Introduction

The human desire to control insects has existed about as long as humans themselves have. Insects annoy people by biting, stinging, and generally getting in the way. However, this desire for insect control increased significantly with two events: the realization that insects can spread human diseases such as malaria and yellow fever, and the rise of agriculture. As the world’s population increases, the need to keep insects from destroying food crops becomes even more urgent; currently, insects destroy 13% of American crops intended for human consumption. That figure is even higher worldwide (Miller, 1998).

Human attempts at insect control have changed over time from natural methods to synthetic chemical control, and now, again, we look to natural methods. While today’s synthetic insecticides are presumably safer and less persistent than those used in the past, such as DDT, they are still a cause for concern. Long-term exposure to modern synthetic insecticides has been associated with cancer, liver damage, immunotoxicity, birth defects, and reproductive problems in humans and other animals (Kegley and Wise, 1998). This is why development of biologically natural methods of insect control, or biopesticides, is preferential today.

A Brief History of Bacillus thuringiensis

What is Bt?

Perhaps the most well known and widely used biopesticide comes from Bacillus thuringiensis (Bt), a bacterium that produces insecticidal proteins during its sporulation. This common soil bacterium, most abundantly found in grain dust from silos and other grain storage facilities, was discovered first in Japan in 1901 by Ishawata and then in 1911 in Germany by Berliner (Baum et al., 1999). It was subsequently found that thousands of strains of B. thuringiensis exist (Lereclus, 1993). Each strain produces its own unique insecticidal crystal protein, or delta-endotoxin, which is encoded by a single gene on a plasmid in the bacterium (Whalon and McGaughey, 1998). The insecticidal activity of the toxins from each Bt strain differs. Nevertheless, the set of Bt delta-endotoxins affects a variety of species from the orders Coleoptera (beetles), Lepidoptera (moths and butterflies), and Diptera (flies and mosquitoes) (Gould and Keeton, 1996). Some Bt delta-endotoxins have a toxicity on par with that of widely used organophosphate pesticides. Unlike organophosphates, which are quite general in their effect, Bt’s toxins are very specific to certain harmful insects and are therefore safe to most beneficial insects and other animals. Additionally, Bt toxins are biodegradable and do not persist in the environment (van Frankenhuyzen, 1993).

A Bt Timeline

Bacillus thuringiensis first became available as a commercial insecticide in France in 1938 and in the 1950s entered commercial use in the United States. For many years, Bt primarily came in the form of a spray to be applied to crops. The non-persistent nature of the insecticide necessitated many reapplications during early use (van Frankenhuyzen, 1993).

In the 1980s, commercial interest in Bt grew very rapidly as many popular synthetic insecticides became ineffective due to insect resistance, or became unusable due to environmental restrictions, and as the field of genetic engineering grew. In 1987 came the first reports of insertion of genes encoding for Bt delta-endotoxins into plants. The first transgenic plants to express Bt toxins were tobacco and tomato plants (van Frankenhuyzen, 1993). The first Bt plant-pesticide, Bt field corn, was registered with the United States Environmental Protection Agency in 1995 (USEPA, 1999b). Today, major Bt transgenic crops also include corn, cotton, potatoes, and rice. The engineering of plants to express Bt delta-endotoxins has been especially helpful against pests that attack parts of the plant that are usually not well-protected by conventional insecticide application. A prime example of this is protection against Ostrinia nubilalis, the European corn borer. Larvae of this lepidopteran bore into the stalk of a corn plant and destroy its structural integrity. In the stalk, the pest is relatively safe from pesticide application. With toxins engineered into the plant, O. nubilalis is exposed and its damage becomes easier to control (Ely, 1993). Overall, because of benefits such as these, Bt has become a major presence in agriculture. In 1997, Bt cotton, corn, and potatoes covered nearly 10 million acres of land in the United States alone. These crops have also been commercialized and are in wide use in Canada, Japan, Mexico, Argentina, and Australia (Frutos et al., 1999). While using Bt in the form of transgenic crops is now very common, the more traditional spray form of Bt is still widely used (Liu and Tabashnik, 1997).

Mode of Action

Bt directly causes mortality in insects, and isolates of the toxin from different strains follow similar modes of action. After the delta-endotoxin crystals are ingested, they are dissolved in the insect midgut, liberating the protoxins of which they are made. These are proteolytically processed into fragments, one of which binds to cells of the midgut epithelium. The activated protein disrupts the osmotic balance of these cells by forming pores in the cell membrane causing the cells to lyse (Van Rie et al., 1992). The gut becomes paralyzed and the insect stops feeding; most insects will die within a few hours of ingestion (Marrone and Macintosh, 1993). The binding affinity of these toxin fragments is often directly related to the toxicity, though binding does not assure toxicity (Whalon and McGaughey, 1998).

Bt Toxins

There are 34 recognized subspecies of B. thuringiensis – some of the most commonly used include subspecies kurstaki (against Lepidoptera), subspecies israelensis (against Diptera, primarily mosquitoes and blackflies), and subspecies tenebrionis (against Leptinotarsa decemlineata, the Colorado potato beetle) (Whalon and McGaughey, 1998). Two general groups of insecticidal crystal proteins made by this wide variety of subspecies have been identified, Cyt (cytolysins) and Cry (crystal delta-endotoxins). Hofte and Whiteley (1989) define four class of Cry genes and two classes of Cyt genes. CryI and CryII toxins are active against lepidopterans, CryII and CryIV against dipterans, and CryIII against coleopterans (Hofte and Whiteley, 1989). While CryIII toxins are produced by subspecies tenebrionis and tolworthi and CryIV by israelensis, generally very little correlation between certain toxins and certain subspecies exists. Cry toxins bind to specific receptors on cells in the insect midgut. Cyt genes are active against dipteran and coleopteran pests, and additionally have shown action against hemipterans (true bugs) and dictyopterans (roaches and termites) (Frutos et al., 1999; Gould and Keeton, 1996). Cyt toxins, unlike Cry toxins, do not recognize specific binding sites (Lereclus et al., 1993).

