notes on the taxonomy, biology and economic importance of


(Bradley & Povolny).´ Jour. Agric. Soc. Trinidad & To-bago. 78: 311-318.

D

ES

V

IGNES

, W. G. 1981. Notes on parasites and predators of

Symmetrischema cap-sica

(Bradley & Povolny).´ Jour. Agric. Soc. Trinidad & Tobago. 81: 50-58.M

ARSH

, P. M. 1979. Family Braconidae, pp. 144-313

in

K. V. Krombein et al. [eds.].Catalog of Hymenoptera in America north of Mexico. Smithsonian InstitutionPress, Washington, DC, Vol. 1, 1198 p.

P

ARASRAM

, S. 1973. The pepper flower bud moth in the Caribbean - an evaluation.Proc. 11th Ann. Meeting Caribbean Food Crops Soc., Barbados, pp. 466-470.

P

OLOVNY

,´ D. 1967. Genitalia of some Neartic and Neotropical members of the tribeGnorimoschemini, Lepidoptera, Gelechiidae. Acta Entomol. Mus. Prague. 97:51-67.

S

CHUSTER

, M. F. 1960. A pepper flower-bud moth

Gnorimoschema gudmanella

. Jour.Econ. Entomol. 52: 117-118.

W

ILSON

, C. E. 1923. Truck-crop pests in the Virgin Islands and methods of combatting

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

them. Bull. Virgin Island (U.S.) Agric. Exp. Sta. 4: 24-25.

Scientific Notes

549

EFFECT OF TIME OF BAIT EXPOSURE ON NUMBER OF WIREWORMS (COLEOPTERA: ELATERIDAE) FOUND AT BAITS

R

ONALD

H. C

HERRY

AND

J

OSE

A

LVAREZ

Everglades Research and Education CenterP. O. Box 8003

Belle Glade, Florida 33430

In Florida, the Everglades Agricultural Area (EAA) encompasses 260,000 hectares(650,000 acres) of the upper Everglades extending from Lake Okeechobee south about50 km. Most of the soils in the EAA are organic (Histosols) generally containing 85%or more organic matter by weight (Snyder et al. 1978). The area is intensively farmed,with sugarcane and winter vegetables being the principal crops.

Wireworms are ubiquitous soil insect pests of most of these crops and soil insecti-cides are frequently applied at planting for wireworm control. Cherry (1993) showed

550



that rolled oats was a simple, attractive bait that could be used for sampling

Melan-otus communis

(Gyllenhal) and

Conoderus

sp. wireworms in the organic soils of theEAA. In that study, baits were left in fallow fields for 14 d. However, the effect of timeof bait exposure on wireworm numbers found at baits was not determined. Moreover,the effect of time of bait exposure on wireworm numbers found at baits has receivedlittle attention in general. Toba & Turner (1983) reported on tests in WashingtonState in which wireworm baits were left in fields for one, two, or three weeks. Resultswere variable with significant differences in wireworm numbers, primarily

Ctenicerapruinina

(Horn), at baits with different time exposures in some tests and not in othertests. Seal et al. (1992) reported on the effect of the postemergence period of bait ex-posure on wireworm numbers in Georgia. Baits removed one week after emergencecontained significantly more wireworms than similar baits removed 2 and 3 weeks af-ter emergence in fields dominated by

Conoderus scissus

Schaeffer and

C. amplicollis

(Gyllenhal). In a field dominated by

C. rudis

(Brown), more wireworms were found fol-lowing increased exposure of the bait. The objective of our study was to determine theeffect of time of bait exposure on numbers of wireworms found at baits under the fieldconditions of the EAA.

All tests were conducted on Histosols at the Everglades Research and EducationCenter at Belle Glade, Florida. Tests were conducted in 10 fallow fields which hadbeen disked following sugarcane. Fields were surveyed by digging to determine thepresence of wireworms before tests were conducted. Each bait consisted of 200 g ofrolled oats placed in a plastic bag. Four baits plus two controls (= no bait) were in eachreplication in a 3

