|
Hoy: Myths of Managing Resistance
INTRODUCTION TO THE SYMPOSIUM,
THE MYTHS OF MANAGING RESISTANCE
MARJORIE A. HOY
Entomology & Nematology Department, University of Florida,
Gainesville, FL 32611-0620
Resistance to pesticides in arthropod pests is a serious and increasingly perplexing
problem 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 we
have learned much during the past 40 years. However, to some degree, much of the
discussion about 'resistance management' has been based on 'myths'. As an organizer
of 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 the
Florida 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try to resistance could not be published in this series.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 resistance
to pesticides in Florida and cite examples of resistance that limit pest management
options.
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 of
their effects on the physiology of arthropods, verify that resistance is a normal re-
sponse to diverse environmental stresses. 'Pesticide resistance' is part of a general
stress response with a long evolutionary history.
Leah Bauer describes what we know about resistance to various toxins of Bacillus
thuringiensis (B.t.) strains. B.t. provides microbial control of an increasingly diverse
group of arthropod species and is an increasingly important tool for integrated pest
management programs. The deployment of transgenic crop plants containing B.t.
toxin genes is likely to be an effective method for inducing resistance in agricultural
pests. Despite the diversity of B.t. toxin genes isolated and cloned, cross resistances
are 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 resistance
management needs a paradigm shift that can best be accomplished if we recognize
that pest management must be changed from a single-tactic strategy to a multi-tactic
mode. Delaying resistance, whether to traditional pesticides or to transgenic plants
with toxin genes, will require that we develop truly integrated pest management pro-
grams, incorporating all appropriate tactics, including host plant resistance, cultural
controls, biological controls, genetic controls, and biorational controls. Pesticides
should be reserved for situations in which they perform best-as tools to resolve an
unexpected pest population outbreak. Effective, fully-integrated IPM programs will
delay resistance because the number and rates of pesticide applications can be re-
duced.
Florida Entomologist 78(3) September, 1995
PESTICIDE RESISTANCE IN FLORIDA INSECTS LIMITS
MANAGEMENT OPTIONS
GARY L. LEIBEE' AND JOHN L. CAPINERA2
'Department of Entomology and Nematology,
University of Florida,
Central Florida Research and Education Center,
Sanford, FL 32771
'Department of Entomology and Nematology,
University of Florida,
Gainesville FL 32611
ABSTRACT
Pesticide resistance in Florida was characterized through a survey and literature
review. 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 impact
of 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, or
household and structural pests. It remains possible to manage most arthropods by us-
ing chemical pesticides, but the current and anticipated lack of efficacious materials
threatens 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 major
insect pests in Florida for which management options have become severely limited
because of insecticide resistance. Both cultural practices (continuous cropping, isola-
tion, transport of infested seedlings) and pesticide use patterns (frequent application
of broad spectrum pesticides) contributed to L. trifolii and P. xylostella resistance de-
velopment. The history of pesticide resistance in these two insects is probably typical
of 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.
RESUME
La resistencia a los pesticides en la Florida fue caracterizada a trav6s de una en-
cuesta y una revision de la literature. La encuesta fue conducida en 1994 entire los en-
tomologos del sector public para determinar el estado present y future, extension,
context, patron e instancias de la resistencia a pesticides insecticidess y acaricidas)
en la Florida. Veinte y cinco ejemplos de resistencia a insecticides y acaricidas fueron
citados por los que respondieron la encuesta sobre plagas agricolas, de ornamentales
y de jardines, de importancia m6dica y veterinaria, o dom6sticas y de otras estructu-
ras. Parece possible manejar la mayoria de los artr6podos usando pesticides quimicos,
pero la falta actual y anticipada de materials amenaza las practices presents en al-
gunas areas. La tendencia en la extension, context, o patrons de resistencia fue
como sigue: cultivos de alto valor, artr6podos frecuentemente tratados, pequefios ar-
tr6podos, y piretroides fueron todos considerados como factors asociados con la resis-
tencia.
Leibee and Capinera: Symposium on Pesticide Resistance 387
La resistenca a los insecticides y su manejo fueron revisados en profundidad para
el minador de las hojas Liriomyza trifolii y para la polilla de la col, Plutella xylostella,
los insects plagas principles en la Florida para los cuales las opciones de manejo se
han tornado severamente limitadas debido a la resisitencia a los insecticides. Tanto
las practices culturales (cosecha continue, aislamiento, transport de plantulas infes-
tadas) como los patrons de uso de pesticides (aplicaci6n frecuente de insecticides de
amplio espectro) contribuyeron al desarrollo de la resistencia de L. trifolii y P. xylos-
tella. La historic de la resistencia a los pesticides en estos dos insects es probable-
mente tipica para la resistencia de las plagas en la Florida, y podria significar
problems futures similares a menos que la dependencia de los pesticides para la su-
presi6n de las plagas sea reducida a trav6s de la adopci6n de filosofia y practices de
MIP.
Insecticide resistance has had an impact on the management of insect pests in
Florida since the mid-1940s following the widespread adoption of synthetic insecti-
cides, especially the organochlorines, organophosphates, and pyrethroids. Numerous
anecdotal reports exist, wherein consistently effective insecticides have become inef-
fective and remained so for several seasons. Such reports have been considered ample
evidence of resistance development (Hoskins & Gordon 1956). In fact, Genung (1957)
provided strong evidence based on anecdotal reports and data from field efficacy trials
for resistance development in the cabbage looper, Trichoplusia ni Hubner, imported
cabbageworm, Artogeia rapae (L.), a Liriomyza sp., and leafhoppers, Empoasca sp., at
a 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 with
laboratory studies in which concentration-mortality response has been used to com-
pare resistant and susceptible strains. Much of this work has been conducted within
the last 10 years and involves species such as cabbage looper (Shelton & Soderlund
1983), diamondback moth, Plutella xylostella (L.), (Leibee & Savage 1992a, Shelton et
al. 1993, Yu & Nguyen 1992), silverleaf whitefly, Bemisia argentifolii Bellows & Per-
ring, (G. L. L. unpublished data), house fly, Musca domestic 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 et
al. 1986), and citrus rust mite, Phyllocoptruta oleivora (Ashmead), (Omoto et al.
1994).
In hopes of providing a better understanding of current pesticide resistance and its
consequences in Florida, we report here the results of a recent survey of public-sector
entomologists conducted to assess the extent of pesticide resistance in Florida, and its
current 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 and
the diamondback moth, for which management options have become extremely lim-
ited because of insecticide resistance.
RESISTANCE SURVEY
During the spring of 1994, 16 public-sector entomologists were sent survey forms
to 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 Florida Entomologist 78(3) September, 1995
sity and USDA entomologists from various backgrounds, representing the fields of
agricultural, ornamental and landscape, medical and veterinary, or household and
structural pest management. Entomologists with considerable field experience, and a
close 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 to
round 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 viewed
to be a critical problem in greenhouses (foliage plants, flowering plants, and some
woody 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 prone
to insecticide resistance problems. Resistance in animal production is a more recent
phenomenon, however, apparently resulting from widespread use of insecticide im-
pregnated ear tags.
Resistance was considered to be a significant, but not critical, problem in vegetable
crops, some field crops, and households. This might be viewed as surprising, because
many vegetable crops, some field crops, and households in Florida receive insecticide
treatments at frequencies similar to the aforementioned situations where pesticide
resistance was judged to be critical. It is likely that the severity of the problem is due
as 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 critical
situation.
For medical pests, which in Florida is principally mosquitoes, the significance of
the resistance problem apparently is related to location. Resistance was reported to be
a 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 high
proportion of the state's population (79%) dwells along the coast, so there is frequent
need 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 them
to avoid development of high pest populations. Such landscape plants often tolerate
considerable defoliation or pest density without obvious symptoms, so chemical treat-
ment is not a regular feature of landscape maintenance. Also, in recent years there
has been a concerted effort to introduce native, hardy, pest-resistant plants into the
landscape, 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 can
present resistance problems in this environment.
Although the number of pests displaying resistance to pesticides has increased
markedly in the last two decades, respondents generally did not see the resistance
problem worsening greatly in the next 10 years. The only exception was the area of
Leibee and Capinera: Symposium on Pesticide Resistance 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 materials
that allow producers to continue with traditional agriculture and pest control prac-
tices despite increasing numbers of pests that have become somewhat resistant to one
or more pesticides. The scientific community has also responded quickly and effec-
tively to the onset of resistance by identifying alternative pest control chemicals and
by 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 responses
were 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 on
whether 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 as
1-10 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 to
5-10 compounds. The range in the number of compounds was given as from 1 product
to 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 crops
were 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 were
those in which frequent or routine pesticide applications were made. Of course this
corresponds to the aforementioned high value crops, but there are also situations in
which 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 favoring
frequent 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 classes
of 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 was
species-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 Florida Entomologist 78(3) September, 1995
distance prone. Similarly, although cross resistance within chemical classes was
acknowledged, respondents indicated that they generally considered each pesticide to
have unique chemical properties, so that development of resistance was difficult to
predict based on chemical taxon. The exception to this generalization seems to be the
pyrethroids, 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 in
Table 1. Surely this is not a complete list, either of pests or problem pesticides, but
serves 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 the
purported cases of resistance are "documented," with the remainder based on anec-
dotal information. However, we carefully selected experienced entomologists and
asked 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 industry
sources and not accessible to us, we have not included this specific information. Note
also 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 resistance
in Florida. A review of insecticide resistance in these two troublesome insects follows.
INSECTICIDE RESISTANCE IN L. TRIFOLII
Past and Present Situation
Prior to 1945, leafminer problems on celery and other vegetables in Florida were
apparently almost nonexistent. Control consisted mainly of clean-up measures and
application 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 naled
which 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 remained
low 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 on
celery 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 celery
was 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 of
leafminer on celery. Permethrin became ineffective for leafminer control on celery in
less than two years. Methamidophos was then considered the only insecticide that
gave 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
Leibee and Capinera: Symposium on Pesticide Resistance 391
TABLE 1. EXAMPLES OF INSECTICIDE AND ACARICIDE RESISTANCE IN FLORIDA CITED BY
RESPONDENTS IN A 1994 SURVEY OF PUBLIC SECTOR ENTOMOLOGISTS.
Date
Arthropod (decade) Pesticide
House fly
Musca domestic (L.)
German cockroach
Blatella germanica (L.)
Cat flea
Ctenocephalides felis
(Bouche)
Horn fly
Haematobia irritans
(L.)
Salt marsh mosquito
Culex nigripalpus
Theobald
Soybean looper
Pseudoplusia includes
(Walker)
Fall armyworm
Spodoptera frugiperda
(J. E. Smith)
Southern green stinkbug
Nezara viridula (L.)
Tobacco budworm
Heliothis virescens
(Fabricius)
Corn earworm
Helicoverpa zea (L.)
Pepper weevil
Anthonomus eugenii
Cano
Beet armyworm
Spodoptera exigua
(Hiibner)
Tomato pinworm
Keiferia
lycopersicella
(Walsingham)
Western flower thrips
Frankliniella
occidentalis
(Pergande)
Mole crickets
Scapteriscus spp.
1940
1950
1970
1980
1950
1960
1970
1980
1950-70
1970-80
1980-90
1980
1950
1960
1990
1970
1970-80
1970
1980
1970
1950-60
1960-70
1990
1980
1970
1980
1990
1980
DDT
chlordane, dieldrin, lindane, malathion
dimethoate, ronnel, tetrachlorvinphos
cyromazine, methomyl, various pyrethroids
chlordane, dieldrin, lindane
allethrin, diazinon, malathion
carbaryl, propoxur
cyfluthrin, cypermethrin
diazinon, malathion
bendiocarb, carbaryl, propoxur
cyfluthrin, fenvalerate, permethrin
fenthion, fenvalerate, flucythrinate,
permethrin, stirophos
DDT
malathion
methoprene
acephate, methomyl, various pyrethroids
malathion, carbaryl, methyl parathion,
diazinon, trichlorfon, fluvalinate, bifenthrin,
tralomethrin
carbaryl, methomyl
endosulfan
ethyl parathion
malathion, diazinon,
ethyl parathion, carbaryl
fenvalerate, permethrin, oxamyl
chlorpyrifos, methomyl
carbaryl
methomyl, fenvalerate
oxamyl
pyrethroids
1970 chlordane
Florida Entomologist 78(3) September, 1995
TABLE 1. (CONT.) EXAMPLES OF INSECTICIDE AND ACARICIDE RESISTANCE IN FLORIDA
CITED BY RESPONDENTS IN A 1994 SURVEY OF PUBLIC SECTOR ENTOMOLOGISTS.
Date
Arthropod (decade) Pesticide
Green peach aphid 1970-80 malathion, diazinon, oxydemeton-methyl,
Myzus persicae (Sulzer) dimethoate
Cabbage looper 1950-60 DDT, toxaphene, parathion
Trichoplusia ni 1960-70 endrin, mevinphos, naled
(Htibner) 1970-80 methomyl
Cowpea curculio 1980 endosulfan
Chalcodermus aeneus
Boheman
Citrus rust mite 1990 dicofol
Phyllocoptruta oleivora
(Ashmead)
Yellow pecan aphids 1980 various pyrethroids
Monellia caryella
(Fitch), Monelliopis
pecanis Bissell
Silverleaf whitefly 1990 bifenthrin, fenvalerate, permethrin,
Bemisia argentifolii endosulfan
Bellows & Perring
Two-spotted spider mite 1980 fenbutatin-oxide
Tetranychus urticae 1990 avermectin
Koch
Melon aphid 1990 acephate
Aphis gossypii Glover
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 an
unusual lack of efficacy occurred in the Everglades area.
Laboratory studies confirmed the presence of a high level of cyromazine resistance
in a suspect strain ofL. trifolii (G. L. L., unpublished data). Larval mortality in the
resistant strain at 300 ppm of cyromazine, the highest label concentration used in the
field, 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 suggested
that the resistance was conferred by a major gene. The resistance was considered un-
stable since sensitivity returned in the resistant strain (from an LC0, of about 440
ppm to an LC0, of about 85 ppm) within 5 generations of laboratory rearing without
selection. This was consistent with a survey of leafminer populations that indicated
susceptibility to cyromazine had returned during the summer of 1990 (J. S. Ferguson,
unpublished data). This reversion was probably due to the immigration of susceptible
individuals during what is traditionally a period of little or no celery production and
very 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 leafminer
in celery. This information contributed to the granting of a crisis exemption (Section
18, 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-
Leibee and Capinera: Symposium on Pesticide Resistance 393
tion for the use of abamectin in celery. Since then, abamectin has been used in celery
under specific exemptions. These specific exemptions are unique in that, in order to
discourage the onset of resistance to abamectin, only two consecutive applications are
allowed, forcing rotation with another insecticide. However, no other effective insec-
ticide was available for rotation except for cyromazine which, due to reversion, had
become efficacious again. Since cyromazine and abamectin have different modes of ac-
tion and no cross resistance was indicated, cyromazine was included in the leafminer
control 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 and
preserving susceptibility to cyromazine and minimizing the possibility of selecting for
resistance to abamectin. Cyromazine use patterns and celery culture were suggested
that would reduce selection of resistant phenotypes and encourage the immigration of
feral, hopefully susceptible, leafminers into resistant populations.
Recommendations included: using noninfested transplants; initiating the spray
program based on a threshold to reduce the number of insecticide applications; start-
ing with abamectin to maximize early control; rotating two sprays of abamectin with
two sprays of cyromazine to avoid excessive use of one insecticide; finishing a planting
with two applications of abamectin to reduce the number of adults emerging from the
soil and the trash after harvest; disking in trash as soon as possible to remove this
source 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 susceptible
individuals when celery acreage is at its lowest. Except for seedling production, July
and August are otherwise free of celery. Not using cyromazine at all in seedling beds
was recommended, since transplanting from infested seedling beds is considered an
important 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 included
in a complex of cabbage caterpillars along with the cabbage looper and the imported
cabbageworm, 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 hot
water (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 to
expense and the perception of less than desirable control. In the early 1980s growers
Florida Entomologist 78(3) September, 1995
switched to the newly available and extremely effective pyrethroids, permethrin and
fenvalerate for control of the cabbage looper and diamondback moth, both of which
had become difficult to control with the other insecticides.
Insecticide resistance had long been suspected as the cause of the poor cabbage
looper 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 attributed
to 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 have
actually 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 no
longer providing effective control of the diamondback moth. University trials reflected
the 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 moth
at 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 of
resistance 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 growers
to 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 moth
populations have become very difficult to control with any of the registered synthetic
insecticides and Btk.
The presence of Btk resistance in Florida was immediately suspected because Btk
resistance 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 of
Btk-resistance, there were essentially no effective insecticides available for control of
many diamondback moth populations in Florida until the recent introduction of B.
thuringiensis subspecies aizawai (Bta)-based insecticides. Bta-based insecticides
(those possessing the CrylC 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 to
reduced pyrethroid use. Growers are not spraying as frequently for diamondback
moth and are able to use Btk-based insecticides, suggesting a return of susceptibility
to 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 immigration
of feral, hopefully susceptible, individuals into resistant populations. These recom-
mendations were based on the following knowledge. Susceptibility to Btk had been
greatly reduced in some populations. Bacillus thuringiensis resistance in diamond-
Leibee and Capinera: Symposium on Pesticide Resistance 395
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 the
same (G. L. L., unpublished data). Bacillus thuringiensis subspecies aizawai-based
products (those possessing the CrylC toxin) appeared to be effective in populations
where Btk susceptibility was reduced. Tank-mixing Bt with mevinphos was shown to
be 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 most
effective 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 the
parasites and predators providing natural control of the diamondback moth.
Crop culture recommendations included: not growing cabbage in the warmest
months (May through September in central Florida) when insect pressure is the high-
est and Bt-based insecticides are the least efficacious; immediately disposing of crop
residues to prevent migration from heavily selected populations into new plantings
and seedling production areas; and using noninfested transplants, which not only
contributes to control but also reduces spread of diamondback moths to new locations.
Diamondback moths that infest purchased transplants may be highly resistant due to
heavy usage of insecticides on the transplants or in fields near transplant production
areas. Producers growing their own transplants are at an advantage because they
have 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 per
wk) to determine the presence of the pest of concern or unexpected pests; beginning
inspections 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 by
intuition; 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 of
adult activity (Baker et al. 1982) for timing insecticide applications.
Specific insecticide recommendations included: using Btk and Bta as the principle
insecticides 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 to
any one product; using only Bta to insure maximum control if Btk resistance was
known to be present or the status of Btk susceptibility was unknown; applying Bt
twice weekly and tank-mixing with mevinphos weekly, starting with the tank-mix to
maximize the control that is critical early in crop; including endosulfan, chlorpyrifos,
and methamidophos as alternatives or substitutes for mevinphos in the tank-mixes
with Bt, especially endosulfan, since it belongs to a different chemical class than
mevinphos; 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 in
Florida 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 in
some 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
396 Florida Entomologist 78(3) September, 1995
same population being exposed to insecticides continually. In addition, the constant
use of insecticides removes susceptible individuals that may be immigrating into the
population, thus preventing the opportunity for reversion.