Insecticide Resistance

While Bt is very unlike other insecticides in its origin, mode of action, and use, it still shares some of the problems of any insecticide. One major problem with insect control via insecticides is the evolution in insects of resistance to those insecticides.

The first reported cases of insecticide resistance to early synthetic insecticides occurred over 50 years ago. About thirty years later, in 1979, the United Nations Environmental Programme declared pesticide resistance one of the world’s most serious environmental problems. Its seriousness to the environment stems from problems of human nutrition due to crop loss, spread of disease by resistant insects, addition to the environment of new and potentially dangerous insecticides after resistance has developed, and application of greater and greater amounts of chemicals to which pests have already gained resistance (Pimentel and Burgess, 1985).

How Resistance Develops

Insects can and have developed resistance to nearly every type of insecticide. Resistance to other insecticides is, in fact, one of the many reasons Bacillus thuringiensis has come into common use today. Insecticide resistance develops due to genetic variation in large insect populations. A few individuals in the original insect population are unaffected by a given insecticide. Generally, unaffected (resistant) individuals differ from affected (susceptible) individuals either in the nature of the insecticide’s target molecules in the insect, or in the method the insect uses to break down toxin molecules (Michaud, 1997). When the insecticide is applied, individuals who are unaffected by it are those who survive to pass their genes on to following generations. Over time, a greater and greater proportion of the insect population is unaffected by the insecticide. In addition to the damage done by the increasingly large number of surviving, resistant insects, attempts to control insecticide resistance can indirectly cause secondary pest outbreaks that do yet more crop damage (Hoy, 1998). Secondary outbreaks will be addressed further later in this paper.

Factors Affecting the Development of Resistance

There are several factors that increase the rate at which insecticide resistance is generally developed. Some factors are related to the insect population itself: species with higher reproductive rates, shorter generation times, greater numbers of progeny, and larger, more genetically varied local populations develop a large resistance population more quickly (Pimentel and Burgess, 1985). Whether the genetic basis of insect resistance is dominant or recessive is also of importance (Wearing and Hokkanen, 1995). Other factors are dependent upon the insecticide. Resistance develops more rapidly to more persistent insecticides; their staying power in the environment increases the chance that susceptible individuals are exposed to the toxin and die, thus not passing on their insecticide-susceptible traits to the next generation. This selects more strongly on resistant insects because only the resistant insects thrive. By similar logic, frequent application of non-persistent insecticides has the same effect (Wood, 1981). Insect populations with little immigration into the gene pool of new, non-exposed susceptible individuals also develop resistance more readily (Comins, 1977). Populations that have in the past been exposed to an insecticide with a mode of action similar to that of a new insecticide are quick to develop resistance to the new toxin. This phenomenon is known as cross-resistance (Pimentel and Burgess, 1985).

The Development of Resistance to Bacillus thuringiensis

Why Is This So Important?

As noted above, insecticide resistance is a major problem – not only in agriculture, but also in health and economics. The development of resistance to Bacillus thuringiensis toxins is, however, particularly unfortunate. Bt toxins are more pest-specific and environmentally safe than conventional pesticides, yet as effective against problem insects. For these reasons, commercial spray formulations of Bt are available to organic farmers and are one of their most valuable biocontrol tools. If Bt-based products become ineffective due to resistance, organic farmers will have lost this irreplaceable resource (McGaughey et al., 1998).

Reports of Resistance to Bt

In 1985, the first evidence of resistance developing in the field against Bt delta-endotoxins was published. Low levels of resistance were found in Plodia interpunctella, the Indianmeal moth, in storage bins of Bt-treated grain. This inspired a laboratory study of resistance to Bt in P. interpunctella. It was shown that in conditions like that in grain storage areas, Bt resistance could be developed in the Indianmeal moth in less than one storage season. Prior to this, Bt delta-endotoxin resistance had been seen in neither the field nor the lab, though attempts were made to select for resistance in laboratory populations (McGaughey, 1985).

Recognition of the potential of the Bt resistance problem became greater when the first reports of high resistance to Bt toxins in the field came in 1990 from Hawaii, Florida, and New York in the United States – thirty years after its commercial debut here. The species found to be losing susceptibility to Bt toxin was Plutella xylostella, the diamondback moth, treated with spray formulations of the toxins. At about that same time, resistance was detected in P. xylostella after intensive use in several other countries, including Japan, China, the Philippines, and Thailand (Liu and Tabashnik, 1997). Malaysia also reported Bt resistance in the diamondback moth in 1990; interviews with local farmers confirmed their personal experiences with this unfortunate situation (Iqbal et al., 1996). Thus far P. xylostella is still the only insect species in which very considerable resistance has been found to develop outside of the laboratory.