×

2 pattern with baits and controls 5 m apart. Ten replications wereused in each field with replications being 10 m apart. A randomized complete block de-sign was used for placement of the baits and controls. Holes were dug 15 cm deep,each bait poured from its plastic bag into the hole, a flag placed through the bait tomark for recovery, and the bait covered with soil. At the time of bait placement in thefield, one unbaited control sample from each replication was dug-up to represent 0days after bait placement. Thereafter, one bait in each replication was dug-up at 7-day intervals up to 28 days after bait placement in the field. The second unbaited con-trol sample was also dug-up at 28 days. By comparing the two control samples (0 vs.28 d), it was possible to determine if wireworm population density changed in theplots during the tests. If the wireworm population density did not change in the plots,this would indicate that wireworm populations at baits were due to wireworm attrac-tion to baits over time and not changes in population levels in the plots over time.Baits were recovered by digging-up the bait and adjacent soil in a 25

×

25

×

20 cm deepsample and placing the sample in a bucket. Samples were stored in a laboratory atabout 23

°

C. Each sample was visually examined for wireworms for 30 min in the lab-oratory. Wireworms were then stored in alcohol and later identified by microscopic ex-amination.

M. communis

(Gyllenhal) was identified using the key of Riley & Keaster(1979).

Conoderus

sp. was identified by J. B. Heppner at the Florida Division of PlantIndustry, Gainesville. Other wireworm species were less than 10% of total wirewormsfound at baits and were not identified. Ten tests were conducted in 10 different fieldsfrom October, 1992 to September, 1993.

Since greater than 90% of the wireworms found at food baits were either

Con-oderus

sp. or

M. communis

, statistical analysis was restricted to these two groups. Apaired t-test analysis was conducted to determine if wireworm numbers were signifi-cantly different in the controls in any of the 10 fields at 0 d versus 28 d. The main ob-jective of our tests was to determine the overall response of wireworms to baits overtime. Hence, data from the 10 fields were pooled for regression analysis. Initially, thetotal number of wireworms at baits was plotted against time (0, 7, 14, 21, 28 d). Visual

Scientific Notes

551

observation indicated a quadratic function for

M. communis

and a cubic function for

Conoderus

sp. Thereafter, these models were used for the two wireworm groups usingthe General Linear Models Procedure (SAS 1990). The quadratic equation for

M. com-munis

was:

Y = a

o

+ a

1

X

i

+ a

2

X

2

i; whereY = Total number of wireworms at baitsX

i

= Days baits left in fieldi = 0, 7, 14, 21, and 28 daysa

o

, a

1

, and a

2

= the intercept, linear term, and thequadratic term, respectively.

The cubic equation for

Conoderus

sp. was:Y = a

o

+ a

1

X

i

+ a

2

X

2

i + a

3

X

3

i; wherea

3

= the cubic term; all other terms were previously explained.

A t-test analysis showed no significant difference (P=0.05) in mean numbers of

Conoderus

sp. or

M. communis

at 0 vs 28 d in unbaited samples in any of the 10 fields.These data indicate that wireworms at baits over time were due to bait attraction andnot changes in population densities during the tests.

Regression results from the estimated equations are shown in Table 1. The equa-tion for

M. communis

is highly predictive of the actual observed wireworm distribu-tion over time (Fig. 1). Both linear (X) and quadratic (X

2

) terms are highly significantstatistically; furthermore, the coefficient of determination is a high 0.9954 and the co-efficient of variation is a low 5.96. None of the coefficients from the equation for

Con-oderus

sp. are statistically significant. However, the equation is very close to theactual distribution of wireworms in the fields (Fig. 2), as shown by the high coefficientof determination (0.9196), but not by the coefficient of variation (39). Both equationsshow that time of bait exposure was a strong predictor of wireworm numbers found atrolled oats baits.

Currently, soil insecticides are frequently applied for wireworm control when var-ious crops are planted in the EAA. In many cases, soil insecticides are not neededsince wireworm populations are too low to cause economic damage (Cherry et al.1993). However, few growers sample for wireworms since this procedure is difficultdue to digging, sorting through soil, and low numbers of wireworms normally found.Cherry (1993) and this study have shown that rolled oats baits are attractive to wire-

T

ABLE

1. R

EGRESSION

R

ESULTS

FROM

THE

E

STIMATED

E

QUATIONS

.

a

Equation

Item

M. communis Conoderus

sp.

Intercept -0.11 16.21X 6.65

b

(0.46) -4.24 (9.70)X

2

-0.15

c

(0.02) 1.08 (0.88)X

3

— -0.03 (0.02)R

2

0.99 0.92C.V. 5.96 38.97

a

Figures in parentheses are the standard errors of the estimated coefficients.

b

Statistically significant at P<0.005.

c

Statistically significant at P<0.01.