These factors are believed to be especially evident with the development of high
levels 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 between
production 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 period
and diamondback moth populations were very low. Crucifer production was thrust
into the warmer, drier parts of the year when diamondback moth became more of a
problem, resulting in increased use of insecticides and subsequent exposure of addi-
tional generations to more selection pressure. With development of the container
grown 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 in
the summer, and be redistributed back to the fields in late summer and early fall.
Therefore, heavy populations of resistant diamondback moth were being perpetuated
locally throughout the year and continually exposed to insecticides. Opportunities for
the 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-grown
transplants were popular and a healthy containerized transplant industry developed
in the south. Much of the transplant production in the south in the winter and spring
supplied more northern growers with transplants to establish stands in the spring
when environmental conditions were not conducive to direct seeding. Transplants
were also produced during summer months in the north to facilitate establishment of
stands 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 that
cause obvious damage and shipping what appeared to be uninfested plants. Adults
flying into the open-sided greenhouses from the field could maintain a supply of eggs
and early mining instars before shipment. In addition, larvae have been observed to
be deep inside the bud of the transplant, out of reach of the insecticide and the eye of
the 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. This
greatly increased the probability of many growers receiving transplants from areas
with resistant diamondback moth populations.
CONCLUSION
The results of the survey attest to the impact of insecticide resistance on past and
present pest management in Florida. The review of resistance problems with L. tri-
folii and diamondback moth illustrates how insecticide resistance can complicate the
management of pestiferous arthropods. Insecticide resistance seems to be pervasive
Leibee and Capinera: Symposium on Pesticide Resistance 397
in Florida, suggesting that we are not adequately considering the consequences of the
way we use pesticides. Resistance management strategies should be integrated with
nonchemical control whenever possible. In the case of the aforementioned L. trifolii
and 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 of
controlling 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. Conserving
pesticide susceptibility takes the development of knowledge and a commitment from
those 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 refine
its 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 nonchemical
techniques could be used to reduce the numbers and frequency of application. This is
best 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 gave
generously of their time to make the survey component of this paper possible: Joe
Funderburk, Jerry Hogsette, Freddie Johnson, Joe Knapp, Phil Koehler, Russ Mizell,
Charlie Morris, Lance Osborne, Jim Price, Dakshina Seal, Dave Schuster, Phil
Stansly, and Simon Yu. We are also grateful to Brett Highland for his critical review
of 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 diamondback
moth (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. Insecticide
resistance 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
398 Florida Entomologist 78(3) September, 1995
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 to
Bacillus 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 leafminer
control 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 resistance
to 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 in
two colonies of Liriomyza trifolii (Diptera: Agromyzidae). J. Econ. Entomol.
83:18-26.
KOEHLER, P. G. 1991. Toxicity of hydramethylnon to laboratory and field strains of
German 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 of
Insecticide Resistance In The 90s. Abbott Laboratories. Sept. 15-17, 1992. Lake
Bluff, 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-Hill
Book 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 methods
for 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 and
management, pp 1-4 in S. L. Poe and J. O. Strandberg [eds.], Plant protection
through 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 and
Sons, Inc., London.
SHELTON, A. M., AND D. M. SODERLUND. 1983. Varying susceptibility to methomyl and
permethrin in widely separated cabbage looper (Lepidoptera: Noctuidae) pop-
ulations within eastern North America. J. Econ. Entomol. 76:987-989.
Leibee and Capinera: Symposium on Pesticide Resistance 399
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 diamondback
moth (Lepidoptera: Plutellidae) to Bacillus thuringiensis subspecies in the
field. J. Econ. Entomol. 86:697-705.
SYED, A. R. 1992. Insecticide resistance in diamondback moth in Malaysia, pp
437-442 in N. S. Talekar [ed.], Proceedings of the second international work-
shop on the management of diamondback moth and other crucifer pests. Asian
Vegetable 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 diamondback
moth, 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. 0. 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 and
Sanford Florida during 1970. Proc. Florida State Hort. Soc. 83:166-167.
Yu, S. J. 1992. Detection and biochemical characterization of insecticide resistance in
fall 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-
81.
4444444444444444444444444444444444444444444444444444
Scott: Symposium on Pesticide Resistance
THE MOLECULAR GENETICS OF RESISTANCE:
RESISTANCE AS A RESPONSE TO STRESS
JULIE A. SCOTT
Knipling-Bushland USDA Livestock Insect Research Laboratory
Agricultural Research Service
Kerrville, TX 78028-9184
ABSTRACT
In this overview of the molecular genetics of resistance, pesticides are regarded as
one of the many environmental stresses against which insects must defend them-
selves to survive. Examined at the genetic level, pesticide resistance appears to be a
preadapted 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 and
a 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 the
development of novel methods to detect and monitor for resistance is examined and
compared to more traditional technology.
Florida Entomologist 78(3) September, 1995
Key Words: Pesticide resistance, molecular, mutation, genetic, adaptation, stress, in-
sect.
RESUME
En esta revision de la genetica molecular de la resistencia, los pesticides son con-
siderados como uno de los muchos estr6ses ambientales de los cuales los insects de-
ben defenders para sobrevivir. Examinada a nivel genetico, la resistencia a los
pesticides parece ser una respuesta preadaptada al estres y no debida a nuevas mu-
taciones causadas por la exposici6n a los pesticides. Son descritas y explicadas las mu-
taciones geneticas-amplificaci6n genetica, regulaci6n g6nica alterada, alteracion
structural de un gen-que produce la resistencia, y es considerado un possible meca-
nismo de distribuci6n de los genes de resistencia. Son detallados los mecanismos de
resistencia, los process biol6gicos y los tipos de mutaciones asociados con cada uno de
ellos. Finalmente, es examinado y comparado a la tecnologia mas traditional el poten-
cial de la tecnologia molecular para el desarrollo de nuevos m6todos de detecccion y
monitoreo de la resistencia.
When we discuss pesticide resistance, we generally refer to our ability to control a
pest, 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 which
must 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 and
nonlethal stresses for as long as they have existed. And, they have evolved effective
defense mechanisms to deal with these stresses. Their potential to adapt and develop
resistance to stress becomes most apparent when examined at the molecular genetic
level. 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 that
resistance 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 readers
to concepts of molecular genetics and pesticide resistance. It is not a technical review
of the molecular biology of specific resistance mechanisms.
STRESS, RESISTANCE AND TOLERANCE: DEFINITIONS AND INTERACTIONS
Before discussing the molecular genetics of resistance, some definition and discus-
sion of the relationships of stress, resistance, and tolerance is required. Stress has
been broadly defined as "any environmental change that acts to reduce the fitness of
an organism" (Koehn & Bayne 1989). Stresses may have physical, biotic, and/or toxic
components (Fig. 1) which, in turn, may be acute, chronic, and/or seasonal and affect
insects at the community, population, and/or individual level. The deleterious effects
of excesses in temperature or humidity are self-evident. Exposure to ultraviolet radi-
ation can influence trophic-level interactions of entire communities to the advantage
of 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 or
failure of a population to occupy a particular ecological niche. Toxic components of the
environment are also stresses that can affect populations and can be divided into
three groups: pollutants, which may be natural or artificial, pesticides, and plant al-
Scott: Symposium on Pesticide Resistance
predation/
parasitism
disease
plant
allelochemicals
Toxicants
Figure 1. Examples of environmental stresses which can act on living organisms
and force individuals, populations, and entire communities to continuously adapt to
new and ever changing conditions in order to survive.
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 (Anonymous
1957). 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 different
events. 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 a
population'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 to
amounts of toxicant which are sublethal to the pre-adapted individuals but may or
may not be sublethal to others in the population. For many years this adaptive event
was difficult to understand; however, we now know that the toxic components of some
pesticides 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 that
detoxify pesticides. Therefore insects which can detoxify certain plant allelochemicals
well are pre-adapted to detoxify and develop resistance to pesticides which have the
same mode of action as the allelochemicals even before the insects are ever exposed
to the pesticides.
Resistance and tolerance are often used interchangeably in the literature and to
define one another; however, tolerance is also used to describe shifts in susceptibility
that occur within a single generation after exposure to stress which is not expressed
by succeeding generations until after exposure to a similar stress. This phenomenon
is different from the one previously described. For example 6-day-old bollworms, He-
Physil
humidity
radiation
pollutants
pesticides
Florida Entomologist 78(3) September, 1995
liothis zea (Boddie), only detoxify relatively small amounts of methyl parathion, but
at 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 detoxify
not 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 exposure
to gossypol. The ability to metabolize methyl parathion is inherent; the increase in
tolerance to it is not. The difference between resistance and tolerance can become
blurred when a population is subjected to a strong selection pressure, such as chronic
pesticide exposure, and the mechanism that is induced by the pesticide exposure to
yield 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 is
heritable 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 of
the resistance gene(s) which may or may not be heritable.
Two other terms which are often used and may be confused with one another are
cross-resistance and multi-resistance. Cross-resistance is resistance to two or more
classes 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 the
coexistence of two or more different resistance mechanisms. For example, a resistant
insect may have both metabolic resistance to organophosphates and target-site resis-
tance to pyrethroids.
THE WAYS AND MEANS OF ALTERING GENETIC MATERIAL
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 same
detoxification mechanism. It does not explain resistance mechanisms which do not
appear 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 parental
generation to stress result in their progeny being more resistant to that stress? The
answer 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 occur
and 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 insect
has ten copies of a gene, then it can make ten times as much product as an insect with
only one copy of the gene. If the amplified gene encodes for a detoxifying enzyme, then
that 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, there
is only one copy of the gene present in the mutated insect but that gene's regulation
is 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 but
in 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 its
toxic form, up-regulation of a detoxifying enzyme will increase resistance. When a
pro-insecticide, i.e. the material must be metabolized first in order to become toxic, is
applied, down-regulation of the metabolizing enzyme will increase resistance.
Scott: Symposium on Pesticide Resistance
(a) (b) (c)
Amplification Altered Structural
Expression Change
EZ oWI Q
DOD 000 0
Figure 2. Graphic representation of the types of genetic mutations which occur
and 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 regulatory
expression of a gene is altered to increase the amount of gene product made (note that
gene expression may also be altered to decrease the amount of product made) (c) the
genetic code is rewritten to produce a structurally different product.
The third type of mutation that can result in resistance is a structural change in
a gene which yields a corresponding structural change in its product. A single point
mutation, i.e. one nucleotide in the gene's coding region is substituted with a different
nucleotide so that a different amino acid is encoded for at a specific position and this
change causes the gene product to have a different three-dimensional structure, can
result in resistance in several ways. It may decrease the ability of the insecticide to
physically bind to its site of action, or increase or decrease the gene product's ability
to metabolize the insecticide. A structural change does not alter the quantity of the
product made but alters the quality of the product made.
It is important to recognize that these alterations to an insect's DNA do not create
new genes. They only affect pre-existing genes. The idea that exposure to a pesticide
causes resistance genes to be "created" has been debated periodically; however, no
substantiating data have ever been proffered. It is much more likely that resistance
genes 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 may
also 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 more
copies of the TE are inserted into the genome) after they have moved. TEs can also
move genes that were previously not mobile and whose functions are not related to
the transposition. The genes that are moved with the TEs are also replicated at a
higher frequency (Berg 1989). Gene amplifications may be initially distributed
throughout a population by TEs or a gene may be transposed to a new location where
its 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 the
gene Met results in insecticide resistance in Drosophila (Wilson & Turner 1992). But
Florida Entomologist 78(3) September, 1995
so far there is absolutely no evidence that such an event has occurred in a field popu-
lation.
RESISTANCE MECHANISMS AND ASSOCIATED TYPES OF MUTATIONS
Interestingly, only specific types of genetic mutations appear to be associated with
specific resistance mechanisms (Table 1). The two types of mechanisms that cause
high 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 to
alter 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 that
several 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 for
the 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 structurally
altered 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
TABLE 1. THE GENETIC MUTATIONS ASSOCIATED WITH ENZYMES AND RECEPTORS THAT
RESULT IN DIFFERENT TYPES OF RESISTANCE.
Associated Genetic Mutations
Types of Gene Altered Structural
Resistance Amplification Expression Change
Metabolic
P450 oxidases ND' + ND
Esterases + ND +
GSTs ND + ?
Target site insensitivity
Acetylcholinesterase ND ND +
GABA receptor ND ND +
Sodium channel ND ? ?
JH receptor ND ? ?
Other
Reduced penetration
Behavioral change
'ND = not detected; + = confirmed or strongly indicated; ? = implied but not confirmed;
= no data available.
Scott: Symposium on Pesticide Resistance
altered to yield resistance, then susceptible and resistant strains will have quantita-
tively different amounts of that P450. Three P450s have been demonstrated to be
over-expressed by resistant insect strains: P450Lpr (Wheelock & Scott 1992),
CYP6A1 (Carifio et al. 1994), and CYP6A2 (Waters et al. 1992), respectively. P450Lpr
has been directly implicated as the major enzyme causing pyrethroid resistance in
one 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 to
determining which P450s are involved in resistance is our inability to distinguish the
activities of individual uncharacterized P450s from each other with a high degree of
accuracy. The activity of a specific P450 towards an insecticide must be determined
because 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 substrates
are 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. Since
many 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 either
gene amplification or altered gene expression. So far, no data have indicated that the
expression 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 mutation
cannot be discounted.
Considerable data show that certain non-specific esterase genes are amplified to
yield resistance. The esterases which cause resistance in Myzus persicae Sulz. and
Culex mosquitoes have been particularly well-studied. In these insects the resistant
esterase genes are highly amplified and up to 250 copies of the same gene may be
found in a single individual (Mouch6s et al. 1986, Poirie et al. 1992). The more the es-
terase genes are amplified, the greater the level of resistance that they provide (Field
et al. 1988, Poirie 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 hydrolyze
the insecticides (Devonshire & Moore 1982, Ketterman et al. 1992, Karunaratne et al.
1993).
How recently these esterases have been amplified in Culex populations has been a
matter of some debate. One group has suggested that two esterases associated with
resistance, 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 B2
have 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 be
identical. 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 were
present in a number of resistant strains (Ketterman at al. 1993), and three amino acid
differences 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
Florida Entomologist 78(3) September, 1995
once long ago and have since diverged. Examination of other amplified esterase genes
from different Culex strains clearly reveals numerous differences at the molecular
level and strongly suggests that multiple amplification events have occurred with
them (Vaughan et al. 1995).
In addition to amplification, esterases maybe 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, a
carboxylesterase appears to be structurally altered in resistant populations to pro-
duce high levels of resistance to malathion (Whyard et al. 1994a, 1994b). In neither
species is more of the enzyme produced. The difference between susceptible and resis-
tant populations is strictly a qualitative difference in the enzyme produced. Whether
or not the carboxylesterases from the blow fly and mosquito are homologous to each
other 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 molecular
level in insects than the P450s and esterases. Elevated GST levels are found in many
resistant insect strains (Motoyama & Dauterman 1975, Ottea & Plapp 1984, Aham-
mad-Sahib et al. 1994, Hoffman & Fisher 1994) and increased GST activity is clearly
the underlying resistance mechanism in some cases (Kao & Sun 1991, Wang et al.
1991, Prapanthadara et al. 1993). But in other resistant populations the increased
GST 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 GST
production (Yu 1992a, Lagadic et al. 1993, Leszczynski et al. 1994), and generalist
plant feeders seem to rely on GST pathways more heavily than do specialist feeders
to metabolize plant allelochemicals (Yu 1992b). Because GSTs can metabolize a wide
variety 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 GST
activity does not appear to be the result of gene amplification. In both the house fly
and Drosophila, several GSTs contribute to resistance and their expression appears to
have 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 GSTs
have been found in several insects, each of which appears to be encoded by a different
gene (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 changes
and is not due to gene amplification in most, if not all, cases.
The second major resistance mechanism is target-site insensitivity, which refers to
an alteration of the molecules) 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 putative
targets 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 the
neurotransmitter acetylcholine. Multiple forms of AChE that confer varying degrees
of 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
Scott: Symposium on Pesticide Resistance
ofAChE for the pesticide has been reduced. Neither gene amplification nor altered ex-
pression of the gene encoding AChE has been detected. Instead, point mutations have
occurred to structurally change the enzyme. Recently, Mutero et al. (1994) identified
five 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 them
were combined, high levels of resistance resulted. Mutero et al. (1994) hypothesize
that 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 single
mutation 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 inferred
that 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, decreased
sensitivity by GABA receptors is due to a structural change of the protein. Neither
amplification 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 be
substituted with another results in high levels of resistance to cyclodienes. Other
point mutations have been detected, but no others appear to cause resistance or are
consistently associated with resistance (ffrench-Constant et al. 1993, Thompson et al
1993).
Voltage-gated sodium channels play an integral role in the transmission of neural
impulses. Pyrethroids disrupt neural transmissions by interrupting the normal func-
tioning of voltage-gated sodium channels. Target-site insensitivity to pyrethroids, a
phenotypic response commonly referred to as knockdown resistance (kdr), results in
these sodium channels becoming less sensitive to intoxication. Although it is clear
that sodium channels are adversely affected by pyrethroids, there is disagreement as
to whether or not kdr is the result of a mutation to the sodium channels or to some
other molecule which is integral to the functioning of the sodium channels. A point
mutation 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 shown
to cause resistance. Other reports indicate that the mutation which causes kdr is
closely linked to (physically close to or a part of) the gene encoding one type of sodium
channel (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 associated
with kdr. On the other hand, a mutation to a regulatory protein or receptor could also
result in kdr. There is some indirect evidence to support this alternative hypothesis
(Rossignol 1991, Osborne & Pepper 1992). A third hypothesis that has been proposed
and for which there is limited electrophysiological evidence is that kdr is caused by
changes to two closely linked genes. One involves an altered sodium channel and the
other may be associated with calcium-activated phosphorylation of a protein(s) in-
volved with neurotransmitter release (Pepper & Osborne 1993). At this time, it can
only be stated that there is no indication that a gene amplification event is associated
with kdr. It is most likely that either the expression of a gene associated with sodium
channel function has been altered and/or a structural mutation to sodium channels
results in kdr.
Juvenile hormone analogs such as methoprene compete with the natural hormone
for 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 target
site insensitivity to JH analogs are in mutants isolated from laboratory colonies of
Florida Entomologist 78(3) September, 1995
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 expression
of the insect growth cycle has been altered. Gene amplification of JH receptors has not
been 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 tactile
or olfactory receptors, with a pesticide. Alone, neither of these mechanisms cause high
levels 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 by
an insect. For example, Raymond et al. (1989) calculated that reduced penetration in-
teracts with any other resistance mechanism multiplicatively. Experimental data
support this conclusion (Hoyer & Plapp 1968, Plapp & Hoyer 1968). The underlying
physiological, genetic and molecular mechanisms that cause these types of resistance
can only be speculated and the genetic mutations which cause them cannot be in-
ferred at this time.