In the fifteen years since Bt resistance was discovered in P. interpunctella, Bt resistance has been selected in laboratory populations of a total of thirteen insect species. Eleven of these species have developed resistance to various strains of Bt toxin in the laboratory but not in the field: Ostrinia nubilalis (the European corn borer), Heliothis virescens (the tobacco budworm), Pectinophora gossypiella (the pink bollworm moth), Culex quinquefasciatus (the mosquito), Caudra cautella (the almond moth), Chrysomela scripta (the cottonwood leaf beetle), Spodoptera exigua (the beet armyworm), Spodoptera littoralis (the Egyptian cotton leafworm), Trichoplusia ni (the tiger moth), L. decemlineata (the Colorado potato beetle), and Aedes aegypti (the yellow fever mosquito) (Huang et al., 1999; Gould et al., 1997; Liu et al., 1999; Tabashnik et al., 1994; Wirth et al., 1997; Frutos et al., 1999; Whalon and McGaughey, 1998). Many other species have been tested in the lab but retained susceptibility to Bt (Whalon and McGaughey, 1998). While none of the species listed here has yet developed resistance in the field, these laboratory studies show that the potential to develop resistance is real. These results should be of no less concern than those of the 1985 and 1990 findings regarding P. interpunctella and P. xylostella.

Mechanism of Resistance

Central to learning to curb resistance to Bt is understanding the mechanism by which an insect resists the toxins. Mechanisms by which insects resist the lethal effects of B. thuringiensis toxins are, naturally, closely related to the mode of action of Bt. As stated earlier, Bt protoxins are activated by proteases in the insect midgut; after activation they bind to receptors on the epithelium. Thereafter a number of steps lead to the death of the insect. The specifics of the mode of action are complex and varied among insect and Bt strains, so complex in fact that prior to 1985 it was thought that the complexity itself would prevent the evolution of resistance (Whalon and McGaughey, 1998). Mechanisms of resistance are equally complex. Because so many steps are involved in the full process of Bt’s mode of action, many ways of stopping the process and resisting the toxin are possible. The actual mechanism of resistance has only been closely studied in a few species in which resistant populations have been selected for in the lab. In even fewer have results been very conclusive.

Thus far studies have most commonly shown the resistance mechanism to involve a change in the membrane receptors to which activated Bt toxins bind. In P. xylostella, this reduced binding of the toxin is the only known resistance mechanism (Tabashnik et al., 1997). A 1992 study indicated that resistance to Bt CryIAb toxins correlated with a reduction in the number of CryIAb receptors in the midgut (van Rie et al., 1992). P. interpunctella appears to also have reduced binding, with a 50-fold decrease in CryIA binding associated with a 100-fold decrease in toxicity (van Rie et al., 1990). There do not seem to be fewer binding sites developing in P. interpunctella, simply less binding affinity (van Rie et al., 1992). In addition to decreased affinity, resistance in P. interpunctella is linked with the absence of a major gut proteinase. Presumably this proteinase is associated with the proteolytic cleavage and activation of Bt protoxins (Oppert et al., 1997).

Both of the above methods of resistance, reduced toxin binding/binding sites and decreased activation of the toxin, are thought to occur together in H. virescens (Michaud, 1997). Studies involving resistance in H. virescens have been inconclusive, however. A 1991 study indicated reduced CryIA binding affinity and increased number of CryIA binding sites in resistant populations; a contradictory study indicated increased CryIA binding affinity and decreased number of CryIA binding sites in resistant populations (MacIntosh, 1991; Tabashnik 1994b).

Finally, Choristoneura fumiferana, the spruce budworm, shows a completely different resistance mechanism: CryIAa toxins are deactivated by precipitation with a protein complex present in the midgut (Michaud, 1997).

Resistance Management

The Goals and Types of Resistance Management

Insecticide resistance has come to be, thanks to the lessons of history, recognized by many as an unavoidable outcome of insecticide use. The goal of what is known as “resistance management” is, then, not to stop resistance entirely, but to slow its development and extend an insecticide’s useful lifespan as long as possible (Comins, 1977). Some scientists have come to refer to this science not as resistance management, but as “resistance mitigation,” as it more appropriately describes the nature of our control of the resistance problem (Hoy, 1998).

It will be necessary to counter resistance in order to preserve the efficacy of Bt. There are three goals of resistance management: avoiding resistance where and if possible, delaying resistance as long as possible, and making resistant populations revert to susceptibility (Croft, 1990). Several possible resistance programs have been conceived in the past 25 years, most of which could potentially be used in conserving susceptibility to Bt. The transgenic plant forms of Bt, the use of which is on the rise, are especially prone to resistance development. Transgenic plants expose insects to toxins continually, even at times when they are not causing economic damage (Mallet and Porter, 1992).

Resistance management programs generally use one of just three basic approaches to delay resistance. One approach seeks to minimize exposure to toxins and/or allow for mating between resistant insects and a large population of susceptible insects, to keep susceptible traits continuing in the gene pool. These strategies include tissue-specific and time-specific expression of toxins, mixtures, mosaics, rotations, refuges, and occasional release of susceptible males into the field. Another approach focuses on combining pest-control techniques and is based on the assumption that an insect is more likely to develop resistance to just one type of control than more than one type of control simultaneously. Strategies in this category include gene stacking, high doses, combinations of toxins with completely different modes of action, and combinations of low toxin dose and natural enemies. The final approach is very different in nature from those listed above. This strategy employs “trap plants” to lure pests away from productive crops.

Keeping a Susceptible Population for Mating with Resistant Individuals

The Release of Susceptible Insects Into an Exposed Population

Among the oldest strategies are those involving the mating of resistant with susceptible insects. The simplest of these ideas is the periodic release of susceptible males, raised in the lab or collected elsewhere, into a local, Bt-treated population. This would theoretically make it possible to keep the frequency of resistance in a population below a predefined level (Curtis, 1981). This method is best used on populations of insects such as mosquitoes, in which insecticides generally target females (Wood, 1981). However, Bt is not a gender-specific pesticide, and there is a risk that many of the susceptible males released would die in the Bt field before mating. Additionally, the feasibility of rearing and transporting large colonies is very questionable.