552



worms in the EAA and that the time of bait exposure affects the number of wirewormsfound at baits. These latter two studies provide basic data for using rolled oats baitsas a sampling tool to determine the necessity of applying insecticides for wirewormcontrol when planting various crops in the EAA.

Florida Agricultural Journal Series No. R-04466.

Figure 1. Regression equation for M. communis. Y = the total number of wire-worms at baits and X = the duration of bait exposure in days.

Figure 2. Regression equation for Conoderus sp. Y = the total number of wire-worms at baits and X = the duration of bait exposure in days.

Scientific Notes

553

S

UMMARY

Our data show that the duration of exposure of rolled oats baits is important in de-termining the numbers of wireworms found at baits under field conditions. Wire-worms of both

Conoderus

sp. and

M. communis

were found in increasing numbersfrom 0 to 21 days and then declined in number at 28 days.

R

EFERENCES

C

ITED

C

HERRY

, R. 1993. Baits for sampling wireworms (Coleoptera:Elateridae) in organicsoils (Histosols) of southern Florida. J. Agric. Entomol. 10: 185-188.

C

HERRY

, R., G. P

OWELL

,

AND

M. U

LLOA

. 1993. Reduced soil insecticide use in sugar-cane planted after rice. Sugar y Azucar. 88: 37-38.

R

ILEY

, T. J.,

AND

A. J. K

EASTER

. 1979. Wireworms associated with corn: identificationof larvae of nine species of

Melanotus

from the North Central States. AnnalsEntomol. Soc. America 72: 408-414.

SAS L

ANGUAGE

. 1990. Version 6. First Ed. Cary, North Carolina. 1042 pages.S

EAL

, D., R. C

HALFANT

,

AND

M. H

ALL

. 1992. Effectiveness of different seed baits andbaiting methods for wireworms (Coleoptera: Elateridae) in sweetpotato. Envi-ron. Entomol. 21: 957-963.

S

NYDER

, G., H. B

URDINE

, J. C

ROCKET

, G. G

ASCHO

, D. H

ARRISON

, G. K

IDDER

, J.M

ISHOE

, D. M

YHRE

, F. P

ATE

,

AND

S. S

HIH

. 1978. Water table management fororganic soil conservation and crop production in the Florida Everglades. Univ.of Florida, Agricultural Experiment Station Bulletin 801, Gainesville, 63 pp.

T

OBA

, H.,

AND

J. T

URNER

. 1983. Evaluation of baiting techniques for sampling wire-worms (Coleoptera: Elateridae) infesting wheat in Washington. J. Econ. Ento-

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

mol. 76: 850-855.

Scientific Notes

553

SOME CORRECTIONS TO THE GENERIC RECORDS OF CENTRAL AMERICAN EUMOLPINAE (CHRYSOMELIDAE:

COLEOPTERA)

R. W

ILLS

F

LOWERS

1

1

Agricultural Research Programs, Florida A&M University, Tallahassee, FL 32307

Seeno & Wilcox (1982) list 56 genera of Eumolpinae (Coleoptera: Chrysomelidae)as occurring in Central America. As part of an ongoing project on the Eumolpinae ofCosta Rica, I have found that eight of these generic records are probably incorrectsince they are based on species that have been transferred to other genera or arebased on very doubtful records. Since study and identification of Central AmericanEumolpinae is difficult enough without worrying about genera that do not in fact oc-cur there, I feel it worth while to review these questionable records.

Agbalus

Chapuis 1874:242. Type species

Agbalus sericeus

Chapuis by monotypy.Valid genus, South America.

Hylax

Lefèvre 1884:xlv. Type species

Amasis chalcaratus

Chapuis by monotypy. Validgenus, Central and South America.

Bechyné (1950a) restricted this genus to its type species,

A. sericeus

Chapuis fromBrazil, transferring all other species to

Hylax

Lefèvre. Those other species, including

554



chalybaeus chiriquiensis

Jacoby,

hoegei

Jacoby,

mexicanus

Jacoby,

puncticollis

Jacoby,

quadriplagiatus

Jacoby,

tenebrosus

Jacoby, and

viridis

Bowditch from Central Amer-ica are listed under

Hylax

in the Bechyné (1953) catalog of the Eumolpinae.