DETECTING AND MONITORING FOR RESISTANCE
Resistance is a widespread phenomenon and resistant populations of nearly all
economically important pests can now be found (Georghiou 1994, Leibee & Capinera
this issue). Where control failures have occurred, the history of pesticide application
usually indicates that chronic pesticide exposure resulted in high levels of resistance
which caused the failures. Does the development of a resistant population then mean
that a control failure is inevitable? Certainly the continuous selection of the same re-
sistance mechanisms) over and over will result in resistance levels that are high
enough to cause a control failure. But if the selection pressure (i.e. the pesticide) for
each resistance mechanism is removed prior to significant resistance developing, then
a control failure may be avoided (for an alternative view on this subject, see Hoy this
issue). Keys to the successful avoidance of a control failure are the detection of resis-
tance at low levelss, the routine monitoring for changes in the levels) and type(s) of
resistance 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 pest
management 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 types
of resistance and able to detect resistance at frequencies as low as 1%. Numerous
methods to detect resistance are currently available but all fall short of being the ideal
technique. 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 a
limited number of biochemical assays, and novel molecular techniques to detect and
monitor for resistance. Certain general features are shared by the detection assays
within each of these two technological groupings, i.e. traditional and molecular; and
consequently, they have similar advantages and disadvantages which are summa-
rized in Table 2.
Scott: Symposium on Pesticide Resistance
In general, detection methods which utilize molecular technology are better able to
distinguish between the different resistance genotypes, i.e. heterozygotes (SR), ho-
mozygous susceptible (SS) and homozygous resistant (RR), than traditional detection
assays. Because they detect only genetic differences, molecular assays can eliminate
the environmental components which often increase the variability in bioassay and
biochemical 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 to
perform 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 are
limited to the detection of known resistance genes and a separate assay must be done
for 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 detect
many types of resistance at this time since with few exceptions we do not know which
specific genes cause resistance. In time, molecular assays will be developed to detect
more resistance genes; however, it is unlikely that molecular assays will completely
replace 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 of
novel 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. specific
genotypes, that regulate or enhance particular defensive mechanisms. How many
TABLE 2. COMPARISON OF THE ADVANTAGES AND DISADVANTAGES OF DETECTION
METHODS THAT ARE BASED ON EITHER MOLECULAR OR TRADITIONAL TECH-
NOLOGY.
Molecular Detection Assay
can distinguish resistant genotypes
even at low levels
distinguishes SS, SR, and PR
genotypes reliably even when
phenotypes are hard to distinguish
generally greater accuracy and less
variability because environmental
components are eliminated
fewer insects can yield more
information because material from one
insect can be used for several assays
currently not adapted for field use
generally less rapid, it may take days
to complete a test
material and equipment often costly;
more technical expertise often needed
limited to detection of known
resistance genes
vs. Traditional Detection Assay
less sensitive; only detects resistant
phenotypes, not genotypes
cannot distinguish SR and RR from
each other unless the phenotypes are
distinct from one another
less accurate and more variable
because environmental components
cannot be separated from genetic ones
requires more insects to get the same
data because the same individuals
cannot be used in several bioassays
many are easily used in the field
very rapid, often only requiring a
couple of hours to do
generally inexpensive and simple to
prepare and execute
can detect any type of resistance, even
if the resistance gene is not known
Florida Entomologist 78(3) September, 1995
genes can cause resistance is not known. Data on various esterases and cytochrome
P450s 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 distributed
within 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 are
of 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 effective
control 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 relevant
manuscript. 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. BERGE AND D. PAURON. 1992. Target
modification as a molecular mechanism of pyrethroid resistance in Drosophila
melanogaster. Pestic. Biochem. Physiol. 44:183-190.
ANDERSEN, J. F., J. G. UTERMOHLEN AND R. FEYEREISEN. 1994. Expression of house
fly CYP6A1 and NADPH-cytochrome P450 reductase in Escherichia coli and
reconstitution of an insecticide-metabolizing P450 system. Biochemistry
33:2171-2177.
ANONYMOUS. 1957. World Health Expert Committee on Insecticides 7th Report. WHO
Technical 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 of
PCR-based monitoring for cyclodiene resistance in Drosophila populations
with insecticide bioassay. Pestic. Biochem. Physiol. 48:229-233.
BAKER, W. L., A. G. CLARK, G. FAULDS AND J. S. NIELSEN. 1994. Multiple glutathione
S-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 and
R. 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 response
to solar ultraviolet-B radiation: influence of trophic-level interations. Science
265-97-100.
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.
CARINO, F. A., J. F. KOENER, F. W. PLAPP AND R. FEYEREISEN. 1994. Constitutive
over-expression of the cytochrome P450 gene CYP6A1 in a house fly strain with
metabolic resistance to insecticides. Insect Biochem. Molec. Biol. 24:411-418.
Scott: Symposium on Pesticide Resistance
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 Staal. 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 substrate
specificity causes organophosphorus, carbamate and pyrethroid resistance in
peach-potato aphids i.1: 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. CARINO AND A. S. DAGGETT. 1990. Biochemistry
and 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 Insect
Science, 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 resistance
in Drosophila: an invertebrate y-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 y-aminobutyric acid sub-
type A receptor locus is associated with cyclodiene insecticide resistance in
Drosophila 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 .I1 : persicae Sulz.) results from
amplification of an esterase gene. Biochem. J. 251:309-312.
FOURNIER, D., J. M. BRIDE, C. MOUCHES, M. RAYMOND, M. MAGNIN, J-B. BERGE, N.
PASTEUR AND G. P. GEORGHIOU. 1987. Biochemical characterization of the es-
terases Al and B1 associated with organophosphate resistance in Culex pipiens
L. complex. Pestic. Biochem. Physiol. 27:211-217.
FOURNIER, D., J. M. BRIDE, M. POIRIE, J-B. BERGE AND F. W. PLAPP. 1992. Insect glu-
tathione S-transferases: biochemical characteristics of the major forms from
houseflies 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 of
chlorpyrifos 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-derived
population of Chironomus riparius (Diptera: Chironomidae): biochemical and
fitness evidence for population divergence. J. Econ. Entomol. 87:318-325.
Florida Entomologist 78(3) September, 1995
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 an
intensifier 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 by
dimethyl 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 and
characterization of a carboxylesterase involved in insecticide resistance from
the 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 insecticide
resistance. 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. USA
91:2483-2487.
KOEHN, R. K. AND B. L. BAYNE. 1989. Towards a physiological and genetical under-
standing of the energetic of the stress response. Biological Journal of the Lin-
nean Society 37:157-171.
LAGADIC, L., A. CUANY, J. B. BERGE AND M. ECHAUBARD. 1993. Purification and par-
tial characterization of glutathione S-tranferases from insecticide-resistant
and lindane-induced susceptible Spodoptera littoralis (Boisd.) larvae. Insect
Biochem. Mol. Biol. 23:467-474.
LEIBEE, G. L. AND CAPINERA, J. L. 1995. Pesticide resistance in Florida insects limits
management options. Florida Entomol. 78:386-399.
LESZCZYNSKI, B., M. MATOK AND A. F. G. DIXON. 1994. Detoxification of cereal plant
allelochemicals 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.
MOUCHES, C., N. PASTEUR, J. B. BERGE, O. HYRIEN, M. RAYMOND, B. R. DE SAINT VIN-
CENT, M. DE SILVESTRI G. P. GEORGHIOU. 1990. Amplification of an esterase
gene 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. CARINO. 1989. Effects
of cotton allelochemicals on toxicity of insecticides and induction of detoxifying
enzymes 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
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 the
housefly larva (Musca domestica. Pestic. Sci. 39:279-286.
PLAPP, F. W. AND R. F. HOYER 1968. Insecticide resistance in the house fly: decreased
rate of absorption as the mechanism of action of a gene that acts as an intensi-
fier of resistance. J. Econ. Entomol. 61:1298-1303.
POIRIE, 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 pesticide
genes: model and experiment. Genetics 123:543-551.
RAYMOND, M., A. CALLAGHAN, P. FORT AND N. PASTEUR. 1991. Worldwide migration
of amplified insecticide resistance genes in mosquitoes. Nature 350:151-153.
ROSE, R. L., F. GOULD, P. E. LEVI AND E. HODGSON. 1991. Differences in cytochrome
P450 activities in tobacco budworm larvae as influenced by resistance to host
plant 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 P450
genes 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 of
cyclodiene 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-resistant
Culex quinquefasciatus. Biochem. J. 305:651-658.
WANG, J-Y., S. MCCOMMAS AND M. SYVANEN. 1991. Molecular cloning of a glutathione
S-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 involvement
of 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-450
gene 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 house
flies. Pestic. Biochem. Physiol. 43:67-77.
414 Florida Entomologist 78(3) September, 1995
WHYARD, S., R. J. RUSSELL AND V. K. WALKER. 1994a. Insecticide resistance and
malathion 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, a
gene in Drosophila involved in resistance to juvenile hormone analog insect
growth regulators, in C. Mullin and J. G. Scott [eds.], Molecular mechanisms of
insecticide 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.
4444444444444444444444444444444444444444444444444444
Florida Entomologist 78(3)
September, 1995
RESISTANCE: A THREAT TO THE INSECTICIDAL CRYSTAL
PROTEINS OF BACILLUS THURINGIENSIS
LEAH S. BAUER
USDA Forest Service, North Central Forest Experiment Station
Pesticide Research Center & Department of Entomology
Michigan State University
East Lansing, MI 48823-5290
ABSTRACT
Insecticidal crystal proteins (also known as 5-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 enemies
and 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 5-endotoxins has the potential to
greatly 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 5-endotoxin crystals are solubilized in the
midgut; proteases then cleave protoxin molecules into activated toxin which binds to
receptors on the midgut brush border membrane. Part of the toxin molecule inserts
into the membrane causing the midgut cells to leak, swell, and lyse; death results
from bacterial septicemia. Insecticides formulated with Bt account for less than 1% of
the total insecticides used each year worldwide because of high cost, narrow host
range, and comparatively low efficacy. Environmental contamination, food safety con-
cerns, and pest resistance to conventional insecticides have caused a steady increase
in 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 resistance
to 5-endotoxins in P. xylostella populations worldwide. The recent appearance ofBt 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
present in all insects, and the heritability, fitness costs, and stability of the resistance
trait are documented in several insect populations. In two strains ofBt-resistant lep-
idopteran species, mechanisms of resistance involve reductions in the binding of toxin
to midgut receptors. Research on other resistant strains suggests that other mecha-
nisms are also involved. Unfortunately, the high stability of the resistance trait, as
well as broad spectrum cross-resistance to other 5-endotoxins, undermines many po-
tential options for resistance management. Genetically engineered plants, expressing
5-endotoxin continuously and at ultrahigh doses, ensure intense and rapid selection
of the target insect population. The efficacy of transgenic plants can be preserved only
by developing an integrated pest management program that is designed specifically to
reduce selection pressure by minimizing exposure to Bt and increasing other mortal-
ity factors, thereby slowing the rate of pest adaptation to Bt.
Key Words: 5-endotoxin, cross-resistance, transgenic plants, resistance management
RESUME
Las proteinas de cristales insecticides (tambien conocidas como 5-endotoxinas)
sintetizadas por la bacteria Bacillus thuringiensis Berliner (Bt) son el ingredient ac-
tivo de various insecticides ambientalmente amistosos que son 1) altamente compati-
bles con los enemigos naturales y otros organismos no objetos de control debido a su
estrecha especificidad de hospedante, 2) inocuos a los vertebrados, 3) biodegradables
en el ambiente, y 4) altamente d6ciles para la ingenieria genetica. El uso de plants
transg6nicas expesando 5-endotoxinas de Bt tiene la posibilidad de reducir grande-
mente los costs ambientales y de salud asociados con el uso de insecticides conven-
cionales. El modo complejo de acci6n de Bt es sujeto de investigaci6n intensive.
Cuando son ingeridos por un insecto susceptible, los cristales de 5-endotoxina son di-
sueltos en el intestine medio; las proteasas abren las mol6culas de proteinas transfor-
mandolas en toxinas activadas que se unen a receptores en el cepillo de la membrana
del intestine medio. Parte de la mol6cula de la toxina se inserta en la membrana cau-
sando que las c6lulas del intesitno medio pierdan el control de la permeabilidad, se
hinchen y rompan, y la muerte ocurre luego por septicemia bacteriana. Los insectici-
das formulados con Bt son menos del 1% del total de los insecticides usados cada afio
en todo el mundo debido a su alto costo, estrecho rango de hospedantes, y eficacia com-
parativamente baja. La contaminaci6n abiental y la resistencia de las plagas a los in-
secticidas convencionales han causado un incremento constant en la demand de
insecticides de Bt. La reciente escalada de interns commercial en Bt ha provocado la
aparici6n de formulaciones mas persistentes y eficaces. Sin embargo, el uso intensive
de insecticides mejorados a base de Bt autorizados para el manejo de la polilla de la
col, Plutella xyllostella (L.), ha traido como reusltado la evoluci6n de resistencia a las
5-endotoxinas en las poblaciones de P. xyllostella en todo el mundo. La reciente apar-
ci6n de reistencia a Bt en el campo, corroborada por los resultados de experiments de
selecci6n de laboratorio, demuestra que existe resistencia gen6tica en varias species
de Lepidoptera, Diptera y Coleoptera. La capacidad gen6tica de evolucionar la resis-
tencia hacia esas toxinas esta probablemente present en todos los insects, y la he-
redabilidad, costo de ajuste, y estabilidad de la resistencia son documentados en
varias poblaciones de insects. En dos species de lepid6pteros resistentes a Bt, los
mecanismos de resistencia incluyen reducciones en la union de la toxina a los recep-
tores del intestine medio. La investigaci6n sobre otras species de insects resistentes
sugiere que otros mecanismos estan tambi6n relacionados. Desafortunadamente, la
alta estabilidad de la resistencia, asi como la resistencia cruzada de amplio espectro
a otras 5-endotoxinas, determine pocas opciones potenciales para el manejo de la re-
sistencia. Las plants transg6nicas expesando 5-endotoxinas continuamente, y las do-
sis muy altas, aseguran la selecci6n intense y rapida de la poblaci6n de insects a
controlar. La eficacia de las plants transg6nicas puede ser preservada solo desarro-
llando un program de manejo integrado de plagas diseiado especifcamente para re-
Florida Entomologist 78(3) September, 1995
ducir la presi6n de seleccci6n, minimizando la exposici6n a Bt e incrementando otros
factors de mortalidad, para disminuir la velocidad de adaptaci6n de la plaga a Bt.
The bacterium Bacillus thuringiensis Berliner (Bt) is a complex of subspecies char-
acterized by their ability to synthesize crystalline inclusions during sporulation. These
crystalline inclusions are comprised of relatively high quantities of one or more glyco-
proteins known as 5-endotoxins or Cry toxins (Table 1). The toxins produced byBt play
a vital role in the pathogenicity of this bacterium to insects and other invertebrates. The
Cry toxins have enormous commercial value as safe, biodegradable pesticides. The
specificity ofBt 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 ofBt-based in-
secticides provide growers and the public with environmentally friendly and effective
alternatives to conventional insecticides (Meadows 1993). Advances in biotechnology
and genetic engineering, as well as the proteinaceous nature of the Cry toxins, led to the
selection of the cry genes as the primary insect-resistance genes transferred into, and
expressed 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 environments
rich with insects, such as grain bins and insectaries (Smith & Couche 1991, Burges &
Hurst 1977). In fact, the first reports ofBt were from colonies of the silkworm Bombyx
mori (L.) (Ishiwata 1901), and from the Mediterranean flour moth, Ephestia kueh-
TABLE 1. BT STRAINS, THEIR RESPECTIVE 6-ENDOTOXINS AND HOST RANGES, IN INSECT
RESISTANCE STUDIES.
Bt Strain' 5-Endotoxin2 Spectrum
kurstaki HD-1 (Btk) CryIA(a), CryIA(b), CryIA(c), CryIIA, Lepidoptera
CryIIB Diptera3
kurstaki HD-73 CryIA(c) Lepidoptera
aizawai HD-112 (Bta) CryIA(a), CryIA(b), CryIC, CryID, CryIG,
CryII4 Lepidoptera
aizawai HD-133 (Bta) CryIA(a), CryIA(b), CryIC, CryID Lepidoptera
thuringiensis HD-2 CrylA, CryIB Lepidoptera
Coleoptera'
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
RBt strains may contain multiple toxins, and composition may differ slightly from those reported here.
Organized by amino acid sequence by Hofte & Whiteley (1989).
CryIIA accounts for the dipteran activity of this strain.
Presence of CryIG uncertain, and CryII type unknown (McGaughey & Johnson 1994).
CryIB is toxic to some coleopterans (Bradley et al. 1995).
Bauer: Symposium on Pesticide Resistance
niella (Zeller) (Berliner 1911). Ecological considerations of Bt as an insect pathogen
and in the environment are discussed by Meadows (1993).
In the United States, commercial development of Bt into a formulated insecticide
did not begin until the late 1950s. Most Bt-based insecticides are formulated mixtures
of 5-endotoxin crystals and Bt spores, which are known to synergize the toxicity of the
crystals. 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 ofBt subsp. israelensis, which is toxic to larval mosquitoes and black
flies (Goldberg & Margalit 1977), and the discovery ofBt subsp. tenebrionis (Krieg et
al. 1983), which is toxic to several beetle species, stimulated sudden and dramatic
commercial interest in Bt. During the 1980s, new biotech companies and large agro-
chemical and pharmaceutical corporations initiated research programs to isolate Bt
from various environmental samples and to screen for toxicity in agriculturally and
medically-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 species
in three insect orders (Lepidoptera, Diptera, and Coleoptera) to species in eight insect
orders (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), and
coleopteran pests (Keller & Langenbruch 1993). The short half-life ofBt, due to ultra-
violet inactivation when topically applied, has stimulated considerable research into
alternative delivery strategies. By far the most controversial strategy is the use of in-
sect-resistant crops expressing Bt 5-endotoxin genes, which are already in the field in
Asia 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% of
insecticides used worldwide each year. The comparatively high production cost of Bt-
based insecticides is a primary impediment to more widespread usage. However, the
major impetus for greater use of Bt in agriculture is the development of resistance to
conventional insecticides (Georghiou 1994, Watkinson 1994). In fact, many growers
typically add Bt to conventional sprays because of concerns about chemical control
failure (Marrone & Macintosh 1993). Because of their environmental safety, microbial
insecticides are one of the few pesticides that can be developed and registered quickly
and cheaply. In addition, resistance to conventional insecticides does not confer cross-
resistance to Bt toxins due to the unique mode of action of 5-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 can
no longer be controlled by one or more pesticides (Georgiou & Lagunes 1988). Resis-
tance is documented in diverse groups of insecticides, including neurotoxins, chitin
synthesis inhibitors, juvenile hormone analogues (National Research Council 1986),
and, most recently, Bt (Tabashnik et al. 1990). Adaptation to individual insecticides is
a 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-
Florida Entomologist 78(3) September, 1995
resistant plants will expedite selection for resistance in the target organisms more
rapidly than has occurred through conventional application methods (Gould 1988a,
1988b, Van Rie 1991, McGaughey & Whalon 1992, Marrone & Macintosh 1993, May
1993, Whalon & McGaughey 1993, McGaughey 1994, Tabashnik 1994a).