Refugia

Based on the simple strategy described above, several resistance programs involve the nearby placement of a susceptible population which, it is hoped, will diffuse into the treated population to mate. This is the basis of the refuge strategy. Refugia can vary in size and placement, and serve as a reservoir of susceptible insects. Ideally, many susceptible individuals will mate with few resistant individuals, creating an overall very low rate of resistance in following generations. The success of refugia depends upon four conditions: that the resistance trait is recessive, that there is random mating, that adults will travel sufficiently between toxic plants, and that there is complete lack of insecticidal action in the refugia. If adults do not travel between the refuge and treated area, resistance will quickly develop in treated areas, while susceptible populations continue to mate amongst themselves in untreated areas. If refugia are subjected to any sort of insecticide, the available susceptible population available for mating with Bt-exposed individuals will decrease (Tabashnik, 1997).

Refugia placed alongside treated areas and external to the field are more successful than rows of refuge plants planted in with rows of Bt plants. The larger the refugium, the longer resistance is delayed (Frutos, 1999). However, resistance will eventually develop when migration of resistant individuals into the susceptible population finally brings the resistant population of the untreated area to too high a fraction to sustain the equilibrium refuges are designed to create (Comins, 1977). A refuge area covering five to ten percent of the total crop area was recommended in a 1992 study (Mallet and Porter, 1992). Other studies confirmed this figure. Computer modeling using information about a lepidopteran life cycle indicated that a crop area with 10% refuge content delays resistance for five to 1120 generations (Tabashnik, 1994a). A program with ten percent refugia was also shown to help P. xylostella maintain susceptibility to Bt subspecies aizawai. Other species, however, did not necessarily respond well to refugia of this size (Liu and Tabashnik, 1997).

While the refuge strategy is successful in a general sense, the correlation between what occurs in laboratory studies and what happens in the ever-variable field is not easy to predict. In 1999, a study indicated that random mating might not necessarily be a safe assumption of insects in the field. Bt-resistant P. gossypiella populations took an average of 5.7 more days to develop fully than susceptible populations. Because more than 80% of P. gossypiella mate within three days of hatching and die soon thereafter, this favors assortative, not random, mating. Susceptible individuals will mate with each other before the resistant individuals even hatch (Liu et al., 1999). Of course, the degree to which this affects P. gossypiella refuge strategy in the field depends on generation overlap and perhaps on other factors as well. Another factor that is rather difficult to predict is the migration of adults when finding a mate. This information is helpful in deciding where to place refugia.

The refugia strategy is imperfect, but its successes are well backed by data. It is important to remember that the susceptible population is a resource that can be depleted (Wood 1981). The refuge method is very often used in combination with other strategies to enhance the effectiveness of both. An unfortunate problem is the potential unwillingness of farmers to set aside area for refugia, which is then not usable cropland. This is a difficult decision to make when competing with other farmers who are not using refugia (Mallet and Porter, 1992).

Seed Mixtures

Seed mixtures, like refugia, act to delay resistance by maintaining a susceptible population for mating. A field planted using this strategy will result in a random mix of Bt and toxin-free plants. Two major studies investigated the efficacy of seed mixtures in comparison with refugia. Mallet and Porter (1992) used computer modeling to show that seed mixtures actually hastened the development of resistance in comparison with fields of solely toxic plants (Mallet and Porter, 1992). Two years later, a contradictory study found that seed mixtures are preferable to pure Bt fields (Tabashnik, 1994a). Both studies agree that refugia are more successful than seed mixtures and, in many cases, even more successful when combined with seed mixtures in a single program (Mallet and Porter, 1992; Tabashnik, 1994a).

The reason for the poor performance of seed mixtures as a resistance strategy is closely related to theoretical advantage of seed mixtures. While individuals from susceptible plants can more easily travel to and mate with Bt-exposed and resistant individuals, the ease of travelling for individuals also puts susceptible insects at risk for too much Bt exposure from neighboring plants. Working in favor of seed mixtures is the potential behavioral pattern of some insect species to preferentially choose toxin-free over Bt plants, thus cutting down on feeding exposure of susceptible insects to Bt. This behavior has been observed in laboratory populations of H. virescens. However, we know neither the extent of this behavior in the field nor the propensity of other species toward this behavior (Mallet and Porter, 1992).

Mosaics

Mosaics are like refugia in that they involve a patchwork of Bt areas and non-Bt areas. However, mosaics are different in that the non-Bt areas, instead of being untreated, are treated with some other insecticide – a different strain of Bt or something completely unrelated. For this strategy to be effective, it is important that there is no cross-resistance between the two insecticides (meaning resistance to one of them cannot be positively correlated with or helpful in developing resistance to the other) (Tabashnik, 1994b). In fact, negative cross-resistance is preferable (Hoy, 1998). The model, like the refuge strategy, relies on the assumption of sufficient migration of insects between the patches of the mosaic. The mosaic strategy has seen little experimental evaluation, but models have suggested that it is not useful because one ends up simultaneously selecting for resistance to both toxins rather than delaying resistance to either (Tabashnik, 1994b; Frutos et al., 1999). Only one early study provides any support for the mosaic model. This study showed that if a mosquito population is evenly distributed across a mosaic of two equally effective insecticides, and if immigration and emigration rates are equal for all plots, then mosaics can delay the development of insecticide resistance (Curtis, 1981).