Chalcoplacis

Chevrolat 1837:433. Type species

Colaspis fulgurans

Klug by mono-typy. [

C. incerta

Dejean, designated as type species by Chevrolat 1845, is a

nomen nudum

. Subsequent designation of

Lamprosphaerus abdominalis

Balyas type species by Baly 1875 is unnecessary and invalid.] Valid genus, SouthAmerica.

Parachalcoplacis

Bechyné 1950a:268. Type species

Lamprosphaerus mrazi

Bechyné by original designation. Synonym of

Chalcoplacis

(Monrós & Bechyné1956).

Lamprosphaerus

Baly 1859:124. Type species


Baly byoriginal designation. Valid genus, Central and South America.

Antitypona

Weise 1921:17. Type species

Lamprosphaerus collaris

Baly by originaldesignation. Valid genus, Central and South America.

Neochalcoplacis

Bechyné 1950a:268. Type species

Chalcoplacis dimidiata

Lefèvre byoriginal designation. Valid genus, South America.

Chalcoplacis

is in the center of a very tangled nomenclatural web. Baly began it bydesignating the species


Baly as the type species firstfor his genus

Lamprosphaerus

(Baly 1859), then for Chevrolat’s

Chalcoplacis

(Baly1865). Weise (1921) resolved this problem by restricting the definition of

Chalcoplacis

to

Chalcoplacis incerta

Chevrolat, the species designated as type species by Chevrolat(1845), and establishing the genus

Antitypona

with Baly’s

Lamprosphaerus collaris

as the type species. Weise also stipulated that all

Lamprosphaerus

then listed in the

Coleopterorum Catalogus

(Clavareau 1914) are

Antitypona

and all

Chalcoplacis

inthat catalog are

Lamprosphaerus

.Bechyné (1950a) erected a new genus

Neochalcoplacis

, designating

Chalcoplacisdimidiata

Lefèvre as the type species, including two species from Central America [

N.constituta

Bechyné and

N. fulvipes

(Jacoby)], and erected the subgenus

Parachalco-placis

for a South American species. In the Bechyné’s 1953 catalog,

Parachalcoplacis

became a valid genus including numerous South American species described byBechyné or transferred by him from

Antitypona

and

Lamprosphaerus

. However, bothof the Central American species (

constituta

and

fulvipes

) were transferred to

Lam-prosphaerus

.

Neochalcoplacis

was now restricted to its type species.Monrós & Bechyné (1956) identified

Colaspis fulgurans

Klug as the type species of

Chalcoplacis

(by monotypy), stated that

Chalcoplacis incerta

Chevrolat was a

nomennudum

, and synonymized

Parachalcoplacis

with

Chalcoplacis

(but they did not syn-onymize

Neochalcoplacis

, as stated by Seeno & Wilcox 1982).

Colaspis fulgurans

hadpreviously been designated as the type species of

Parachalcoplacis

(Bechyné 1950a).Thus, both

Chalcoplacis

and

Neochalcoplacis

are valid genera but neither is knownfrom Central America at present. On the other hand,

Lamprosphaerus

and

Antity-pona

are valid and widespread Central American genera.

Corysthea

Baly 1865:336. Type species

Corysthea ferox

Baly by monotypy. Validgenus, South America.

The single Central American species,

Corysthea violacea

described by Jacoby(1882), was transferred to

Hylax

by Bechyné (1953) and renamed

pseudoviolaceus

(Bechyné 1955) since the specific epithet

violaceus

was preoccupied by

Hylax auratusviolaceus

(Jacoby). No species of

Corysthea

are presently known from Central Amer-ica.

Coytiera

Lefèvre 1875:116. Type species

Coytiera marginicollis

Lefèvre by monotypy.Valid genus, South America.

Scientific Notes

555

Euphrytus

Jacoby 1881:124. Type species

Euphrytus aeneus

Jacoby, designated byBechyné (1950a). Valid genus, Central America.

Bechyné (1953) transferred

Colaspis melancholica

Jacoby to this genus. In 1957,he transferred three other Jacoby

Coytiera

species,

costata

Jacoby,

fulvipes

Jacoby,and

rugipennis

Jacoby to the genus

Euphrytus

Jacoby, thus leaving

melancholica

asthe only valid Central American

Coytiera

. In 1969, Bechyné & Bechyné again moved

melancholica

, this time to the genus

Freudeita

Bechyné, based, in their opinion, onthe structure of the antenna (see below for further discussion of this species). No validspecies of

Coytiera

are presently known to exist in Central America.