Our understanding of the nature of Bt insecticidal crystal proteins has advanced
rapidly 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 an
abundance of new research results, methodologies, and discoveries. In this paper, my
objectives are to provide researchers, even those only peripherally involved or knowl-
edgeable of Bt 5-endotoxins, with a greater understanding of 1) Bt mode of action, 2)
resistance and cross-resistance to Cry toxins, and 3) resistance management, as they
relate to the use of both improved Bt insecticides and genetically-engineered plants
expressing 5-endotoxin genes. The research results presented here will compliment
and 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 of
IPM and management of Bt resistance. The design, validation, and implementation of
effective 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 a
susceptible host reveals various levels at which resistance might evolve in an insect
population (Fig. 1). The high degree of host specificity, as well as the complexity ofBt
mode 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 the
midgut was the primary site of 5-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, Honee & Visser 1993, Knowles & Dow 1993, Yamamoto & Powell 1993,
Federici 1993, Visser et al. 1993). A small family of Bt 5-endotoxins known as the cy-
tolytic or Cyt toxins, an important component ofBt subsp. israelensis (Table 1), is not
covered 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 5-endotoxin. Detection
and behavioral avoidance of food treated with Cry toxins have also been reported for
many 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 after
treatment.
Crystal Solubilization
Following ingestion, solubilization of crystalline 5-endotoxin is a prerequisite to all
subsequent events in the intoxication pathway (Tojo & Aizawai 1983, Du et al. 1994).
Bauer: Symposium on Pesticide Resistance
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 extent
of crystal solubilization greatly influence toxicity levels in different hosts (Aronson et
al. 1991, Bradley et al. 1995), as well as the rate of intoxication (Bauer & Pankratz
1992, Koller et al. 1992).
Enzymatic Processing
The Cry proteins are synthesized as protoxins that require processing by midgut
enzymes to generate activated toxin (Ogiwara et al. 1992). The larger protoxins of
about 130-140 kDa (e.g. Cryl) are proteolytically cleaved, exposing the activated toxin
which is a protease-resistant core of about 55-65 kDa (H6fte & Whiteley 1989). Many
other toxins (e.g. CryII, CryIII, CryIVD) are synthesized as 70 kDa proteins and are
similar to the N-terminal half of the larger protoxins. Enzymatic processing of these
smaller 70 kDa toxins also occurs, with amino acids cleaved from the N terminus
(Carroll et al. 1989).
Receptor Binding
The action of Cry 5-endotoxins on the midgut epithelium begins with binding of
the 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 brush
border membrane, although most toxins bind to more than one receptor (Van Rie et al.
1989, 1990a, Denolfet al. 1993, Estada & Ferr6 1994). Amino acid sequence similarity
in the receptor-binding domain of the toxin molecule is a useful predictor of overall
host specificity (Van Rie et al. 1990, Cummings & Ellar 1994). The binding domain is
also 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 most
animals (Ellar 1994, Garczinski & Adang in press).
Binding, while essential, is not sufficient to produce mortal damage, as shown by
several 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,
Ferr6 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 (Wolfersberger
1991) or membrane permeability (Carroll & Ellar 1993). They stressed that, although
receptors play an essential role, post-binding factors are required for successful intox-
ication by Bt 5-endotoxins.
Intercalation, Pore Formation, and Cell Lysis
After binding to a receptor on the cell surface, the toxin then inserts or intercalates
into the plasma membrane (English & Slatin 1992, Knowles & Dow 1993). Evidence
from 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)
420 Florida Entomologist 78(3) September, 1995
Ingestion
v
Solubilization
v
Processing
v
Binding
v
Intercalation
v
Pore Formation
v
Cell Lysis
v
Septicemia
Death?
Figure 1. Generalized flow chart of events leading to Bt intoxication in a suscepti-
ble host.
Bauer: Symposium on Pesticide Resistance
also detected toxin-toxin aggregation, supporting evidence that the toxin acts as a
multimer on the cell surface. Pores in the plasma membrane are estimated at 1-2 nm
diam, disrupting the actively maintained osmotic balance, causing the cells to swell
and burst by a process known as colloid-osmotic lysis (Knowles & Ellar 1987). Carroll
& Ellar (1993) demonstrated that 5-endotoxin-induced changes in cell permeability is
non-selective to cations, anions, neutral solutes, and water. A three-dimensional
model of the CryIIIA protein structure supports the hypothesis that the toxin causes
pores 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 5-endotoxin, on insect mor-
tality leads to speculation that 5-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 to
gain access to the body cavity. The insect hemolymph provides an excellent medium
for bacterial growth. Death caused by bacterial septicemia usually occurs within 2-3
days 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 capability
of midgut epithelial cells, can allow damaged regions of the midgut to heal. The actual
recovery 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 et
al. 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 suggest
otherwise. The slow development of field resistance in the past may have resulted
from low selection pressure exerted by early formulations and usage patterns (Stone
et al. 1991). Other researchers believed that the complex mode of action of Bt, often
involving multiple toxins and Bt spores, provided protection against resistance be-
cause a single mutation in the insect would be unlikely to affect susceptibility (Boman
1981, Briese 1981, de Barjac 1987). McGaughey & Whalon (1992), however, suggested
that at high levels of selection, the multicomponent-toxicity pathway merely expands
behavioral 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 biopesticide
potency 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 (Hiibner), the Indianmeal moth, and up to a 15-fold difference in Cadra
cautella (Walker), the almond moth. However, the underlying cause of this variation
Florida Entomologist 78(3) September, 1995
is unknown because the history of Bt applications in these grain storage facilities was
not reported. In a subsequent study over a five-state area, the mean median lethal
concentration (LC,,) for populations ofP. interpunctella from grain bins treated with
Bt was 1.2-fold higher than the mean LC0, 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 15
generations. 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). In
addition, the dark, stable and closed environment of grain bins favored selection for
resistance by long toxin residual times because of no UV exposure and by minimizing
the 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. Resistant
populations ofP. 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, Ferr6 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 that
these unique bacterial toxins may be rendered useless as pest management tools, par-
ticularly with the imminent commercialization of transgenic plants expressing single
activated toxic fragments (May 1993, Whalon & McGaughey 1993).
Laboratory Selections
The increased effort researchers are now devoting to Bt resistance is reflected in
the many experiments designed to select for Bt resistance in the laboratory (Table 2).
The results summarized in Table 2 represent selection experiments, performed and
ongoing during the last decade, which achieved significant levels of resistance to Bt
preparations containing either a mixture of spores and native 5-endotoxin crystals, or
individual Cry toxins in various forms. These results represent only a few of the more
than 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 resistance
ratios ranging from 1.1 to >1000.
Concern about rapid adaptation of insects to 5-endotoxin-based transgenic plants
has 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 & Ferr6 1994). Researchers found that resistance ratios were consistently
higher, and resistance developed more rapidly, in insect populations selected with in-
dividual toxins than in populations selected with Bt-insecticides that contain live
spores and multi-toxin crystals, such as Bt subsp. kurstaki (Btk) (Tables 1 and 2). The
results of these experiments suggest that deployment of the high dose, single toxin
strategy in the design of transgenic plants will quickly generate resistant populations
of 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 from
either Btk spore/crystal preparations or purified CryIC toxin (Table 2). After 20 gen-
Bauer: Symposium on Pesticide Resistance
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-fold
resistant after 21 generations. This confirms that the complex mode of action, present
in most commercially Bt-based insecticides available today, is important in the low
prevalence of field resistance for pest populations treated with Bt. It may also explain
the lack ofBt-resistance generated in laboratory experiments which used crude or for-
mulated Bt preparations containing both 5-endotoxins and spores for selections
(Tabashnik 1994a). Before 1985, large quantities of a single, pure Cry toxin, made
possible 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 populations
are 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 5-endotoxin is
widespread in insects. Although some efforts to select for resistance have failed, this
may reflect either insufficient selection pressure, a genetic bottleneck due to lack or
loss 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 that
is caused by additive genetic variation (Falconer 1989). Estimated heritability was
used 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 relatively
high 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 the
heritability estimates in assessing resistance risk, i.e., predicting the rate at which a
pest will evolve resistance (Tabashnik 1992, Keiding 1986, Tabashnik & McGaughey
1994). Plodia interpunctella has a high estimated h2 and adapts readily to Bt in the
laboratory, 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 do
not achieve significant levels of resistance to Bt in laboratory selections (Devriendt &
Martouret 1976, Krieg & Langenbruch 1981). However, moderately resistant field
populations of P. xylostella quickly reach high levels of resistance during laboratory
selections (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 in
populations of mosquitoes (Gill et al. 1992), P. interpunctella (McGaughey & Johnson
1992), 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-
Florida Entomologist 78(3)
a)
c0
C)
C0
C t
-^-~a
bfl b
2
cPc
C)
Cta
3t c c c (
C) acc. cc
3 m 3 c
s' ct! .. a c
^c~c0f ccjn n-^^
0
o i c o
gto
0
Arr
c)
(U
'V
C
Cl Ct Ct Ct Cte -ic' O- -I(=cl 't
SC to q I A cl cc OO
rm
c)
il
m
i m
.S
C)
C-
oa>
tr
'"
cc 0
'cc A-
0cc
C
ctC
^0
z
'Vy
Cu
CD
)a
cc
-EVj
U
Y
September, 1995
S
cs
; 3
o
-s '
&F a
Sg -I **
a S c a
S
F4m
IF I
ai
-c- m o.
_a ,M ^
PI i
d -3
a- : 2
I I IU I ac
o No
0 o 0
; 3L F 1 ,
*h q, a 0 3 -
I I'i Ig
S
5 q, 0
c t >iB a-
a I I l *a 1
*g ~ S" ^-o? "" S
^~ .a g " -
a)
d~j CC
0 0
abb~~bb
Bauer: Symposium on Pesticide Resistance
to
a)
a)
tC)
a)
C)
NE
'C
ccc
O o
g
A A
rt _
-S w
oo
A A
t &1 M&LO
- 00 COD fM COD CO
cc
==
,,
uu
.s
s .,a
de
F1
-"f~
u?
a)
C C a
00
C)
o M
m
Sm
QIcnQ
t~oto
1^^
'S '73 '
";3
I
I .F
-s 0
00
0c 0
I a0
0 ~C
Sg *I **
S a -.-& a
a
"s
.^ i I l
cCe
_a M
B o
S3
>- I 0
a 0 q,' q,
S- 0 (- r q,
-h q, a .3 s
II a
ct ,s
m 0 >iB^ a-
a~ gl ^ & g
^~ .a a *"
,tr ,: : 1 1 ;
.a- .s s a .
*g' o a | g .' .
a a^'a^ ^^'a^
&si&S^S&SS3
c 00
a
cq
a
g
<-0
S
o 09
ss
Florida Entomologist 78(3) September, 1995
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 species
within 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 withBt 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 that
the susceptibility of intensively treated populations varied up to 40-fold, whereas the
susceptibility 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 in
North 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 among
nine spruce budworm field populations from Ontario with no previous Bt exposure re-
veal substantial familial differences in sensitivity to Bt, whereas differences between
populations are minimal. This finding suggests that Bt tolerance is genetically based,
and that spruce budworm has the potential to adapt to Bt. One laboratory selection
experiment (Table 2) illustrates an increase in frequency of the resistance trait (van
Frankenhuyzen et al. 1995).
To gain baseline data on the Bt susceptibility of two pests targeted by transgenic
cotton, Heliothis virescens (F.) and Helicoverpa zea (Boddie), Stone & Sims (1993) bio-
assayed populations from 14 states with activated CryIA(c) toxin and a commercial
Btk-based insecticide. Although previous exposure to Bt was not identified, significant
differences within populations of both species were detected. Moreover, the variability
in Bt tolerance was consistently higher for the activated toxin in both species. This
again 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 and
managing for resistance will be an exciting challenge when insect-resistant cotton, ex-
pressing Bt 5-endotoxin, is deployed.
Mechanisms
An understanding of the mechanisms of resistance to Bt toxins will prove essential
in the future design and management of transgenic plants containing Bt toxin genes.
At present, the primary mechanism of resistance reported for P. interpunctella (Van
Rie et al. 1990b) and P. xylostella (Ferr6 et al. 1991) is a reduction in the binding of
toxin 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 5-endotoxins in Lep-
idoptera (Hoffman et al. 1988b).
Plodia interpunctella, selected for resistance to Btk (see Table 1), shows a 50-fold
reduction 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 at
least 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 one
receptor population may compensate for diminished function in the other. In a strain
Bauer: Symposium on Pesticide Resistance
ofP. 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 field
populations of P. xylostella demonstrated greatly reduced or lack of toxin binding to
midgut receptors, suggesting a change or complete loss of the receptor (Ferr6 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 other
than receptor binding can contribute to resistance. For example, two separate studies
of resistance in H. virescens found no relation between resistance to CryIA(b) or
CryIA(c) and toxin-receptor binding (Macintosh et al. 1991, Gould et al. 1992). In ad-
dition, studies of Trichoplusia ni (Hiibner) 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 Ferr6 1994). Resistance mechanisms
in 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 ofBt 5-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 5-endotoxins are available (Adang 1991), evidence is mounting that selection
for resistance to one or more 5-endotoxins causes resistance to others (Tables 3 and 4).
This phenomenon, known as cross-resistance, typically occurs when mechanisms of
toxicity are similar.
Many of the resistant insect populations generated in the laboratory and the field
were selected with Bt-based insecticides containing multiple toxins; the most studied
products are formulated with Btk strain HD-1, which is comprised of live spores and
a mixture of five toxins (Table 1). In many studies, cross-resistance is referred to as an
increase 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 to
control 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 & Johnson
1987). Resistance was highest among various Btk isolates, suggesting some degree of
specificity. Many Bt isolates overlap considerably in their 5-endotoxin composition. In
the context of the following discussion, cross-resistance is defined as an increase in
the 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 present
in 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, showed
some level of resistance and cross-resistance to six 5-endotoxins tested (McGaughey &
Johnson 1994) (Table 3). Btk tended to select for high levels of resistance to the entire
complex of CrylA toxins which are 82 to 90% homologous in their amino acid se-
quences (H6fte & Whitley 1989). Resistance was highest to CryIA(b) and CryIA(c).
Evidence suggesting that they share the same binding site (Wolfersberger 1990) is
also 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 for
CryIA(b) because cross-resistance is significantly greater than the selected resis-
Florida Entomologist 78(3)
September, 1995
So u 0o 0
CM CM ^-
c-l tol &
M - "I0 O -
tv -~~t
t-- 01 0 Lc~
CM 0M LOM
o N
r- -H &1
c N eq c:
CD toU
&1 1 &
t N^C 0 t^
CD-' --l
10
B
00
.
Fq cc
>m-
sa -
. -F
o
S
II0
-m-~m
B-~ -
000 -B
F1
0
.S
-c
'rV
cccl
a r
C)
acc
Bauer: Symposium on Pesticide Resistance
tance. Cross-resistance to CryIB and CryIC was significant, although very low, which
is consistent with their relatively low amino acid sequence similarity to the CrylA 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 broader
spectrum of cross-resistance in the treated insects than did isolates producing similar
Cry toxins such as Btk HD-1. The complexity in the pattern of resistance results, in
part, from differential selective pressure exerted by different levels of toxicity and the
amounts 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 CryIC
was 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 to
CryIA(b) (resistance ratio 236), showed no resistance to Btk, CryIA(a), CryIA(c) and
no cross-resistance to CryIB, and CryIC (Ferr6 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 narrow
spectrum of resistance is somewhat unique and suggested that this population may
represent a biotype present in the Phillipines, not a case of field selection by Btk.
To understand the complexity of these emerging cross-resistance patterns, we
must 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 into
an 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 of
H. 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 in
the concentration of CryIA(c), or CryIA(b) binding sites or binding affinities, were de-
tected between selected and unselected H. virescens, suggesting other mechanisms of
resistance are involved. Similar broad-spectrum cross-resistance was also reported
for two species of Spodoptera and two coleopterans, Cliry.,,nclo scripta F. (Table 4)
and L. decemlineata (Whalon, unpublished results).
In contrast, cross-resistance in Trichoplusia ni (Htibner) selected with CryIA(b)
had a higher degree of specificity within the CryIA group of toxins (Estada & Ferre
1994). 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 toxins
for 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-resistance
to CryIA(c) was detected, suggesting CryIA(c) toxicity results from other binding sites
or alternative mechanisms.
Overall, cross-resistance patterns and their underlying physiological mechanism
are very complex and somewhat unpredictable, even within a closely related group of
Florida Entomologist 78(3) September, 1995
cc
'V
C)
uC
C
o
'V
b
0 cc
0
0
o.
oW
00 0 00 ~ ,0
0~ ^ 0 1
0 0 g 0 0 0 3 s ^
< ,, -s, a -, ,a ^ l al
.i^ ^^ ^|
cS cS cc
ct ct ct
cI cS cS
cI cI
Fc cFF
cS cS
F0~
I I rt
I rt F1
-d 2
d 0F
"- 0
B -
0 B
-6
Bauer: Symposium on Pesticide Resistance
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 selecting
5-endotoxins for design of transgenic plants that do not favor broad-spectrum cross-
resistance.
Adaptation to 5-Endotoxin
The recent appearance of resistance to conventional Bt-based insecticides was cor-
related with improved formulation and more intensive usage patterns (Tabashnik et
al. 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 rapid
selection (Table 5). These attributes include multivoltinism, short generation time,
and populations that tend to be isolated and stable. These attributes are typical of
many agricultural and medical pests, and they temper enthusiasm for bioengineered
plants which produce continuous and high levels of 5-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 understanding
of the inheritance of a resistance trait in the pest population (Gould 1986). This was
demonstrated by Tabashnik (1994b) using a population genetics model to simulate
the response of a pest population to different resistance management strategies. The
best strategy, i.e., one that delays resistance the longest, must be customized to the
number 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 Two
LEPIDOPTERANS.
Factors P. interpunctella P. xylostella
Pest Attributes and Management
Short generation yes yes
Generations/year 5-6 8-10
Population isolation yes yes
Realized heritability high low
Crop rotation no no
Operational Use of Bt
Multiple toxins yes yes
Applications/generation 5-6 5-8
Frequency/generation 2-4 days 1.5-2 days
Selection pressure high high
Florida Entomologist 78(3) September, 1995
ulations. In both P. interpunctella and P. xylostella, resistance is autosomally
inherited (no maternal effects or sex linkage), partially recessive (progeny suscepti-
bility more similar to unselected parent), and apparently due to one or a few major
loci (McGaughey 1985, McGaughey & Beeman 1988, Hama et al. 1992, Tabashnik et
al. 1992). The genetic basis of resistance to the CrylA toxin-complex in H. virescens is
partially recessive and due to a single locus or set of tightly linked loci (Gould et al. in
press). In L. decemlineata, resistance to CryIIIA is also autosomally inherited, and
conferred by one incompletely dominant gene (Rahardja & Whalon 1995). Knowledge
of the genetic basis of resistance is critical to our understanding of the stability of the
trait within the selected population.
Stability
Perhaps one of the simplest resistance management strategies involves providing
the 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 resistance
trait after Bt exposure ceases. The rate of reversion is dependent on the inheritance
of 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, one
Btk-resistant population declined from 29-fold to 1-fold after 32 generations without
selection (Tabashnik et al. 1991). Similar results were reported in other Btk-resistant
populations of P. xylostella (Tabashnik et al. 1991, 1994, Hamma et al. 1992) and in
other resistant species, including P. interpunctella (McGaughey & Beeman 1988), H.
virescens (Sims & Stone 1991), and L. decemlineata (Rahardja & Whalon 1995). In a
recent study, rapid and complete reversal of resistance occurred after 13 generations
without 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 significant
fitness costs of resistance. Despite the rapid reversion to the susceptible genotype, the
population responded rapidly to reselection (Tabashnik et al. 1994a). Rapid resur-
gence of resistance in relaxed populations is typical, indicating the persistence of a
low number of highly resistant individuals. If the alleles for resistance become fixed
in the population, or other alleles compensate for losses in fitness, resistance becomes
stable and reversion to susceptibility is unlikely.