Rotations

An alternative to the mosaic strategy is the rotation method. Here, a pattern of two or more insecticides is arranged temporally rather than spatially. Using two or more insecticides sequentially rather than simultaneously has been shown to delay resistance relatively (Wood, 1981). Like mosaics, rotations cannot be successful if cross-resistance is present between the insecticides. The premise of rotations is that, because there is a fitness cost associated with resistance, that whenever the population is not exposed to a certain insecticide, the frequency of resistance to that insecticide will drop (Hoy, 1998). Unfortunately, it has been shown that after removal of selection pressure for resistance to Bt (during periods where Bt is in use), the frequency of resistance in the population remains stable or only decreases very slowly (Tabashnik, 1994b). Additionally, fitness costs associated with resistance have generally been negligible (Hoy, 1998). Thus, there would probably not be considerable pressure selecting for Bt-susceptible individuals during the non-Bt periods of the rotation.

Tissue-specific and Time-specific Toxin Expression

Tissue-specific and time-specific toxin expression are management strategies which seek to minimize Bt overexposure by engineering transgenic plants to express toxin genes only at times when needed or only in parts of the plant which are most economically important or most vulnerable (Frutos et al., 1999). This is similar to refugia and seed mixtures in that the plant itself provides its own refuges (Mallet and Porter, 1992). Plants could be developed to only produce toxins after a certain threshold of damage. Or, in tissue-specific expression, studies on the interactions between a particular pest and its target plant could suggest strategies to develop. For instance, cotton plants attacked by boll worms could produce toxin only in young boll tissues, as this is the most important part of the plant. In addition to specifically protecting the critical plant tissue, this would affect only one generation of boll worms (that which lives at the time the boll is young), avoiding the constant selection pressure that hastens evolution of resistance (Gould, 1988). Tissue-specific expression would not be a viable option for some pests which target nearly the entire plant, such as the European corn borer (Monsanto, 2000). For most species, not enough is known about field behavior, and therefore effective methods of carrying out tissue-specific or time-specific expression of Bt are not yet known. Because specific toxin expression, as noted above, provides refuge in a way similar to seed mixtures, the same negative results could result with use of specific expression as have resulted with seed mixtures.

Combining Insect Control Methods

The next general approach to resistance management combines elements of both old and new ideas. It is assumed that resistance is less likely to evolve to two control methods simultaneously than to only one method. Thus, with two methods, resistance to the combination will be delayed more than using each individually in a temporal or spatial arrangement. Using two or more control methods at a time is like having insurance in case one of them begins to lose efficacy. Similar to this is a method of heightening effectiveness of a toxin by using a high dosage – like having one dose as the first control method and a second dose as the other, though the underlying theory is different.

Use of Multiple Insecticides

The first of these strategies is a simple combination of two or more insecticides on the same field, such as a Bt and non-Bt toxin. Necessary to this approach is a lack of cross-resistance between the two. However, it is difficult to assume that no degree of cross-resistance will be present between any two insecticides, as it has been shown to occur even between some insecticides with completely different modes of action (Hoy, 1998). No experimental evidence indicates that use of insecticide combinations in lethal doses delays resistance to Bt (Tabashnik, 1994b). However, some work with synergists looks somewhat promising. Synergists are methods that are not necessarily lethal or toxic alone by when used with an insecticide increase the toxicity of that insecticide. Serine protease inhibitors increase toxicity of Bt against some species, but there is no evidence that this greater toxicity brings with it lessened resistance development (Tabashnik, 1994b).

Gene Stacking

Gene stacking is like the combination method explained above, but involves only combinations of two or more different Bt Cry toxins, or a Bt Cry and a Bt Cyt toxin. The toxins are delivered simultaneously but each recognizes a different binding site in the insect midgut (Frutos et al., 1999). This strategy is based on the same assumptions and criteria as the combination method. Some studies show Cry and Cyt toxins working together, with Cyt toxins as a synergist. In a 1998 study, use of Bt CytIAa helped overcome 5000-fold resistance to Bt CryIIIAa toxins in C. scripta (Frutos et al., 1999). Similarly, sublethal doses of CytA toxins, when combined with CryIV toxins, appear to suppress and/or reduce up to 1000-fold resistance in CryIV-resistant populations of C. quinquefasciatus (Wirth et al., 1997). However, considerable evidence works against any strategy using combinations of solely Cry toxins. H. virescens shows cross-resistance to many strains of Bt Cry toxins; P. xylostella and P. interpunctella in the field and the laboratory readily evolve resistance to up to five or six Bt Cry toxins simultaneously (Tabashnik, 1994b). This may be partially explained by the 1997 finding that a single gene in P. xylostella is responsible for resistance to four individual Bt Cry toxins (Tabashnik et al., 1997). However, with the inclusion of external refugia, this strategy seems more effective than mosaics or rotations of different Cry toxins. It must be noted that for even the small amount of resistance delay the gene stacking strategy may provide, refugia are necessary (Caprio, 1998).