Freudeita

Bechyné 1950b:241. Type species

Colaspis parallina

Erichson, by originaldesignation. Valid genus, South America.

Bechyné (1953) and Wilcox (1985) list the species F. dentifera Bechyné from CostaRica. Bechyné & Bechyné (1969) also transferred Coytiera melancholica (Jacoby) andColaspis balyi Jacoby to Freudeita, and noted the similarity of both species to F. den-tifera. Blake (1976) treated melancholica, balyi, and six closely related species as aspecies complex within Colaspis. She ignored (as was her custom) all Bechyné’s workand mentioned neither F. dentifera nor other South American Freudeita that Bechynédescribed as similar to dentifera. In Bechyné’s (1950b) original description of Freude-ita, the defining character is described as antennae with middle segments noticeablywider than either basal or apical segments. This character is readily apparent in theSouth American species originally placed in Freudeita (Bechyné 1950b) but is veryweakly developed in melancholica—as the Bechynés themselves admitted. AlthoughBlake’s across-the-board dismissal of everything Bechyné wrote was, on occasion, ex-cessive and incorrect, I agree in this case that the melancholica complex is better leftin Colaspis. Based on published descriptions, dentifera is close to C. melancholica andC. spinigera Blake (all three have a subapical spine on the hind tibia of the male); ac-cordingly Bechyné’s species is now Colaspis dentifera (Bechyné) new combination.

Ischyrolamprina Bechyné 1950a:265. Type species Spintherophyta lampros Lefèvreby original designation. Valid genus, South America.

Ischyrolampra Lefèvre 1885:166 [Name for Eulampra Baly which was preoccupied]Type species Eulampra batesi Baly by monotypy. Valid genus, South America.

Agrosterna Harold 1875:103. Type species Agrosterna buphthalma Harold by mono-typy. Valid genus, Central and South America.

This was originally described as a subgenus of Ischyrolampra Lefèvre, which in-cluded I. panamensis Jacoby from Panamá. In his catalog, Bechyné (1953) transferredI. panamensis to the genus Agrosterna Harold. Wilcox (1983) apparently missed thischange and listed panamensis as Ischyrolampra, and this was perpetuated in theSeeno & Wilcox list. All valid species now included in both Ischyrolamprina and Is-chyrolampra are South American.

Longeumolpus Springlová 1960:5. Type species Eumolpus imperalis Baly, by origi-nal designation. Valid genus, South America, Martinique.

I have found no records from continental Central America but Springlová (1960),in her revision of the genus, lists the type species, L. imperalis (Baly), as occurring onMartinique in the Lesser Antilles. To be consistent with other listings in Seeno & Wil-cox (1982), this record should be considered “W. Indies”.

Lycaste Gistel 1848:123. Type species Lycaste trichoa Gistel by monotypy.Callicolaspis Bechyné 1950a:275. Junior synonym.

Bechyné (1950a) mentioned a specimen of Lycaste cuneiformis (Bechyné) in theFrey Museum from Mexico, a record he considered erroneous for this otherwise SouthAmerican genus. This appears to be the only published Central American record. As


Lycaste species are among the largest and showiest of the Neotropical Eumolpinae, itis not likely that collectors in Mexico have been missing them all these years.

SUMMARY

Eumolpinae genera Agbalus, Chalcoplacis, Corysthea, Coytiera, Freudeita, Ischy-rolamprina, Longeumolpus, and Lycaste do not occur in Central America, contrary toprevious records in the literature. Current synonymies and known ranges for each ofthese genera are given.

REFERENCES CITED

BALY, J. S. 1859. Descriptions of new genera and species of phytophagous insects.Ann. Mag. Nat. Hist., ser. 3 4:55-61, 124-128, 270-275.

BALY, J. S. 1865. Descriptions of new genera and species of Phytophaga. Trans. Ento-mol. Soc. London, ser. 3 2:333-357.

BECHYNÉ, J. 1950a. Les générotypes des Eumolpides de l’Amérique du Sud et Centreavec les diagnoses des formes nouvelles (Col. Phytoph. Chrysomeloidea). Mitt.München Entomol. Ges. 40: 264-292.