RESISTANCE MANAGEMENT
In an effort to preserve the utility of these unique insecticidal proteins, knowledge
of 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 in
a pest population is probably the inevitable consequence of insecticide use (Denholm
& Rowland 1992). The goal then becomes how to design and manipulate operational
strategies that best conserve susceptibility, thereby delaying resistance. The imple-
mentation of integrated pest management (IPM) strategies that optimize the goals of
resistance management involves 1) diversifying the sources of mortality to avoid se-
lection for a single mechanism, 2) reducing selection pressure for the major mortality
factors, 3) maintaining susceptible individuals by providing refuges and encouraging
immigration, 4) monitoring for increasing resistance to any one of the mortality
agents, and 5) responding to resistance through management strategies designed to
Bauer: Symposium on Pesticide Resistance
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 IPM
strategies, because the short residual time and host specificity help reduce selection
pressure associated with the Bt toxins. Insects surviving Bt exposure are generally in
a weakened condition, facilitating their exploitation by other mortality factors such as
beneficial insects (Tabashnik 1986, 1994a) and pathogens (Krieg 1971, Jacques &
Morris 1981). Other stressing agents, such as adverse weather conditions and low
plant nutritional quality, will also cause higher mortality in insects recovering from
Bt 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 5-endotoxins in transgenic plants is considered analogous, in
some 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 ofBt 5-endotoxin genes into plants. There was considerable optimism several
years ago that these plants would remain durable (Gould 1986,Gould 1988b, Rousch
1989).
Early optimism and excitement over genetically-engineered plants expressing 5-
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 with
it, conventional uses of Bt. Resistance management strategies in the context of Bt
were 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 plant
or in different plants, or expressed serially over time (Gould 1986); 2) synergists to
increase 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) ultrahigh
doses 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) to
simulate the effect of several resistance management strategies on resistance (single
locus with two alleles) in a pest such as Heliothis. Transgenic plants expressing 5-en-
dotoxin were planted as 1) pure stands of toxic plants, 2) seed mixtures with varying
proportions of toxic and toxin-free plants, 3) toxin-free plants in refugia, and 4) seed
mixtures + 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 because
refugia reduce selection without altering dominance (Mallet & Porter 1992). Refugia
will 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.
Florida Entomologist 78(3) September, 1995
The distinction between seed mixtures and refugia is the spatial distribution of
toxin-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 planted
with a seed mixture, whereas movement from toxic stands to toxic-free refugia is less
likely. The tissue-specific expression of toxin through gene regulation is analogous to
seed mixtures, because larvae can move freely between toxic and toxin-free tissue
(Gould 1988a). Tabashnik (1994b) recommends maximizing spatial refuges, as well as
temporal refuges, such as alternative crops and controls. The only method known to
prolong the efficacy of any insecticide is to minimize the exposure of the target pests
to the toxin in space and time (Denholm & Rowland 1992). Resistance management
tactics 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 5-endotoxins
is now well documented in many species within eight different insect orders. Although
high-level field resistance is known only in P. xylostella, much has been learned from
studying 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 in
different 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 5-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 in
conjunction with conventional insecticides as a backup for control failure, or, as a last
resort once resistance to other registered insecticides has occurred. Many insect pests,
therefore, are already adapted to mixtures of 5-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 not
kept pace with those made in biotechnology. Developing and validating realistic resis-
tance management plans that preserve the durability of 5-endotoxins deployed in
transgenic plants may prove far more complex than the theory and techniques that
actually 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 research
into the development of strategies designed to prolong the durability of these unique
bacterial toxins in transgenic plants. In general, resistance management seeks to
minimize 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 alternating
crops or control measures (Hoy 1995). Substantial benefits to the environment will be
gained if insects can be successfully managed through the careful deployment of ge-
netically-engineered insect-resistant plants. However, only few Bt 5-endotoxins, all
with 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 the
loss of Bt-based insecticides for many of the pests they are targeted to control.
Bauer: Symposium on Pesticide Resistance
ACKNOWLEDGMENTS
An overview of this paper was presented in a symposium entitled "The Myths of
Managing Resistance" at the Florida Entomological Society held 8-11 August 1994 in
Stuart, 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. van
Frankenhuyzen, 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 technical
support 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 for
biological control of pests and vectors. Boca Raton, Florida:CRC Press.
ALY, C., M. S. MULLA, AND B. A. FEDERICI. 1985. Sporulation and toxin production by
Bacillus 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 of
inclusion 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. MENSUA, G. W. RIETHMACHER, AND JUAN FERRE.
Lack of cross-resistance to other Bacillus thuringiensis crystal proteins in a
population 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, C('lir/,rncio scripta, for resistance
to Bacillus thuringiensis var. tenebrionis 5-endotoxin. p. 68 in Abstracts VI' In-
tern. Coll. Invertebr. Pathol.
BECKER, N., AND J. MARGOLIT. 1993. Use ofBacillus thuringiensis israeliensis against
mosquitoes, 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. Ober die Schlaffsucht der Mehlmottenraupe (Ephestia kuehniella
Zell), und ihren Erreger Bacillus thuringiensis n. sp. Zeitschrift fiir 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 diseases
1970-1980. 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.
Florida Entomologist 78(3) September, 1995
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 of
Bacillus 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 in
E. 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 Insects
and Mites. New York:Academic Press.
BURGES, H. D., AND J. A. HURST. 1977. Ecology of Bacillus thuringiensis in storage
moths. 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. New
York:Marcel Dekker.
CARROLL, J., AND D. J. ELLAR. 1993. Analysis of Bacillus thuringiensis 5-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 a
coleopteran-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. de
Barjac and D. Sutherland [eds], Bacterial Control of Mosquitoes and Blackflies:
Biochemistry, Genetics, and Applications ofBacillus thuringiensis and Bacillus
sphaericus. 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 in
wheat cultivars. Environ. Entomol. 15: 24-31.
CUMMINGS, C. E., AND D. J. ELLAR. 1994. Chemical modification ofBacillus thuring-
iensis activated 5-endotoxin and its effect on toxicity and binding to Manduca
sexta midgut membranes. Microbiol. 140: 2737-2747.
DE BARJAC, H. 1987. Operational bacterial insecticides and their potential for future
improvements, 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 in
arthropods: 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 brush
border membrane of the european corn borer, Ostrinia nubilalis (Htibner) (Lep-
idoptera: Pyralidae). Appl. Environ. Microbiol. 59: 1828-1837.
DEVRIENDT, M., AND D. MARTOURET. 1976. Absence de resistance a Bacillus thuring-
iensis chez la teigne des crucif6res, Plutella maculipennis. Entomophaga 21:
189-199.
DU, C., P. A. MARTIN, AND K. W. NICKERSON. 1994. Comparison of disulfide contents
and 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 5-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.
Bauer: Symposium on Pesticide Resistance
ENGLISH, L., AND S. L. SLATIN. 1992. Mode of action of delta-endotoxin from Bacillus
thuringiensis: 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. FERRE. 1994.
Occurrence of three different binding sites for Bacillus thuringiensis 5-endot-
oxins in the midgut brush border membrane of potato tuber moth, Phthorimaea
operculella (Zeller). Arch. Insect Biochem. Physiol. 26: 315-327.
ESTADA, U., AND J. FERRE. 1994. Binding of insecticidal crystal proteins of Bacillus
thuringiensis to the midgut brush border of the cabbage looper, Trichoplusia ni
(Hiibner) (Lepidoptera: Noctuidae), and selection for resistance to one of the
crystal 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.
FERRE, J., M. D. REAL, J. VAN RIE, S. JANSENS, AND M. PERFEROEN. 1991. Resistance
to the Bacillus thuringiensis bioinsecticide in a field population ofPlutella 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) 5-endotoxin
binding 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 5-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 and
other microorganisms as Bacillus thuringiensis delivery systems, pp. 89-104 in
P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs [eds], Bacillus thuringien-
sis, an environmental biopesticide: theory and practice. New York:Wiley, 311
pp.
GEORGHIOU, G. P. 1990. Resistance potential to biopesticides and consideration of
countermeasures, 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 Agricultural
Organization, 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, Culex
univittatus, 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 of
the 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.
Florida Entomologist 78(3) September, 1995
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-73
delta-endotoxin on growth, behavior, and fitness of susceptible and
toxin-adapted strains Heliothis virescens (Lepidoptera: Noctuidae). Environ.
Entomol. 20: 30-38.
GOULD, F., A. ANDERSON, D. LANDIS, AND H. VAN MELLAERT. 1991. Feeding behavior
and 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 and
genetic analysis of a Heliothis virescens (Lepidoptera: Noctuidae) strain with
high 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. Resistance
to 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. Fitness
costs of resistance to Bacillus thuringiensis in the diamondback moth (Plutella
xylostella). Evol. 48: 197-201.
HAMA, H., K. SUZUKI, AND H. TANAKA. 1992. Inheritance and stability of resistance to
Bacillus 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 bacteria
in 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 the
cabbage butterfly (Pieris brassicae). Eur. J. Biochem. 173:85-91.
HOFFMAN, C., H. VANDERBRUGGEN, H. HOFTE, 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.
HOFTE, H., AND H. R. WHITELEY. 1989. Insecticidal crystal proteins of Bacillus thur-
ingiensis. Microbiol. Rev. 53: 242-255.
HONEE, G., AND B. VISSER. 1993. The mode of action of Bacillus thuringiensis crystal
protein. 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 Sanbshi
Kaiho 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:Academic
Press.
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 National
Research 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.
Bauer: Symposium on Pesticide Resistance
Cory, M. J. Bailey, and S. Higgs [eds], Bacillus thuringiensis, an environmental
biopesticide: 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. Bioessays
15: 469-475.
KNOWLES, B. H., AND D. J. ELLAR. 1987. Colloid-osmotic lysis is a general feature of
the 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 the
pH-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 5-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:Academic
Press.
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: Academic
Press.
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 insecticidal
delta-endotoxin from Bacillus thuringiensis at 2.5 A 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 ofHeliothis virescens.
Proc. Natl. Acad. Sci. USA 88: 8930-8933.
MALLET, J., AND P. PORTER. 1992. Preventing insect adaptation to insect-resistant
crops: are seed mixtures or refugia the best strategy? Proc. R. Soc. London B
250: 165-169.
MARRONE, P. G., AND S. C. MACINTOSH. 1993. Resistance to Bacillus thuringiensis
and 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 of
Bacillus thuringiensis isolates. Appl. Environ. Microbiol. 55: 2437-2442.
MASSON, L., A. MAZZA, R. BROUSSEAU, AND B. TABASHNIK. 1995. Kinetics of Bacillus
thuringiensis 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.
Florida Entomologist 78(3) September, 1995
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 in
colonies 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 plant
variation 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 of
5-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 in
Abstracts VIh Intern. Coll. Invertebr. Pathol.
MOLDENKE, A. F., R. E. BERRY, J. C. MILLER, J. G. WERNZ, X. H. LI. 1994. Toxicity of
Bacillus thuringiensis subsp. kurstaki to gypsy moth, Lymantria dispar, fed
with alder or Douglas fir. J. Invertebr. Pathol. 64: 143-145.
MULLER-COHN, J., J. CHAUFAUX, C. BUISSON, N. GILOIS, V. SANCHIS, AND D. LERE-
CLUS. 1994. Spodoptera littoralis resistance to the Bacillus thuringiensis cryIC
toxin and cross-resistance to other toxins. p. 66 in Abstracts VIt Intern. Coll.
Invertebr. Pathol.
NATIONAL RESEARCH COUNCIL. 1986. Pesticide Resistance: strategies and tactics for
management. 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. New
York: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 gut
juices 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 ofPlodia interpunctella. Biochem. Biophys. Res. Comm.
198: 940-947.
PARENT, P, V. MANUELA, G. M. HANOZET, M. TASCA, AND B. GIORDANA. 1995. Inter-
action of the insecticidal crystal protein CryIA from Bacillus thurigiensis with
amino acid transport into brush border membranes from Bombyx mori larval
midgut. J. Invertebr. Pathol. 65: 35-42.
Bauer: Symposium on Pesticide Resistance
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, and
D. 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 5-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 5-
endotoxin gene. Environ. Entomol. 23: 1030-1041.
ROUSCH, R. T. 1989. Designing resistance management programs: how can you
choose? Pestic. Sci. 26: 423-441.
SALAMA, H. S., AND M. M. MATTER 1991. Tolerance level to Bacillus thuringiensis
Berliner 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 susceptibility
levels 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 the
toxicity and receptor binding properties of Bacillus thuringiensis CryIC 6-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 of
Manduca sexta aminopeptidase and phosphatase enhances Bacillus thuring-
iensis insecticidal CryIA(c) toxin binding and "Rb6-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 to
Bacillus 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 the
field. J. Econ. Entomol. 86: 697-705.
SIMS, S. R., AND T. B. STONE. 1991. Genetic basis of tobacco budworm resistance to an
engineered Pseudomonas fluorescens expressing the 5-endotoxin of Bacillus
thuringiensis 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 virescens
and 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 for
resistance to a genetically engineered Pseudomonas fluorescens containing the
delta-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.
Florida Entomologist 78(3) September, 1995
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 Bacillus
thuringiensis 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 in
Plutella 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 thuringiensis
delta-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, an
environmental biopesticide: theory and practice. New York:Wiley.
VAN FRANKENHUYZEN, K., C. W. NYSTROM, AND B. E. TABASHNIK. 1995. Variation in
tolerance to Bacillus thuringiensis among and within populations of the spruce
budworm (Lepidoptera: Tortricidae) in Ontario. J. Econ. Entomol. 88: 97-105.
VAN RIE, J. S. 1991. Insect control with transgenic plants: resistance proof? Trends
Biotechnol. 9: 177-179.
VAN RIE, J., S. JANSENS, H. HOFTE, 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. HOFTE, D. DEGHEELE, AND H. VAN MELLAERT. 1990a. Re-
ceptors on the brush border membrane of the insect midgut as determinants of
the 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. HONEE. 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.
Bauer: Symposium on Pesticide Resistance
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 channel
activity 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 forBt 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, New
York: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 5-endotoxins
to gypsy moth larvae is inversely related to the affinity of binding sites on the
midgut brush border membranes for the toxins. Experientia 46: 475-477.
WOLFERSBERGER, M. G. 1991. Inhibition of potassium-gradient-driven phenylalanine
uptake in larval Lymantria dispar midgut by two Bacillus thuringiensis 5-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 toxicity
between CytA and CryIVD proteins using inclusions produced from cloned
genes 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.
4444444444444444444444444444444444444444444444444444
Hoy: Symposium on Managing Resistance
MULTITACTIC RESISTANCE MANAGEMENT: AN APPROACH
THAT IS LONG OVERDUE?
MARJORIE A. HOY
Department of Entomology and Nematology
University of Florida, Gainesville, FL 32611-0620
ABSTRACT
"Resistance management" tactics have been much discussed, but such tactics have
provided surprisingly limited practical results for pest management programs to
date. We have learned a great deal about pesticide resistance mechanisms, the mode
of inheritance of resistances, the molecular basis of resistance and cross resistance
mechanisms, and how to evaluate the impact of resistance on fitness. However, it re-
mains difficult to "manage" resistance once resistant individuals make up more than
5 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 product
less 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
Florida Entomologist 78(3) September, 1995
economic injury levels so that pesticides are applied only when needed, biological con-
trol, host plant resistance, cultural controls, biorational pest controls, and genetic
control 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 enemies
can be effective components of a program to delay the development of resistance in
pest arthropods.
Key Words: Pesticide resistance, integrated pest management, biological control, pes-
ticide selectivity, resistance management.
RESUME
Las tacticas de manejo de la resistencia han sido muy discutidas pero sorprenden-
temente, hasta la fecha, tales tacticas han dado resultados practices limitados en pro-
gramas de manejo de plagas. Hemos aprendido much sobre los mecanismos de
resistencia, el modo de heredarse, sus bases moleculares y los mecanismos de resis-
tencia cruzada, asi como a evaluar el impact de la resistencia en el ajuste genetico.
Sin embargo, es aun dificil manejar la resistencia cuando los individuos resistentes
integran mas del 5-10% de la poblaci6n. Generalmente, lo mejor que se ha logrado ha
sido retrasar en various afos el desarrollo de altos niveles de resistencia en la pobla-
ci6n, mas a menudo usando menos f recuentemente el product que la provoca. Una
estrategia mas efectiva de manejo de la resisitencia combinaria una variedad de tac-
ticas eficaces de manejo de plagas con la de reducir el numero y dosis de los pesticides
aplicados. Las tacticas efectivas de manejo inculyen el monitoreo, la evaluaci6n del
dano econ6mico de modo que los pesticides sean aplicados solamente cuando es nece-
sario, el control biol6gico, la resistencia de las plants hospedantes, el control cultu-
ral, el control biorracional de plagas, y los m6todos de control genetico. Como parte de
esta estrategia multitactica, es crucial evaluar el impact de los pesticides en los ene-
migos naturales. A veces, los enemigos naturales resistentes pueden ser conponentes
efectivos de un program para retrasar el desarrollo de la resistencia en los artr6po-
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 to
have equivalent goals and methods. When we accept that the goals and tactics are
similar, we will develop effective resistance management programs for arthropods.
Effective management of resistance and effective IPM programs require an holistic
and multitactic strategy. A key component of this holistic and multitactic approach in-
cludes enhancing the I ..... .." of pesticides and biological control agents (Hoy
1992).
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 major
classes of such products (Georghiou & Saito 1983, Georghiou 1986, Roush & Tabash-
nik 1990). Resistance to pesticides in weeds, plant pathogens, and nematodes also is
increasing, although somewhat more slowly (National Academy of Sciences 1986,
Denholm et al. 1992). While my commentary focuses on resistance to insecticides and
acaricides, 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
Hoy: Symposium on Managing Resistance
$60 million per successful compound. Thus, pesticide producers should be increas-
ingly interested in extending the economic life of their products in order to maximize
a 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 in
minor crops, which are increasingly being ignored by pesticide companies because
they are such a small market. These so-called 'minor' crops of fruits and vegetables
are a major component of Florida's agriculture. Registration of new pesticides is likely
to be more difficult and expensive in the future, which could leave some pest manage-
ment specialists with extremely limited options for managing certain recalcitrant
pests.
A few environmentalists have argued that we do not need pesticides, that they will
soon 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. Pesticides
are the most effective tools for fighting outbreaks and emergency arthropod pest prob-
lems, and they are often required to control plant pathogens, weeds, or nematodes
that cannot be controlled by alternative methods.