Combining Bt with Natural Enemies

Another potential strategy uses natural enemies, instead of other toxins, in combination with low doses of Bt. This strategy cannot be used with most insecticides, as they have broad-range effects and will kill natural enemies as well as pests (Chilcutt and Tabashnik, 1997). Natural enemies used can be predators or parasitoids (parasites which kill their prey). The assumption underlying this strategy is one earlier mentioned as part of the rotation strategy. Theoretically, the fitness costs of resistance will make resistant pests more susceptible to attack by natural enemies (Frutos et al., 1999). Any natural enemy used must, of course, be unaffected by exposure to Bt. Already, as part of many integrated pest management programs, low doses of Bt are used in conjunction with natural enemies when Bt is not strong enough in the preferred dosage to control the pest population. A 1997 study investigated the toxic effects of a combination of Bt and the parasitoid Cotesia plutella on P. xylostella. The interaction of the two did not significantly affect mortality of P. xylostella populations resistant to Bt, though Bt-susceptible populations were affected by the combination more than by either the parasitoid or Bt used alone (Chilcutt and Tabashnik, 1997). However, no studies in the field or the laboratory have actually shown a slowing of resistance development by using a Bt and natural enemy combination. As mentioned previously, Bt resistance has not been shown to have a detectable fitness cost. Additionally, there is no guarantee that a natural enemy will prefer to attack Bt-resistant pests, even if they do show detectable fitness costs. If natural enemies attack susceptible and resistant pests, or prefer susceptible pests, they may even hasten resistance development when used with Bt (Frutos et al., 1999).

High Dose

The high dose strategy hopes to delay resistance by using a high enough dosage of toxin to kill heterozygous insects – those with alleles for both resistance and susceptibility. Insects with a resistance allele have varying degrees of resistance depending on whether they are carrying one or two alleles for resistance. Theoretically, an increasingly higher dose of toxin will kill insects with an increasing degree of resistance, with a high enough dose killing even those individuals who are resistant homozygotes (Tabashnik, 1994b). A dose high enough to be 100% lethal is, however, not feasible. Therefore the high dose strategy aims to kill just heterozygotes (and will also kill individuals homozygous for the susceptibility allele).

The success of the high dose strategy depends on rare and recessive or partially recessive resistance alleles (Huang et al., 1999). Controlling the dose carefully is also key to using this strategy. If the dose is allowed to degrade, heterozygotes will survive and pass on the resistance gene. Additionally, heterozygotes must have no advantage over insects homozygous for the susceptibility allele, or the development of resistance can be hastened (Curtis 1981).

In use, the high dose strategy must be combined with refugia to provide susceptible individuals for mating with the resistant individuals that remain after high dose application. In addition, using a combination of toxins helps delay resistance further. Consideration of problems with the refugia and toxin combination strategies must be taken even when these are used in combination with the high dose strategy (Frutos et al., 1999).

Resistance has generally been found to be a recessive or partially recessive trait, though some evidence indicates that some resistance alleles may be dominant or codominant (Tabashnik et al., 2000). In one study, resistance to Dipel ES, a spray formulation of Bt using toxins of subspecies Berliner, appears to be an incompletely dominant trait in O. nubilalis (Huang et al., 1999). This evidence undermines the usefulness of the high dose strategy. However, no similar results have been found in studies of resistance to transgenic plants, which use different toxins (Tabashnik et al., 2000). Another potential problem with the high dose strategy involves natural enemies. If pest populations are eliminated temporarily, natural enemies in the ecosystem which fed upon these pests may leave or die. This leaves room later for a secondary pest outbreak when pests return and the population is not controlled naturally by enemies (Hoy, 1998).

To achieve high doses, synthetic development or enhancement of Bt toxins is possible. Another method recently studied involves expression of toxins in different parts of the cell. With tobacco plants engineered to express CryIIAa2 toxins in the chloroplasts, resistance was dramatically reduced in populations of three different species already showing significant resistance to nuclear expression of CryIIAa2 (Kota et al., 1999).

Trap Plants

Trap Plants

The trap plant strategy is somewhat similar to refugia, but differs in theory and is therefore in a category all its own. Andow and Alstad developed this theory in 1995, using computer simulations of the behavior of O. nubilalis. This strategy uses transgenic Bt crops, not spray formulations. A patch of Bt crop is planted as a trap, and a non-Bt productive crop is grown nearby. The Bt patch is planted earlier and matures earlier than the crop, luring in insects, which are then killed by the toxin. Later, the crop matures, but insects have been controlled already by the Bt trap (Andow and Alstad, 1995). This strategy is optimized for O. nubilalis because the moths emerge in June, an early part of the growing season, after overwintering as mature larvae. At this time they prefer the most mature plants available for laying eggs, which are the trap plants. Unfortunately, computer simulations with species other than O. nubilalis showed failure of this strategy. The strategy also would not be successful with species such as H. virescens which, as mentioned earlier, have shown some avoidance of Bt plants when given a choice between toxin and toxin-free plants (Frutos, et al., 1999).

Current Issues in Resistance Management

Currently Used Strategies

After a glance through documents on the subject from the United States Environmental Protection Agency (EPA) it seems apparent that the focus of management strategies and most concern is on Bt transgenic crops and not sprays. More attention has been focused in the area because of the consistent nature of the exposure to the toxin insects receive from Bt plants, which produce toxin during the entire growing season. Furthermore, Bt genes, unlike formulations developed in private laboratories, are considered part of the public domain and therefore worthy of extra protection (USEPA, 1999a).

Current management programs focus on a high-dose/refugia strategy, usually used in combination with gene stacking. Sometimes mosaics or rotations are used in place of or as well as refugia. The EPA releases new standards for Bt crops each growing season to the producers of Bt crops, such as Monsanto and DeKalb.