BECHYNÉ, J. 1950b. Notes sur les Chrysomeloidea de l’Amérique du Sud et du Centre.(Col. Phyotph.). Entomol. Arb. Mus. G. Frey 1:237-269.

BECHYNÉ, J. 1953. Katalog der neotropischen Eumolpiden (Col. Phytoph. Chrysome-loidea). Entomol. Arb. Mus. G. Frey, 4:26-304.

BECHYNÉ, J. 1955. Reise des Herrn G. Frey in Südamerika: Eumolpidae (Col. Phy-tophaga). Entomol. Arb. Mus. G. Frey 6:569-657.

BECHYNÉ, J. 1957. Eumolpides Neotropicaux de la collection du Museo Civico de Sto-ria Naturale “Giacomo Doria” di Genova. Ann. Mus. Genova 69:226-247.

BECHYNÉ, J. B., AND B. SPRINGLOVÁ DE BECHYNÉ, J. 1969. Notas sobre PhytophagaAmericanos (Coleoptera). Rev. Fac. Agron. (Maracay). 3:5-64.

BLAKE, D. H. 1976. Colaspis melancholica Jacoby and its close relatives (Coleoptera:Chrysomelidae). J. Washington Acad. Sci. 65: 158-162.

CHAPUIS, F. 1874. In Histoire Naturelle des Insectes. Genera des Coléoptères. T. La-cordaire and F. Chapuis, eds. 10:1-455.

CHEVROLAT, L. A. A. 1837. In Dejean, P. F. M. A., Catalogue des coléoptères de la col-lection de M. le comte Dejean. livr. 5, pp. 361-443.

CHEVROLAT, L. A. A. 1845. In d’Orbigny, C. 1841-1849. Dictionnaire universel d’His-toire Naturelle, Paris, 5:1-792.

CLAVAREAU, H. 1914. Coleopterorum Catalogus, Chrysomelidae: Eumolpinae. pars59(11):1-215.

GISTEL, J. 1848. Naturgeschichte des Thierreichs für höhere Schulen, 216pp.HAROLD, E. VON. 1875. Über Chrysomelidae aus Cordova. Coleopterologische Hefte

XIV:95-106.JACOBY, M. 1881. Biologia Centrali-Americana, Insecta, Coleoptera, Eumolpidae,

Chrysomelidae. 6:73-144.JACOBY, M. 1882. Biologia Centrali-Americana, Insecta, Coleoptera, Eumolpidae,

Chrysomelidae. 6:145-224.LEFÈVRE, E. 1875. Descriptions d’eumolpides nouveaux ou peu connus. Revue et Ma-

gasin de Zoologie, ser. 3:102-138.LEFÈVRE, E. 1885. Eumolpidarum hucusque cognitarum catalogus, sectionum con-

spectu systematico, generum sicut et specierum nonnullarum novarum de-scriptionibus adjunctis. Mém. Soc. roy. Sci. Liège, ser. 2, 11:1-172.

MONRÓS, F., AND J. BECHYNÉ. 1956. Über einige verkannte Chrysomeliden-Namen.Entomol. Arb. Mus. G. Frey 7:1118-1137.

SEENO, T. N., AND J. A. WILCOX. 1982. Leaf beetle genera (Coleoptera: Chrysomel-idae). Entomography 1:1-221.

Scientific Notes 557

SPRINGLOVÁ, B. 1960. Essai monographique du genre Eumolpus. Mem. Inst. Roy. Sci.Nat. Belgique 2nd Ser. Fas. 60. 79 pp.

WEISE, J. 1921. Wissenshaftliche Ergebnisse der schwedischen entomolgischen Reisedes Herrn. Dr. A. Roman in Amazonas 1914-1915. 6. Chrysomelidae. Ark. Zool.14:1-205.

WILCOX, J. A. 1983. Checklist of the beetles of North and Central America and theWest Indies. Vol. 8. The leaf beetles and the bean weevils. Family 129. Chry-somelidae. E. J. Brill. New York. 166 pp.

Erratum

633

ERRATUM

L. S. Bauer—

Resistance: A Threat to the Insecticidal Crystal Proteins of

Bacillus thu-ringiensis.

Change first sentence in conclusions (p. 434) to read: The genetic capacity of insect

populations to evolve resistance to

Bt

δ

-endotoxins is now well documented in many

species within

three

different insect orders.

hoy: myths of managing resistance 385 introduction to...

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