RESEARCH APPROACHES TO RESISTANCE MANAGEMENT
Scientists have approached the problem of pesticide resistance in a variety of
ways. 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 developed
to 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 resistance
remains controversial and field-tested experimental data available are not strong
enough to support either model (Roush & Daly 1990, Tabashnik et al. 1992). The hy-
pothesis that reduced fitness, which is often associated with resistance alleles, could
be used in management programs continues to be controversial and may have limited
application (Tabashnik 1990). Not all resistance alleles confer lowered fitness (for ex-
ample, Hoy & Conley 1989, Hoy 1990) and natural selection can select for modifying
genes that restore fitness to individuals carrying resistance alleles. Various monitor-
ing techniques have been developed to identify resistant individuals and detect their
establishment and spread (ffrench-Constant & Roush 1990). These methods are par-
ticularly useful for documenting that resistance has occurred. However, monitoring
methods that would allow us to detect rare resistant individuals in populations in suf-
ficient time that operational programs could be altered remain difficult and expensive
to execute (Brent 1986).
Resistance management research programs and IPM research programs have had
fairly distinct identities to date (Denholm et al. 1992, Croft 1990b, Hoy 1992). Because
they have been distinct, an effective paradigm for resistance management has not
been adopted in US agriculture. The current scenario usually goes something like
this: A pesticide is registered and used, resistant individuals are detected in popula-
tions, people begin to discuss developing and implementing a resistance management
program. With this short-sighted approach, it is exceedingly difficult to develop and
execute 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-
Florida Entomologist 78(3) September, 1995
quency of resistant individuals in populations, detect cross resistances, and evaluate
mode 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 at
least 5% of the population (Brent 1986). Thus, by the time resistant individuals are
detected, selection by additional pesticide applications is likely to increase their fre-
quency in the population. This scenario is particularly familiar with multivoltine and
highly 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 in
arthropods. 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 them
may 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 may
not be cost effective if employed other than as a method to document a problem once
it has developed. Waiting until the pest becomes resistant before instituting a resis-
tance management program is ineffectual (Hoy 1992).
RESISTANCE 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, multitactic
endeavor (Hoy 1992, Fig. 1).
It seems reasonable, conservative, and fiscally-responsible to assume that nearly
all 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 of
resistance development in arthropod pests during the past 40 years. Resistance to
stress 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 most
pesticides, if subjected to appropriate and sustained selection. While new pesticide
classes have been proclaimed to be potential 'silver bullets', and not amenable to re-
sistance development, these hopes have been misplaced to date. It seems appropriate
to 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 (Hoy
1992).
Growers and pest management experts can not afford to rely on pesticides as their
primary 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 increase
the use of nonchemical control tactics such as host plant resistance, biorational meth-
ods, cultural controls, and biological controls (National Research Council 1989, Office
of Technology Assessment 1992). There is an increasing priority by research scien-
Hoy: Symposium on Managing Resistance
Figure 1. Effective resistance management programs will incorporate multiple
tactics as part of a fully integrated IPM program. Pesticides should be used only when
needed, at the lowest rates possible, and in a manner to reduce negative impacts on
arthropod natural enemies. Pesticide labels should include information about its im-
pact on natural enemies.
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 is
critical for improving pest management and environmental quality, and for managing
resistance to pesticides. Enhancing the compatibility of pesticides and biological con-
trol agents is complex and sometimes difficult (Croft 1990a, Hoy 1985a, 1990), but can
reward 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 achieving
improved integrated pest management. For example, some pesticides are relatively
nontoxic 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 exert
unnecessary selection for resistance in the pest. Under these circumstances, it may be
appropriate for the label to contain two different directions for use; one rate could be
recommended 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 the
pesticide to a selected list of biological control agents be determined for each cropping
system. This information should be provided, either on the label or in readily-available
computerized data bases, perhaps via the internet. Without such information, use of
these pesticides could disrupt benefits derived from effective biological control agents.
Florida Entomologist 78(3) September, 1995
This often results in unnecessary use of pesticides, leading to more rapid evolution of
pesticide 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 agents
be 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 on
the impact of pesticides on natural enemies, but additional data are also buried in
many publications or reports that are difficult to find. Unfortunately, even if the data
can 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 impact
of pesticides under field conditions based on laboratory assays (Hoy 1990, Hassan et
al. 1991, Robertson & Preisler 1992). Consequently, the recommendation that labels
or data bases be developed with information on the impact of pesticides on natural en-
emies requires considerable discussion and additional research. Should pesticide
companies 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 not
new, and in Europe standardized bioassays already are being conducted on selected
natural enemy species by a scientific working group (Hassan et al. 1991). Whether
this concept can be imported to the USA and Florida should be explored. Increased in-
ternational consultation and cooperation between scientists, regulatory agencies, and
pesticide 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 the
essential elements of any resistance management program (Croft 1990b, Tabashnik
1990, Metcalf 1994). Thus, the compatibility of pesticides and biological control
agents is a crucial issue in pesticide resistance management.
Attempts to manage pesticide resistance generally has involved making relatively
minor tactical shifts in use patterns. What we need is a major shift in thinking about
pesticide development and use, if we are to develop effective resistance management
tactics. It is time to recognize that effective resistance management begins before the
product 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 application
rates 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 some
cases, new products may not be developed because they are toxic to biological control
agents and thus could disrupt effective IPM programs already in place. This would
only speed up the development of resistance in specific pests.
Some people suggest that financial incentives may have to be provided to induce
pesticide companies to develop and register products that are harmless to biological
control agents (= physiological selectivity). Others argue that many pesticides can be
applied in a manner that affords substantial selectivity (= ecological selectivity) if the
timing, location, rates, and methods of application are altered (Hull & Beers 1985).
Relying on ecological selectivity is more likely to be cost effective than developing
large 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.
Hoy: Symposium on Managing Resistance
THE ROLE OF PESTICIDE-RESISTANT NATURAL ENEMIES
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 a
practical and cost effective tactic for the biological control of spider mites (Hoy 1985a,
b, 1990). Field tests have been conducted with several laboratory-selected phytoseiid
species and some are being used in IPM programs. Laboratory selection of resistant
strains of parasitoids and insect predators currently lags behind efforts with predatory
mites, but several laboratory-selected insect natural enemies are being evaluated for
incorporation 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 is
time consuming and expensive and should not be considered before exploring other,
less expensive, options for IPM and pesticide resistance management.
CONCLUSIONS
We need to maintain a source of pesticides because they are powerful and effective
pest management tools. Pesticides can be highly selective, rapid in their impact,
adaptable to many situations, and relatively economical. Thus, preserving pesticides
is essential.
Effective paradigms for resistance management are not yet deployed in US and
Florida agriculture. This is because resistance management and IPM have been con-
sidered separate issues. We need to recognize that effective resistance management is
based 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 growth
regulators, and mass trapping. A key issue should always be whether pesticides can
be 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.
ACKNOWLEDGMENTS
This is University of Florida, Institute of Food and Agricultural Sciences, journal
series number R-04510. I thank D. C. Herzog for comments on the manuscript and the
Florida Entomological Society for the opportunity to organize the symposium, "Myths
of Managing Resistance". A brief essay containing many of the ideas presented here
has been published (Hoy 1992). The references cited provide an entire to the extensive
literature on resistance and are not complete.
REFERENCES CITED
BRENT, 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.
CORBETT, J. R., K. WRIGHT AND A. C. BAILLIE. 1984. The Biochemical Mode of Action
of Pesticides. 2nd ed., Academic Press, New York.
Florida Entomologist 78(3) September, 1995
CROFT, B. A. 1990a. Arthropod Biological Control Agents and Pesticides.
Wiley-Interscience Publ., New York.
CROFT, B. A. 1990b. Developing a philosophy and program of pesticide resistance
management, pp. 277-296 in R. T. Roush and B. E. Tabashnik [eds.] Pesticide
Resistance in Arthropods, Chapman and Hall, New York.
DENHOLM, I., A. L. DEVONSHIRE AND D. W. HOLLOMON [eds.]. 1992.Resistance 91:
Achievements and Developments in Combating Pesticide Resistance, Elsevier
Applied Science, London.
FFRENCH-CONSTANT, R. H. AND R. T. ROUSH. 1990. Resistance detection and docu-
mentation: the relative roles of pesticidal and biochemical assays. pp. 4-38.
GEORGHIOU, 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.
GEORGHIOU, G. P. AND T. SAITO [eds.] 1983. Pest Resistance to Pesticides, Plenum
Press, New York.
HASSAN, S. A., F. BIGLER, H. BOGENSCHUTZ, E. BOLLER, J. BRUN, J. N. M. CALIS, P.
CHIVERTON, J. COREMANS-PELSENEER, C. DUSO, G. B. LEWIS, F. MANSOUR, L.
MORETH, P. A. OOMEN, W. P. J. OVERMEER, L. POLGAR, W. RIECKMANN, L.
SAMSOE-PETERSEN, A. STAUBLI, G. STERK, K. TAVARES, J. J. TUSET AND G. VIG-
GIANI. 1991. Results of the fifth joint pesticide testing programme carried out
by the IOBC/WPRS-Working Group "Pesticides and Beneficial Organisms".
Entomophaga 36: 55-67.
HOY, 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. IB. Elsevier, Amsterdam.
HOY, M. A. 1985b. Recent advances in genetics and genetic improvement of the Phy-
toseiidae. Annu. Rev. Entomol. 30: 345-370.
HOY, M. A. 1990. Pesticide resistance in arthropod natural enemies: variability and
selection responses, pp. 203-236 in R. T. Roush and B. E. Tabashnik [eds.] Pes-
ticide Resistance in Arthropods, Chapman and Hall, New York.
HOY, M. A. 1992. Guest editorial: Proactive management of pesticide resistance in ag-
ricultural pests. Phytoparasitica 20(2): 93-97.
HOY, 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.
HOY, M. A. AND J. CONLEY. 1989. Propargite resistance in Pacific spider mite (Acari:
Tetranychidae): stability and mode of inheritance. J. Econ. Entomol. 82: 11-16.
HULL, L. A. AND E. H. BEERS. 1985. Ecological selectivity: modifying chemical control
practices 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 Tactics
for Management, National Academy Press, Washington, D.C.
METCALF, R. L. 1994. Insecticides in pest management, pp. 245-284 in R. L. Metcalf
and 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 Tactics
for Management. National Academy Press, Washington, D.C.
NATIONAL RESEARCH COUNCIL. 1989. Alternative Agriculture, National Academy
Press, Washington, D. C.
OFFICE OF TECHNOLOGY ASSESSMENT. 1992. A New Technological Era for American
Agriculture, 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.] Pesticide
Resistance in Arthropods, Chapman and Hall, New York.
SODERLUND, D. M. AND J. R. BLOOMQUIST. 1990. Molecular mechanisms of insecticide
resistance, 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 in
Arthropods, Chapman and Hall, New York.
TABASHNIK, B. E. AND B. A. CROFT. 1985. Evolution of pesticide resistance in apple
pests and their natural enemies. Entomophaga 30: 37-49.
TABASHNIK, B. E., J. A. ROSENHEIM AND M. A. CAPRIO. 1992. What do we really know
about management of insecticide resistance? pp. 124-135 in I. Denholm, A. L.
Devonshire and D. W. Hollomon [eds.]. 1992. Resistance 91: Achievements and
Developments in Combating Pesticide Resistance, Elsevier Applied Science,
London.
THEILING, K. M. AND B. A. CROFT. 1988. Pesticide side-effects on arthropod natural
enemies: a database summary. Agric. Ecosys. Environm. 21: 191-218.
Florida Entomologist 78(3) September, 1995
IMMUNOLOGICAL STUDY OF JUVENILE HORMONE
BINDING PROTEIN FROM HEMOLYMPH OF THE FALL
WEBWORM (LEPIDOPTERA: ARCTIIDAE)
I. H. LEE', HAK R. KIM1 AND RICHARD T. MAYER2
'Department of Biology, Korea University, Seoul, Korea
2USDA, ARS, U.S. Horticultural Research Laboratory,
2120 Camden Road, Orlando, FL 32803-1419
ABSTRACT
The juvenile hormone binding protein (JHBP) was purified from the hemolymph
ofHyphantria cunea (Lepidoptera: Arctiidae) using anion exchange, gel filtration, and
Mono P FPLC chromatofocusing chromatography. The protein is a single polypeptide
(M, 32,000) with an apparent dissociation constant of 0.43 PM for juvenile hormone
III. An antibody developed against hemolymph JHBP (hJHBP) was prepared, and use
of it showed that a protein immunologically identical with hJHBP occurs in fat body
and 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.
RESUME
La protein de union de la hormonajuvenil (JHBP) fue purificada a partir de la he-
molinfa de Hyphantria cunea. Para purificar la JHBP fueron usados el intercambio de
aniones, la filtraci6n en gel y cromatografias cromatofocales Mono P FPLC. La pro-
teina es un polip6ptido simple (M, 32,000) con una constant de disociaci6n aparente
de 0.43 pM para la hormona juvenile III. Fue preparado un antibi6tico desarrollado
contra la hemolinfa JHBP (hJHBP) y su uso demostr6 que una protein inmunol6gi-
camente id6ntica a la hJHBP esta present en el cuerpo graso y el ovario. La hJHBP
de H. cunea no estuvo inmunol6gicamente 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 et
al. 1977). Generally, high M, (over 200,000) and low M, (20,000-40,000) JHBPs are
present in the hemolymph (de Kort & Granger 1981; de Kort et al. 1983). JHBP from
hemolymph 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 that
hemolymph juvenile hormone binding protein (hJHBP) is closely related to
apoprotein-I of lipophorin (Rayne & Koeppe 1988; Koopmanschap & Dekort 1988).
Proteins with JH affinity were found to be present in ovary and fat body and their
characteristics were also reported (van Mellaert et al. 1985; Koeppe et al. 1987; Shem-
Lee et al.: JHBP of H. cunea
shedini & Wilson 1993). However, information on the relationship between hJHBP
and JH binding protein in these tissues is still limited, thus additional information is
needed 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 M1 hJHBP from the
fall webworm, Hyphantria cunea Drury (Lepidoptera: Arctiidae). In addition, the
physicochemical characteristics of hJHBP are described.
MATERIALS AND METHODS
Insects
Fall webworms, Hyphantria cunea, were reared on artificial diet (Dong Bang Oil
Co., Ltd., Seoul, Korea) at 28 1C 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 4C
to remove hemocytes and cell debris. The supernatant was stored at -70C.
Ovaries were dissected from 5-day-old pupae in cold Ringer's solution (128 mM
NaC1, 1.8 mM CaC,, 1.3 mM KC1; pH 7.4). They were used directly for electron mi-
croscopic observation or homogenized and centrifuged at 10,000 g for 10 min and the
supernatant used as the sample.
Fat body was dissected from last instar larvae in Ringer's solution. These tissues
were stored at -70C until used.
Dextran Coated Charcoal (DCC) Binding Assay
A DCC suspension (0.5% active charcoal and 0.05% dextran, M, 80,000) in buffer
(10 mM Tris, 5 mM MgCl,, 50 mM KC1, pH 7.4) was used to separate protein-bound
JH from unbound hormone (Engelmann 1981). [3H]JH III (NEN Corp., Wilmington,
DE; specific activity: 440 GBq/mmol; 50,000 cpm) was put into a microcentrifuge tube
and the solvent evaporated with a gentle stream of nitrogen gas. Buffer (100 pl; 10
mM Tris, 5 mM MgCl,, 50 mM KC1, 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 pl)
was added to the tube, the contents incubated for 2 min and then centrifuged at
10,000 g to remove free hormone. An aliquot (10 pl) of supernatant was transferred to
10 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 100C, Palo Alto, CA).
Purification of JHBP
Anion exchange chromatography. Hemolymph (10 ml) was diluted 1:1 with TPN
buffer [10 mM Tris, 0.1 mM phenylmethylsulfonyl fluoride (PMSF), 0.01% NaN3, pH
8.2] and dialyzed against the same buffer for 4 h. Dialyzed hemolymph was applied to
a DEAE cellulose (DE 52, Whatman, Hillsboro, OR) column (1.8 x 20 cm) and eluted
with 1 column volume of TPN buffer. This was followed by a NaC1 concentration gra-
dient (0-0.5 M) in buffer (10 mM Tris, 0.01% NaN,, 0.1 mM PMSF, pH 8.2, total 300
Florida Entomologist 78(3) September, 1995
ml) at a flow rate of 18 ml per h and then monitored for the presence of JHBP by the
DCC binding assay. Fractions containing JHBP were pooled, dialyzed against 10 mM
Tris buffer (0.01% NaN 0.1 mM PMSF, pH 8.2) and applied to a Sepharose CL-6B
(Pharmacia, Piscataway, NJ) column (0.9 x 15 cm). The column was washed with the
same buffer using a gradient of 0.05-0.35 M NaC1. 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 and
applied to a Sephadex G-100 column (1.8 x 55 cm) and eluted with buffer (25 mM
bis-Tris, pH 7.2) at a flow rate of 12 ml per h. Each fraction (2.6 ml) was monitored for
JHBP by the DCC binding assay. Fractions containing JHBP were pooled, concen-
trated by lyophilization, and applied to a Sephadex G-75 column (0.9 x 20 cm) and
eluted with buffer (25 mM bis-Tris, pH 7.2) at a flow rate of 8 ml per h. Fractions of
1.5 ml were collected.
Mono P FPLC Chromatofocusing ( ......, .. .. "'' Fractions containing JHBP
from the Sephadex G-75 column were subjected to FPLC chromatofocusing chroma-
tography on a Mono P column (Mono P HR 5120, 5 x 200 mm, Pharmacia). Free pro-
teins were eluted from the column with 25 mM bis-Tris buffer (pH 7.2, 10 ml), and
then Polybuffer (pH 7.4, 52 ml) was used to elute resin-bound proteins. Each fraction
was monitored for JHBP by the DCC binding assay.
Determination of Molecular Weight and Isoelectric Point
The molecular weights were determined on sodium dodecyl sulfate-polyacrylamide
gel (10%) as described by Weber & Osborn (1969). Standard molecular weight markers
were: bovine serum albumin, M, 66,200; hen egg albumin, M. 45,000; bovine carbonic
anhydrase, M1 31,000; soybean trypsin inhibitor, M, 21,500; hen egg white lysozyme,
M, 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 were
mixed 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. The
fourth injection was conducted 1 week after the third injection, and a booster injection
was 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 the
booster injection and centrifuged at 10,000 g for 10 min. Antibody against JHBP was
purified 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 plates
were stained in 1% amido black 10B and destined 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. Gels
were stained with silver nitrate according to Wray et al. (1981).
Lee et al.: JHBP of H. cunea
Western blotting was used to detect JHBP on SDS-polyacrylamide gels. Samples
were electrophoresed as above and transferred to nitrocellulose sheets in Tris-glycine
buffer (25 mM Tris, 92 mM glycine, 30% methanol, pH 8.3) at 100 V for 2 h (Towbin
et al. 1979). After transfer, the nitrocellulose sheets were equilibrated in TBS (20 mM
Tris, 500 mM NaC1, pH 7.5) for 10 min and incubated in blocking solution (3% gelatin
in 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-fold
dilution of the primary antibody against JHBP. This sheet was again washed with
TTBS 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 horseradish
peroxidase color development reagent containing 4-chloro-l-naphthol in 20 ml
ice-cold methanol + 0.015% HO, 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 prefixed
in 2.5% glutaraldehyde for 2 h at 4C. Tissues were washed with 0.1 M phosphate
buffer (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 an
Epon-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 was
reacted with the primary antibody solution 30-fold diluted with antibody buffer and
again washed with TBS 3 times to remove nonspecifically attached antibody. This grid
was then reacted with antirabbit IgG combined with protein-A gold particles (30 nm
in 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 80
kV.