For Bt field corn in the 2000 growing season, the EPA recommends 20% refugia in areas where Bt cotton is not grown. In areas where Bt cotton is grown, a 50% refugia is necessary. Refugia must not be treated with any insecticide. However, this requirement can be waived at a certain economic threshold. At the threshold, growers are allowed to apply a non-Bt insecticide to refuges, resulting in a management strategy more closely resembling mosaics. In addition to regulations regarding size and treatment of refugia, the EPA specifies placement of refugia. Untreated external refugia must be within half a mile of the Bt field; treated external refugia must be within a quarter of a mile of the Bt field. Growers may also choose to plant refugia as strips within a Bt field. Bt and non-Bt strips must be arranged in alternating increments of six rows each (Anderson, 1999). Decisions regarding placement of the refuge within the field or externally can be made based on what is known about the travel habits and mating behavior of target species (USEPA, 1999a). Specific varieties of Bt corn producing the Cry1Ac toxin cannot be planted in areas where Bt cotton using the same toxin are grown. This is due to special concerns of resistance development in the corn earworm, a pest which targets both cotton and corn, sometimes within the same growing season (USEPA, 1999b). Bt sweet corn does not have structured refuge requirements, as the early harvest of this crop is thought to eliminate the field presence of potentially resistant larvae (USEPA, 1999a).

Similarly, growers of Bt potatoes must plant a 20% refuge. Potato growers are also advised to grow Bt potato fields as far as possible from the location of the previous year’s Bt fields. Bt cotton refuges are less conservative, with a 4% non-treated refuge or a 20% treated refuge plan available. If a grower lives within an area where Bt cotton accounts for 75% of the total cotton acreage, he must plant his refuges within a mile of his Bt fields (USEPA, 1999a). Controversy exists over these requirements, however. Bt cotton targets three cotton pests, the tobacco budworm, pink bollworm, and cotton bollworm. The toxin dose produced by the plant is high enough to effectively kill all heterozygous tobacco budworms and pink bollworms, but is only a moderate dose for the cotton bollworm. Without the resistance protection of a high toxin dose, cotton bollworms will very quickly lose susceptibility to Bt. Many endorse more conservative management regulations for Bt cotton to attempt to counter this problem (USEPA, 1999b).

In addition to the above, some farmers use rotation strategies, although this is not regulated or required. Rotations can alternate hybrid varieties of the same crop, or different crops altogether (Benbrook, 1999). Rotating crops is acceptable in combination with the other strategies outlined in the EPA regulations above, but is not considered sufficient alone.

Refuge requirements have become stricter over time as more is learned about the effects of the strategy on the large scale – outside of the computer models, laboratories, and small plots where refugia are first designed. Future plans also aim toward stricter standards. For instance, the EPA plans further specifications for Bt potato growers, quantifying the distance refuges must be from the toxin field and the distance one year’s Bt potato field must be from the previous year’s Bt potato field. EPA’s future plans for cotton growers include six to ten percent larger refuge areas, but the refugia may still be treated with non-Bt insecticides, making this more of a mosaic strategy (USEPA, 1999b).

Education and Compliance Issues

Any resistance management strategy can only be successful if it is properly implemented. It is crucial that growers understand resistance strategies, what the EPA mandates or recommends, and how to follow through with these tactics. Monitoring of practices and education are left to Bt producers, as the EPA does not have the resources to monitor all Bt growers. Producers then report compliance to the EPA (USEPA, 1999a).

Abuse of refugia is commonly reported or suspected. Some refuges that are meant to be untreated are sprayed with insecticides. The success of refuges can also be undermined by growers who plant them on their poorest land and give them less care than their other plants, making them potentially less attractive to insects (USEPA, 1999b).

Planting Bt crops when they are not needed also presents a problem. Bt corn, for instance, provides protection against the European corn borer. Significant outbreaks of this pest only occur about one in five years. Because it is difficult to predict when outbreaks will happen, some growers plant Bt too often. Some growers report that the genetics they desire in corn are only available in Bt varieties, so they plant Bt even though the toxin is not necessary (Benbrook, 1999).

Education of growers has not been as successful in all cases as is necessary to keep refugia within standards. Some growers have reported not receiving any information regarding the need for refugia or how to plant them. Others are confused by conflict between planting requirements from Bt producers, which requires tilling of old stalks, and local soil conservation guidelines. Clearly this indicates a need for improvement in communication. Growers should also be educated to monitor their crops for success or failure of transgenic crops for their particular needs (Benbrook, 1999).

Conclusion

Management Strategies

Many recurring themes are evident in the above discussion of management strategies. A study or studies showing that the method will fail accompanies every strategy. Disclaimers are made on nearly every study that field conditions are unpredictable, and that there is not enough information on the behavior of pests to ensure that they will comply with our models. Much research has been done in the areas of resistance inheritance, pest behavior within the ecosystem, frequency of resistance alleles in natural populations, toxin cross-resistance, and even economic impacts of these strategies. However, it still has not been nearly enough to guarantee or even successfully predict what will happen in the field. In this sense, none of the strategies given, even those currently in use, are completely acceptable if we are serious about preserving Bt as a safe and effective insect control method.

The Problem with Transgenic Bt Crops

Several benefits can be cited for the use of transgenic crops. Labor is cut, because the insecticide does not need to be reapplied several times per season. The effects of weathering on the insecticide are lessened, and parts of the plant that cannot be reached by foliar sprays, such as the inside of the stalk, are covered. The benefits which are most often stressed however, are the following: the whole plant is covered, and the toxin is produced for the entire season (Monsanto, 2000). As far as resistance development is concerned, these are not benefits, and are in fact the complete opposite.

As evidenced by the continually growing refuge requirements, it is very difficult to compensate for the damage that can be done to toxin efficacy by continually exposing pests and their environment to that toxin. Additionally, there will eventually be a point at which growers are not willing to accept the yield losses that can result from growing refugia. Cutting yield losses by allowing insecticide treatment of refugia is not an answer, as it only decreases the available population of Bt--susceptible insects. The wisest management strategy will not just attempt to provide susceptible insects that may or may not choose to mate with resistant insects, but reduce the toxin exposure of all insects to minimize selection pressure as much as possible while still remaining an effective control tool.