RESULTS
Purification of JHBP from Hemolymph
The DCC binding assay for each fraction showed that JHBP was eluted from the
DE-52 column with a linear NaC1 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 NaC1) (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 were
applied to Mono P FPLC chromatofocusing columns and 4 peaks isolated by using A,
(Fig. ID). The fourth peak exhibited high binding activity for [3H]JH III (Fig. 1D, in-
set). Because the JHBP fractions were contaminated with other hemolymph proteins
after Mono P chromatofocusing, gel filtration on Sephadex G-75 was necessary (data
not shown). Fractions were applied to a 10% SDS gel and completely purified JHBP
was confirmed (Fig. 2A).
Characterization of hJHBP
JHBP was electrophoresed with a low molecular weight standard marker to deter-
mine the M, of the JHBP subunit. The M2 was estimated to be 32,000 (Fig. 2B). Also,
456 Florida Entomologist 78(3) September, 1995
A B
3 5
05
0.9-
0,6-
E Ec
1 10 20 30 40 50 60 70 0
FRACTION NO. FRACTION NO.
Figure 1. Chromatography elution profiles for purification ofH. cunea hemolymph
JHBP. A, the first anion exchange chromatography of dialyzed hemolymph proteins
on DE52. Hatched region (35-46) indicates fractions containing JHBP; B, the second
anion exchange chromatography on Sepharose CL-6B. Fractions (47-55, hatched re-
gion) showing radioactivity in DCC binding assay were pooled and lyophilized for the
next purification step; C, gel (Sephadex G-100) filtration of partially purified JHBP.
JHBP containing fractions were confirmed (hatched region, 31-37); D, Mono P FPLC
chromatofocusing 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.
the pi was determined to be 5.3 (data not shown). Saturation analysis of the purified
hJHBP revealed a high affinity for JH III (K, of 0.43 pM) (Fig. 3).
Immunological Studies
Antibodies were made against fractions containing JHBPs (Fig. 4A, lane b) that
had 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 hJHBP
antiserum (Fig. 4B).
A protein that is immunologically identical with hJHBP was found in ovaries by
immunodiffusion and western blotting (Fig. 5). However, the hJHBP ofH. cunea was
neither immunologically related to the lipophorin from H. cunea nor was it similar to
the hJHBP of Bombyx mori and Periplaneta americana (Fig. 5). The western blots
Lee et al.: JHBP of H. cunea 457
A a b c d e B 97.4
a5 .
JHBP
--S
'h.. .. 21.5
--- -14.4
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 from
the posterior portion of the chromatography was confirmed on lane e. The gel (10%)
was stained with silver nitrate. B, SDS-polyacrylamide gel (15%) electrophoresis to
determine the molecular weight of JHBP. The gel was stained with coomassie bril-
liant blue. Right lane contains the low molecular weight standard markers and the
arrow in left lane indicates JHBP
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 immunodiffusion
test (Fig. 5, right) showed an immunopreciptation line extending from the JHBP well
through 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 of
ovary.
DISCUSSION
The DCC binding assay was employed to purify a low M1 JHBP. Fractions showing
radioactivity according to the DCC binding assay were electrophoresed and silver
stained and found to be a single polypeptide with an M, 32,000. The protocols em-
ployed to purify hemolymph JHBP from H. cunea are similar to those used to purify
JHBP of Diatraea grandiosella (Dillwith et al. 1985) and Manduca sexta (Kramer et
al. 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 NM
Florida Entomologist 78(3)
401
20-
September, 1995
50 100 150 200
250
Free 3H JH IIl(hM)
Figure 3. The saturation analysis (Scatchard plot) of the binding of JH III to the
purified hJHBP. ***, total binding; ooo, nonspecific binding; AAA, specific binding.
The specific binding was used for the Scatchard analysis (inset).
JHBP has an M, 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 PM. This
value is similar to those of pyralid moth's 0.08-0.28 pM (Lenz et al. 1986), D. grandi-
osella 0.31 pM, (Dillwith et al. 1985), and M. sexta, 0.44 pM (Kramer et al. 1976). The
JHBP purified from hemolymph of H. cunea is of low M. and high affinity.
Thus far, most JHBP antibody production employed large M, proteins. Prepara-
tion of antibodies for the low M, JHBP was difficult because of the small amount of
0.21-
0.1 5-
i I I
10 20 30 40
Bound(nM)
A
__n-A
rl__ I I I
Lee et al.: JHBP of H. cunea
A
a b
-p. i- *
urwv w.I
qWr ~liq
B a
.1
SJHBP
.~a;h
;' 5'
: . - ,.:
;:
'i
Figure 4. Western blot of purified JHBP. The JHBP band was detected by enzyme
immunoassay 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 three
peaks in Fig. 1D, lane a) that lack JHBP were used for the immunoprecipitation to re-
move other antibodies except antibody against JHBP.
cuv
Y'
,u~-
a
460 Florida Entomologist 78(3) September, 1995
H J OB
2' "
Al
Figure 5. Left panel: Western blots of proteins from the hemolymph (H), JHBP (J),
ovarian extracts (0) ofH. cunea, the hemolymph (B) of B. mori, the hemolymph (P) of
P. americana. The JHBP bands were detected by enzyme immunoassay using anti-
body against the hemolymph JHBP. In three lanes (H,J,0), a single M 32,000 protein
corresponding to the JHBP was detected by the antibody. Right panel: Immunodiffu-
sion test to determine the relationship of several antigens (J, JHBP, 6 gg 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 gg protein; P,
hemolymph of P. americana, 1.2 mg protein; Ab, antibody against JHBP in
hemolymph ofH. cunea).
material in hemolymph. In the present work with H. cunea, immunoprecipitation was
very effective in purifying antibody against low M. JHBP from other antibodies.
hJHBP was confirmed to be lipophorin in Leucophaea maderae (Koeppe et al.
1988) and Locusta migratoria (Koopmanschap & de Kort, 1988). Immunodiffusion
studies conducted here indicate that the hJHBP was neither immunologically related
to lipophorin in H. cunea nor the hJHBP of Bombyx mori and Periplaneta americana.
The hJHBP ofH. cunea appeared to be species-specific.
hJHBP is synthesized by fat body and released into hemolymph. Protein with JH
affinity (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 (van
Mellaert 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. More
information on the relationship between hJHBP and JHBP in the cytosol is required
to better understand the transport of JH from secretary 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.
Lee et al.: JHBP of H. cunea
A *- X ?7 i ty'
C '. -*. '. .' ^^ '*
'- ^-^ '* 4 k-**' ? r'***- *. A S
t7 ^ ,;' <'f. A"< ,
4 .r
** '"ifl^
. .., .
9..
** 'W-.'
:* 'Krt
"A^
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 pm (A), 0.5 pm (B).
REFERENCES CITED
DE KORT, C. A. D., AND N. A. GRAGER. 1981. Regulation of the juvenile hormone titer.
Annu. Rev. Entomol. 26: 1-28.
Florida Entomologist 78(3) September, 1995
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 potato
beetle: a re-examination. Insect Biochem. 13: 481-487.
DILLWITH, J. W., S. D. MANE, AND G. M. CHIPPENDALE. 1985. High affinity juvenile
hormone 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 fat
bodies 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 III
binding 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 the
hemolymph 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 juvenile
hormone-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 and
secretion of juvenile hormone-binding protein by fat body from the cockroach
Leucophaea maderae: protein a immunoassay. Insect Biochem. 17: 1027-1032.
KOOPMANSCHAP, A. B., AND C. A. D. DE KORT. 1988. Isolation and characterization of
a high molecular weight JH III transport protein in the hemolymph ofLocusta
migratoria. 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 juvenile
hormone 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 hemolymph
of 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, 0. 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 dependent
changes in the binding and hydrolysis of juvenile hormone in the hemolymph
of last instar larvae of Galleria mellonella. Insect Biochem. 13: 435-441.
PRESTWICH, G. D., AND J. K. ATKINSON. 1990. Rapid purification and N-terminal
amino 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 hormone
binding 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. Insect
Biochem. 9: 569-575.
Lee et al.: JHBP of H. cunea 463
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 proteins
from 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 binding
proteins in Sarcophaga bullata hemolymph and vitellogenic ovaries. Insect
Biochem. 15: 655-661.
WEBER, K. AND M. OSBORN. 1969. The reliability of molecular weight determination
by dodecyl sulfate-polyacrylamide gel electrophoresis. J. Biol. Chem. 244:4406-
4412.
WHITMORE, E., AND L. I. GILBERT. 1972. Haemolymph lipoprotein transport ofjuve-
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
DEGRADATION OF RED IMPORTED FIRE ANT
(HYMENOPTERA: FORMICIDAE) YOLK SPHERES
AMADOU S. BA AND SHERMAN A. PHILLIPS, JR.
Department of Plant and Soil Science
Texas Tech University
Lubbock, TX 79409-2134
ABSTRACT
Transmission electron microscopy reveals that the eggs of the red imported fire
ant, Solenopsis invicta Buren, contain yolk spheres that are evenly distributed
throughout the center of the egg. Yolk degradation occurs sequentially as evidenced
by the simultaneous observation of three stages of yolk platelet degradation within
the eggs.
Key Words: Insecta, Solenopsis invicta, egg, oogenesis.
RESUME
La microscopia electr6nica de transmisi6n revela que los huevos de la hormiga de
fuego Solenopsis invicta Buren, contienen esferas de yema que se distribuyen pareja-
mente a lo largo del centro del huevo. La observaci6n simultanea de tres etapas de de-
gradaci6n en las esferas de yema dentro de los huevos evidencia que la degradaci6n
de la yema ocurre secuencialmente.
The most abundant subcellular organelles of mature oocytes in arthropods are
yolk granules, constituting approximately 80% of the total protein present (Medina et
al. 1988). Arthropod yolk spheres (granules or platelets) are phosphoglycolipoproteins
with varying amounts of phosphorus, lipids, and carbohydrates (Yamashito & Indra-
Florida Entomologist 78(3) September, 1995
sith 1988). Much of the yolk material consists of vitellin, a protein that is derived from
vitellogenin (Kunkel & Nordin 1985; Tata & Smith 1979; Wallace 1985). Vitellin,
which comprises 60-90% of the total mass of yolk proteins (Kunkel & Nordin 1985), is
transported 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 been
determined (Bownes & Hames 1977; Yamashita & Indrasith 1988). For example, yolk
spheres of the fruit fly (Drosophila) constitute about 80% of the total protein of the
mature oocyte (Vallejo et al. 1981). However, the degradation of yolk in arthropods is
not a process that is well understood.
The red imported fire ant, Solenopsis invicta Buren, was introduced into the
United States from South America around 1940 and has rapidly become a major pest
in 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 understand
the mode of action of the insect growth regulator fenoxycarb on the red imported fire
ant (Glancey & Banks 1988). Fenoxycarb caused a lack of cell differentiation involv-
ing oocytes, trophocytes, and follicular epithelial cells. In addition, because nurse cells
did not develop, oocytes did not contain the necessary yolk spheres. Therefore, eggs
did 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 lack
of 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 yolk
sphere formation in oocytes of the red imported fire ant. Our paper, herein, is the first
to describe the appearance of naturally occurring yolk degradation in fertilized eggs
of ants, and in particular, of the red imported fire ant.
MATERIALS AND METHODS
Red imported fire ant colonies were collected from Abilene and Victoria, Texas, in
the 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 wetting
agent 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 phosphate
buffer (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 dehydrating
material (CuSO4). Eggs were embedded in Spurr's low viscosity medium (Spurr 1969).
Sorvall MT-2B and glass knives were used for ultramicrotomy. Sections were stained
with methanolic uranyl acetate and lead citrate, and micrographs were taken using
a Hitachi HS-9 transmission electron microscope.
RESULTS AND DISCUSSION
Electron micrographs showed the eggs to be filled with large and dense storage or-
ganelles (yolk spheres). Yolk platelets were distributed evenly throughout the center
of 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 from
tissue of the developing embryo. We observed three categories of yolk granules: 1)
BA & Phillips: Fire Ant Yolk Sphere Degradation
Figure 1. Transmission electron micrographs of cross sections of red imported fire
ant eggs showing sequential degradation of yolk spheres: us = undegraded yolk
spheres; ps = partially degraded yolk spheres; ds = degraded yolk spheres; ex = exo-
chorion; en = endochorion; vtm = vitellin membrane (Fig. 1A, bar = 36 pm; Fig. 1B, bar
= 50 pm; Fig 1C, bar = 52 pm; Fig. 1D, bar = 67 pm).
large, membrane bound, undegraded dense yolk spheres; and 2) partially degraded
and 3) degraded yolk spheres (Fig. 1C). Not all yolk platelets in red imported fire ant
eggs 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 ant
eggs support earlier studies showing sequential yolk degradation in other arthropods.
For example, proteinase activity is involved in yolk granule organelle degradation
during embryogenesis in Drosophila, and this activity is developmentally regulated
(Medina et al. 1988). Fagotto (1991) states that yolk spheres of the eggs of an African
soft 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 yolk
during reabsorption of developing oocytes in red imported fire ant queens after treat-
Florida Entomologist 78(3) September, 1995
ment with an insect growth regulator, proteinase is inactive during those stages of egg
formation. Therefore, normal yolk degradation would not be observable. Proteinase is
only 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 these
degradation mechanisms are applicable to other systems (Yamashita & Indrasith
1988). However, the general mechanism for yolk degradation in many insects has
been shown to be sequentially controlled (Yamamoto & Takahashi 1993), and proteins
within these yolk spheres are differentially hydrolyzed.
ACKNOWLEDGMENT
This study was supported by the Texas State Line Item for Fire Ant Research. We
thank H. Thorvilson, E. White, and C. Landry for reviewing our manuscript.
REFERENCES CITED
BOWNES, M., AND B. D. HAMES. 1977. Accumulation and degradation of three major
yolk proteins in Drosophila melanogaster. J. Exp. Zool. 200: 149-156.
FAGOTTO, F. 1991. Yolk degradation in tick eggs: III. Developmentally regulated acid-
ification of the yolk spheres. Develop. Growth and Differ. 33: 57-66.
GLANCEY, B. M., AND W. A. BANKS. 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.
KUNKEL, J. G., AND J. H. NORDIN. 1985. Yolk proteins, pp. 83-111 in G. A. Kerkut and
L. I. Gilbert [eds.] Comprehensive Insect Physiology, Biochemistry and Phar-
macology, Vol. 1. Pergamon Press, Oxford.
MCGREGOR, D. A., AND B. G. LOUGHTON. 1974. Yolk protein degradation during em-
bryogenesis of the African migratory locust. Canadian J. Zool. 52: 907-917.
MEDINA, M., P. LEON, AND C. G. VALLEJO. 1988. Drosophila cathepsin B-like protein-
ase: A suggested role in yolk degradation. Arch. Biochem Biophysics 263: 355-
363.
POSTLETHWAIT, J. H., AND F. GIORGI. 1985. Vitellogenesis in insects, pp. 85-126 in W.
L. Browder [ed.]. Developmental Biology: A Comprehensive Synthesis, Vol. 1:
Oogenesis. Plenum Press, New York.
SPURR, A. R. 1969. A low-viscosity epoxy resin medium for electron microscopy. J. Ul-
trastructure Res. 26: 31-43.
TATA, J. R., AND D. F. SMITH. 1979. Vitellogenesis: A versatile model for hormonal reg-
ulation of gene expression. Rec. Prog. Horm. Res. 35: 49-95.
VALLEJO, C. G., R. PERONA, R. GARESSE, AND R. MAROCO. 1981. The stability of the
yolk granules ofArtenia. An improved method for their isolation and study. Cell
Diff. 10: 345-356.
WALLACE, 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.
YAMASHITA, 0., AND L. S. INDRASITH. 1988. Metabolic fates of yolk proteins during
embryogenesis in arthropods. Develop. Growth and Differ. 30: 337-346.
YAMAMOTO, Y., AND S. Y. TAKAHASHI. 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
ORGAN CULTURE OF SALIVARY GLANDS OF MALE
ANASTREPHA SUSPENSE (DIPTERA:TEPHRITIDAE)
REJANE R. DE MORAES', JAMES L. NATION AND JAMES E. MARUNIAK
Department of Entomology and Nematology Hull Road, Building 970
University of Florida
Gainesville, FL 32611
'Fellow of National Research Council (CNPq). Brasilia, Brazil
ABSTRACT
Salivary glands from male Caribbean fruit flies, Anastrepha suspense, 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 successfully
cultured. Two different procedures were used to evaluate viability. A trypan blue dye
test revealed that no more than 10% of the cultured cells were non-viable, since they
acquired blue coloration. In the second set of experiments, acid phosphatase activity
of the cultured glands was measured colorimetrically from absorbance at 415 nm of
liberated p-nitrophenol. This method indicated that the cultured glands stayed met-
abolically active, with enzyme activity equal to that of freshly excised glands on the
fifth, eighth, and fifteenth days of the experiment. Scanning electron microscopy
(SEM) showed no visual differences in tissue organization and size of individual cells
when freshly excised salivary glands and cultured glands were compared. SEM
showed that the cells of cultured glands were neither swollen nor shrunken, and
close-up views showed no evidence of cell deterioration or lysing.
Key Words: Salivary glands, organ culture, Caribbean fruit fly, Tephritidae, Anas-
trepha suspense, acid phosphatase.
RESUME
Las glandulas salivares de los machos de la mosca caribefia de las frutas, Anas-
trepha suspense, fueron aisladas individualmente por disecci6n y cultivadas en medio
de Schneider que conteniene 10% de suero fetal bovino y antibi6ticos. Un total de 103
glandulas salivares fueron cultivadas con exito. Dos process diferentes fueron usa-
dos para evaluar la viabilidad. Una prueba de tinci6n de trypan azul revel6 que menos
del 10% de las celulas cultivadas no era viable porque adquirian una coloraci6n azul.
En una segunda series de experiments, la actividad de fosfatasa acida de las glandu-
las cultivadas fue estudiada colorimetricamente mediante la absorbancia de p-nitro-
fenol liberado a 415 nm. Este m6todo indic6 que las glandulas cultivadas permanecian
metabl6icamente activas, con actividad enzimatica en los dias quinto, octavo y d6ci-
moquinto del experiment igual a la de las glandulas reci6n extirpadas. La Microsco-
pia Electr6nica de Barrido (MEB) no mostr6 diferencias visuales en la organizaci6n de
los tejidos y en el tamaio de las c6llulas individuals, cuando fueron comparadas las
glandulas frescas y las cultivadas. La MEB mostr6 que las celulas de las glandulas
cultivadas no estaban hinchadas ni contraidas, y las vistas tomadas de cerca no evi-
denciaron deterioraci6n o rompimiento cellular.
Organ culture is a technique in which whole organs or representative parts are
maintained as tissues and retain their intrinsic distribution, numerical and spatial
Florida Entomologist 78(3) September, 1995
orientation of explanted cells (Freshney 1987). Most organ cultures do not grow or, if
they 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 isolated
cells.
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 primarily
with insect endocrinological studies (Marks 1976). In vitro cultures of imaginal discs
of 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 of
20-hydroxyecdysone and ecdysone in development and metabolic activity. Organ cul-
tures of chironomid salivary glands have been extensively utilized in investigations of
polytenic cell differentiation, cytogenetics, biochemistry and physiology (Firling &
Kobilka 1978; Firling & Hou 1980).
The salivary glands in Anastrepha suspense (Loew) are sexually dimorphic, with
males having much enlarged glands that terminate on each side of the body in a large
ball 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 same
time 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 not
they are involved in pheromone production, if a culture technique for these glands
was 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 pheromone
from 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 artificial
medium for at least seven days; and to establish protocols for monitoring the viability
of cultured glands.
MATERIALS AND METHODS
Male A. suspense were obtained from the mass rearing facility at the Florida State
Department of Agriculture and Consumer Services, Gainesville, FL. Organ cultures
of 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 dissect
the glands were sterilized. Microscopes, lamps and wax dishes were sterilized with
70% 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 after
each fly was dissected. Glands were dissected under sterile conditions in a sterile
hood. Male flies were surface-sterilized by immersion in 70% ethanol for 10 min and
rinsed in sterile saline solution for 10 min. Just prior to each dissection, the head and
legs of each fly were removed to diminish movement and to facilitate salivary gland
removal. Each fly was pinned ventral side up in a wax dish containing sterile saline
solution. A longitudinal cut was made through the cuticle and the lateral sides of the
abdomen 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 series
of dissections were complete, then they were transferred to the culture medium.
Moraes et al.: Salivary Gland Cultures
Organ Culture
Schneider's medium (Schneider, 1964) containing 10% Fetal Bovine Serum and
the antibiotics penicillin (100 units per ml), streptomycin (100 Pg per ml), amphoteri-
cin B (0.2 pg per ml), gentamycin (50 pg per ml), geocillin (100 pg per ml), kanamycin
(100 pg per ml) and tetracycline (10 pg per ml) were used to culture the salivary
glands. [Schneider's medium has been used successfully to culture Drosophila cells
(Schneider & Blumenthal, 1978).] A24-cluster-well tissue culture plate (Corning) was
used to culture the glands with 0.5 ml medium and two salivary glands per well. The
salivary gland cultures were kept in an incubator at 27'C. Four experiments were
conducted 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 oftrypan blue dye was used as an indicator of
viability of the cultured glands, based on the principle that dead cells allow the dye to
penetrate while live cells exclude the dye. Each day during one week of culture, 250
pl of trypan blue was added to the culture medium in one of the wells. Penetration or
exclusion of the dye in the 2 glands was determined by microscopic examination. In
experiments 2, 3 and 4, viability was monitored by determining the activity of acid
phosphatase on the fifth, eighth and (only in experiment 4) on the fifteenth day. On
each of these days, about 10 glands were removed from culture, rinsed with saline so-
lution, and homogenized in 250 pl of 0.1 M citrate buffer, pH 5.5. Seventy p of the
glandular homogenate was added to each of two tubes containing 1 ml of 7 mM
p-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 were
the 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 cultured
glands were prepared to receive 70 pl of the fresh gland homogenate positive controls.
All tubes were incubated at 30C for 10 min. The reaction was stopped by adding 2 ml
of 10% NaCO, to each tube. This altered the pH, stopped acid phosphatase activity
and caused a shift in the electron structure of the released p-nitrophenol so that it
gave a yellow color and absorbed light strongly at 415 nm. The amount of
p-nitrophenol released was determined at 415 nm in a Spectronic 20D colorimeter. A
standard 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. The
substrate was dissolved in 0.1 M citrate buffer (pH 5.5) just before use. Incubation
tubes 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 by
adding 3 ml of 10% NaCO, 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's
fixative, washed in 70% ethanol to remove excess picric acid, and dehydrated through
an alcohol series to 100% ethanol. Glands were transferred from 100% ethanol to hex-
amethyldisilasane (HMDS) (Nation 1983), transferred once to fresh HMDS and air
dried. The dried tissues were mounted on stubs, gold coated with a sputter-coater, and
observed in a Hitachi S 570 instrument with filament voltage at 15 or 20 KV.
Florida Entomologist 78(3) September, 1995
RESULTS AND DISCUSSION
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 15
days. Scanning electron microscopy (SEM) showed no visual differences in tissue or-
ganization and size of individual cells between freshly excised salivary glands and
glands cultured for 8 days (Fig. 1). Cells in the cultured glands did not shrink, collapse
or 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 the
gland 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 the
glands were alive and apparently functioning normally. Some blue cells were ob-
served on the periphery of the glands, but their percentage was never greater than
10%. In a preliminary experiment to measure acid phosphatase activity, we found
that 57 pg of p-nitrophenol per gland per h was released by glands cultured for 5 days
post-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 by
cultured 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 that
acid phosphatase activity of freshly excised glands and cultured glands did not differ
in glands that were 5, 8 or 15 days old. The substrate, p-nitrophenol phosphate, added
to 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 preliminary
data above.
The surface sterilization of adult flies, and the antibiotics in the culture medium
were effective in maintaining sterility, since only one preparation showed evidence of
contamination 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% ethanol
for several min prior to isolating their wing discs for organ cultures, and Oberlander
& Leach (1979) surfaced sterilized larvae ofP. interpunctella (Hubner) by submerging
them 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 activity
and protein synthesis have often been used to measure the viability of cultured organs
and cells. Firling & Kobilka (1978) measured the ability of cultured Chironomus ten-
tans (Fabricius) salivary glands to incorporate "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 of
0.12% trypan blue, while injured cells did not exclude the dye (Firling & Kobilka
1978). The few blue cells observed on the surface of the male salivary glands in the
present experiments may well have been injured in the dissection or handling proce-
dures, thus causing them to leak and allow uptake oftrypan blue dye. Monitoring for
trypan blue dye uptake is easier than measuring acid phosphatase activity and might
be the method of choice in any future experiments with salivary glands from fruit
flies.
Moraes et al.: Salivary Gland Cultures
Figure 1. Scanning electron microscopy comparing cultured (8-days-old) and
freshly excised male salivary glands (from 8-day-old males) from Caribbean fruit flies
(Anastrepha suspense). A and B, 150 x magnification; C and D, 500 x magnification.
ACKNOWLEDGMENTS
We thank Kathy Milne for technical assistance in the laboratory, Drs. Herbert
Oberlander 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.
Florida Entomologist 78(3) September, 1995
TABLE 1. COMPARISON OF PHOSPHATASE ACTIVITY BETWEEN CULTURED AND FRESHLY
EXCISED SALIVARY GLANDS.
P-Nitrophenol
Salivary Glands pg per Gland per Hour
Mean' SD
5 Days 8 Days 15 Days
Cultured 7.36 0.21 7.26 0.23 6.69 0.02
Fresh 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.
REFERENCES CITED
FIRLING, C. E., AND B. KOBILKA. 1979. A medium for the maintenance of Chironomus
tentans salivary glands in vitro. J. Insect Physiol. 25: 93-103.
FIRLING, C. E., AND A. HOU. 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.
FRESHNEY, R. I. 1987. Culture of animal cells: A manual of basic technique. 2nd ed.
Alan R. Liss, Inc, New York, NY. 397pp.
MARKS, E. P. 1976. The uses of cell and organ cultures in insect endocrinology, in E.
Kurstak, and K. Maramorosch, [eds.], Invertebrate tissue culture, applications
in medicine, biology, and agriculture. Academic Press, New York. 398pp.
NATION, J. L. 1972. Courtship behavior and evidence for a sex attractant in the male
Caribbean fruit fly, Anastrepha suspense. Ann. Entomol. Soc. America. 65:
1364-1367.
NATION, J. L. 1974. The structure and development of two sex specific glands in male
Caribbean fruit flies. Ann. Entomol. Soc. America. 67: 731-734.
NATION, J. L. 1981. Sex-specific glands in tephritid fruit flies of the genera Anas-
trepha, Ceratitis, Dacus, and Rhagoletis (Diptera: Tephritidae). Int. J. Insect
Morphol. & Embryol. 10: 121-129.
NATION, J. L. 1983. A new method using hexamethyldisilazane for preparation of soft
insect tissues for scanning electron microscopy. Stain Technology. 58:347-351.
NATION, J. L. 1989. The role of pheromones in the mating system ofAnastrepha fruit
flies, pp. 189-205 in A. S. Robinson and G. Hooper, [eds], Fruit Flies Their Bi-
ology, Natural Enemies and Control, Vol. 3A, Elsevier Press.
OBERLANDER, H. 1976. Hormonal control of growth and differentiation of insect tis-
sues cultured in vitro. In Vitro. 12: 225-235.
OBERLANDER, H., AND C. E. LEACH. 1979. The action of P-ecdysone and juvenile hor-
mone in organ cultures of lepidopteran imaginal disks. TCA Manual. 5: 993-
995.
OHMORI, K., AND T. OHTAKI. 1973. Effects of ecdysone analogues on development and
metabolic 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 the
larva of the blow fly, Calliphora erythrocephala. J. Insect Physiol. 20: 329-347.
RAIKOW, R., AND J. W. FRISTROM. 1971. Effects of P-ecdysone on RNA metabolism of
imaginal disks ofDrosophila 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.
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 of
Drosophila, Vol 2A, Academic Press, New York.
SRINIVASAN, A., J. A. COFELT, AND H. OBERLANDER. 1979. In vitro maintenance of the
sex 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)
TODD E. SHELLY' AND ETHEL M. VILLALOBOS2
'Hawaiian Evolutionary Biology Program,
University of Hawaii, Honolulu, HI 96822
'Biology Department,
Chaminade University, Honolulu, HI 96816
ABSTRACT
Laboratory tests were conducted to assess the effect of the parapheromone cue
lure on the mating behavior of male Bactrocera cucurbitae (Coquillett). Exposure to
cue lure resulted in a short-term mating advantage. For wild flies, treated males that
fed on cue lure on the day of testing, or 1 day prior to testing, mated more frequently
than control males that had no prior exposure to cue lure. However, control and
treated males had similar mating success in tests performed 3 or 7 days after the
treated 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, though
this advantage was evident for only 1 day following exposure. Cue lure appeared to
enhance 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 exposed
to cue lure 1 week prior to release were less likely to be captured (in Steiner traps
baited with cue lure and naled) than unexposed males. These findings suggest that
exposure of sterile males to cue lure might improve the effectiveness of sterile insect
release as well as enable simultaneous control programs of sterile insect release and
male annihilation.
Key Words: Bactrocera cucurbitae, parapheromone, sterile insect release, Hawaii.
RESUME
Se estudio en el laboratorio el efecto de la paraferomona "cue lure" en el compor-
tamiento sexual de los machos de Bactrocera cucurbitae (Coquillett). La exposici6n a
la "cue lure" tuvo como resultado ventajas en los apareamientos a corto plazo. Los ma-
chos salvajes que fueron alimentados con "cue lure" el mismo dia, o el dia anterior al
experiment se aparearon mas frecuentemente que los machos del grupo control que
no habian sido expuestos a la paraferomona. Sin embargo, los machos de ambos gru-
pos tuvieron 6xitos de copulaci6n similares en experiments conducidos 3 o 7 dias des-
pu6s de haber sido expuestos a la "cue lure". La exposici6n a la "cue lure" de los
Florida Entomologist 78(3) September, 1995
machos criados en el laboratorio e irradiados tambien aument6 el exito de copulaci6n
en comparaci6n con los machos salvajes que no habian recibido la paraferomona, pero
esta ventaja sexual fue efectiva s61o un dia despues del tratamiento. Al paracer la "cue
lure" aumenta la efectividad del comportamiento sexual porque estimula la actividad
vibratoria de las alas, pero no afecta la calidad atractiva de la serial quimica en si. Un
studio de campo indic6 que los machos irradiados expuestos a la "cue lure" la semana
anerior 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 est6riles a la "cue lure" podria aumentar la
efectividad de las liberaciones de insects est6riles, asicomo permitir el uso simulta-
neo de programs de control basados en la liberaci6n de insects est6riles y aniquila-
ci6n de machos.
The males of many tephritid species are strongly attracted to specific chemical
compounds, termed male lures or parapheromones, which either occur naturally in
plants or are (presumed) synthetic analogues of plant-borne substances (Chambers
1977; Sivinski & Calkins 1986; Fletcher 1987). Several well-known examples include
the attraction of male Mediterranean fruit flies, Ceratitis capitata (Wiedemann), to
trimedlure, male oriental fruit flies, Bactrocera dorsalis (Hendel), to methyl eugenol,
and male melon flies B. curcubitae (Coquillett) to cue lure. Because they are powerful
attractants, parapheromones are used in current control programs of tephritid pests
for detection and monitoring of populations and for eradication via male annihilation
(Chambers 1977; Economopoulos & Haniotakis 1994).
Recent studies (Shelly & Dewire 1994; Shelly 1994) onB. dorsalis suggest another
potential use of male lures in control efforts. Data collected in the laboratory showed
that (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 eugenol
was 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 2
factors: treated males signaled (wing-fanned) more frequently and for a given level of
signaling attracted more females per min than control males. In addition, field tests
showed that treated males were less likely to be captured (in traps baited with methyl
eugenol and naled) than control males for as long as 35 days after exposure. These
studies suggest that the effectiveness of control efforts might be enhanced by exposing
sterile 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 parapheromones
and sexual behavior by investigating the influence of cue lure on the mating behavior
of 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 males
and, if so, for how long after exposure. As will be described, the mating trials did, in
fact, show that cue lure enhanced mating performance, and data were then gathered
to 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 the
probability of capturing (in traps baited with cue lure and naled) irradiated males fol-
lowing their release in the environment.
Shelly & Villalobos: Melon Fly Behavior 475
MATERIALS AND METHODS
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 earlier
reports. Mating occurs at dusk in male aggregations termed leks. Males perch singly
on leaf under surfaces of both host and non-host plants and defend their site against
intruding 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 et
al. 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.) After
several 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 the
rectal 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 plants
and 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 and
to 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 200-300
adults reared from Coccinia grandis (L.) collected in Waimanalo, Oahu, Hawaii. The
present study was conducted during July-September, 1994, and consequently the flies
were 3-5 generations removed from the wild. (Though this stock may have undergone
genetic 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 m
by 0.6 m) with superabundant food (a mixture of honey and protein hydrolysate) and
water. 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 days
of 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 individuals
per 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 stock
will hereafter be referred to as "sterile males"). Pupae that had been exposed to 10
krad of gamma radiation from a "Co source were obtained 1 day before eclosion. Sexes
Florida Entomologist 78(3) September, 1995
were separated within 3 days of eclosion [sexual maturity in this stock is attained at
about 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 placed
with 3 wild females in screen cages at least 4 h before sunset. Six to 9 cages were used
on a given day. The cages were 30-cm cubes with one side open and fitted with a cloth
sleeve. Before placement in the cages, males from the 2 groups were cooled for several
minutes 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 wild
males denied access to cue lure (control males). Treated males (21-25 days old) were
given unrestricted access during a 2-h period to a cotton wick (5 cm long) to which 1.5
ml of pure cue lure had been applied (cue lure obtained from the USDA/ARS Fruit and
Vegetable 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 direct
contact 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 males
was 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 midday
period 1 day or 3 days prior to testing. In the latter two experiments, males of both
groups were used during their first 2 weeks of sexual maturity (21-30 days and 12-21
days of age for wild and sterile males, respectively). In the final experiment, wild and
sterile 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 production
and attractiveness, we placed treated and control males in "minicages" within a
larger flight cage and simultaneously monitored signalling by, and female attraction
to, males (following the basic protocol of Poramarcom & Boake [1991]). Because wild
males 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 in
the 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 day
prior 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 were
covered 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 selected
branches 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 days
of 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 lights
were then extinguished, and the flight cage, which was near a west-facing window, re-
ceived only natural light.
Shelly & Villalobos: Melon Fly Behavior 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 the
minicages. Attractiveness of male signaling was thus based on the number of female
sightings 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 of
an observation period, individuals of both sexes were discarded, and a new set of
males 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 Hawaii
at 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 in
a circle (50-m radius) around a central release point. The same plants were used in all
replicates. Traps were suspended in the canopy (2 m above ground) by a 30-cm wire
fastened to a branch. Each trap contained a 5-cm long cotton wick to which 2.0 ml of
cue lure (5% naled) had been applied.
Treated sterile flies (12-15 days old) were given access to a wick containing 1.5 ml
of cue lure for a 2-h period 1 day prior to release. Similarly-aged control males (18-23
day old) that had no exposure to cue lure were also released. Prior to release, control
and treated males were marked on the thorax with different color combinations of
enamel paint (a given combination was used only once). Flies were released between
1000-1100 hours by removing the screen top and gently tapping the bucket to induce
flight. Traps were checked 48 h after release, and captured flies were examined indi-
vidually for markings in the laboratory. Seven replicates were conducted with 100
males 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.
RESULTS
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 males
tested 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 between
control 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 that
were permitted only to approach a cue lure source without contacting it did not have
a mating advantage over control males. Treated males obtained only 51% (38/74) of all
matings (t=0.2; P > 0.05; binomial test).
Cue lure also influenced the relative mating competitiveness of wild and sterile
males. In the second experiment (where cue lure was not given to either wild or sterile
males), 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 flies
exposed 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-
Florida Entomologist 78(3) September, 1995
SControl f Treated
50
40
C
30
20
10
w/1
'/ /
'/ /1
/*
/I
I//
/
//
I1
Days since exposure
Figure 1. Numbers of matings obtained by control vs. treated wild males in trials
conducted 0, 1, 3, or 7 days after treated males were exposed to cue lure.
ing also displayed increased mating success and obtained nearly 1/4 (22/95=23%) of
the 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 with
Yates correction). Despite their improved performance, wild males still accounted for
a 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 the
relative 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) of
all 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 cue
lure was not given to either group of males (P > 0.05 in both comparisons; G test with
Yates correction).
Signal Production and Attractiveness
Males exposed to cue lure called more frequently than control males (Fig. 2). An
average of 70 instances of wing-fanning (maximum value possible=135=3 males per
minicage over 45 checks) was recorded for minicages with treated males compared to
only 54 instances for minicages with containing control males (U=149; P < 0.001;
Mann-Whitney test). In addition, more females were sighted on minicages with
treated 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 with
control males (U=161; P < 0.001; Mann-Whitney test). This difference in female arriv-
Shelly & Villalobos: Melon Fly Behavior
* Control x Treated
0
*0
40 50 60
70 80 90 100
Male wing-fanning
Figure 2. Relationship between female sightings and wing-fanning for control (0)
and treated (*) males. Each point represents a minicage that contained three males.
Abscissa represents the total number of wing-fanning instances recorded for all 3
males/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: control
males -Y=1.05X-25.2; r=0.75; treated males -Y=0.81X-0.6; pooled (shown) -Y=1.07X-
22.6; rO=0.64.
als appeared to reflect the differential calling activity between control and treated
males 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 for
control 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 than
sterile 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).
DISCUSSION
The present study shows that, under laboratory conditions, exposure to cue lure
confers a mating advantage to male B. cucurbitae. However, whereas a positive effect
of methyl eugenol was evident one month after exposure (Shelly & Dewire 1994), cue
lure provided a mating advantage for only one day following feeding. In the melon fly,
100-
80-
0)
--
| 60-
r-
40-
E
i)
LL
0O
30
|
RSS