Current transgenic Bt products do not appear to be part of the solution to the resistance management problem. Two solutions exist which involve the presence of Bt toxins only when needed. One involves further engineering of plants, the other involves less engineering of plants.

One solution is time-specific expression. As mentioned in the discussion on this method, there are no currently developed methods for expressing toxin at only certain times. The best time-specific expression of toxin would be expression only in damaged tissues. Plants undamaged by pests would not produce toxin. Exposure to toxin would occur only when necessary.

The other solution is to use Bt only as a foliar spray. The benefits of transgenic Bt crops are not significant enough to make up for the toxin overexposure caused. Bt is just as effective when properly applied in spray form. Because it is so specific, Bt would not even need to be applied extremely often, and in fact it would be best if it were not. This contradicts the argument that Bt crops significantly cut labor. Growers could apply the toxin only when potentially economically damaging outbreaks of target pests occur and only in those areas where they occur. Organic and small-scale growers have been employing this technique successfully for years.

The Future

Future research should focus on the gaps in our knowledge listed above, especially regarding pest behavior and the characteristics of insect resistance that we do not yet fully understand. This will lead to more reliable resistance programs. Care must also be taken that growers are fully aware of the details of these resistance programs and what they should do to ensure their crops are in compliance.

Research into time-specific expression should also continue. Combined with the above research on behavior and resistance, this will help us decide if time-specific expression is a successful way to mitigate resistance. Because no methods have yet been developed to express Bt only when tissues are damaged, more research into this is also necessary.

An understanding of Bacillus thuringiensis can aid us in finding more bacteria that might have similar applications in agriculture. Other soil bacteria are the most likely to have developed the insecticidal characteristics of Bt. The discovery of more bacteria with the effectiveness and safeness of Bt, or the discovery of ways to implement currently-known bacteria as effective insecticides would be very fortunate, and an insurance for the future, when resistance to Bt does develop.

In conclusion, biotechnology can be used for or against our advantage in resistance management. Further research can tell us which way to go. In the meanwhile, the EPA and Bt producers and growers should be conservative and careful in creating, enforcing, complying with, and monitoring current resistance strategies. Bt is nearly an ideal pesticide and the loss of its use would be an extremely unfortunate occurrence.

Alstad, D. N. and Andow, D. A. 1995. Managing the evolution of insect resistance to transgenic plants. Science 268: 1894-1896.

Anderson, J. L. 1999. Letter to Bt corn registrant. Available at: http://www.epa.gov/oppbppd1/biopesticides/otherdocs/bt_corn_ltr.htm

Baum, J. A., Johnson, T. B., and Carlton, B. C. 1999. Bacillus thuringiensis: Natural and recombinant bioinsecticide products. In: Biopesticides: Use and Delivery, Hall, F. R. and Menn, J. J., Eds., Humana Press, Totowa, NJ, 189-210.

Benbrook, C. 1999. EPA’s plant pesticide refuge program: Questions from the Chicago workshop. Available at: http://www.biotech-info.net/meeting_report.html.

Caprio, M. A. 1998. Evaluating resistance management strategies for multiple toxins in the presence of external refuges. J. Econ. Entomol. 91: 1021-1031.

Chilcutt, C. F. and Tabashnik, B. E. 1997. Independent and combined effects of Bacillus thuringiensis and the parasitoid Cotesia plutellae (Hymenoptera: Braconidae) on susceptible and resistant diamondback moth (Lepidoptera: Plutellidae). J. Econ. Entomol. 90: 397-403.

Comins, H. N. 1977. The development of insecticide resistance in the presence of migration. J. Theor. Biol. 64: 177-197.

Croft, B. A. 1990. Developing a philosophy and program of pesticide resistance management. In: Pesticide Resistance in Arthropods, Roush, R. T. and Tabashnik, B. E., Eds., Chapman and Hall, New York, NY, 277-296.

Curtis, C. F. 1981. Possible methods of inhibiting or reversing the evolution of insecticide resistance in mosquitoes. Pestic. Sci. 12: 557-64.

Ely, S. 1993. The engineering of plants to express Bacillus thuringiensis delta-endotoxins. In: Bacillus thuringiensis, An Experimental Biopesticide: Theory and Practice, Entwistle, P. F., Cory, J. S., Bailey, M. J., and Higgs, S., Eds., John Wiley & Sons, Chichester, UK, 105-124.

Frutos, R., Rang, C., and Royer, M. 1999. Managing insect resistance to plants producing Bacillus thuringiensis toxins. Critical Reviews in Biotechnology 19: 227-276.

Gould, F. 1998. Evolutionary biology and genetically engineered crops. BioScience 38: 26-33.

Gould, F., Anderson, A., Jones, A., Sumerford, D., Heckel, D. G., Lopez, J., Micinski, S., Leonard, R., and Laster, M. 1997. Initial frequency of alleles for resistance to Bacillus thuringiensis toxins in field populations of Heliothis virescens. Proc. Natl. Acad. Sci. USA 94: 3519-23.

Gould, J. L. and Keeton, W. T. 1996. Biological Science (6th Ed.) New York: W. W. Norton and Company.

Hofte, H. and Whiteley, H. R. 1989. Insecticidal crystal proteins of Bacillus thuringiensis. Microbiol. Rev. 53: 242-255.

Hoy, M. A. 1998. Myths, models and mitigation of resistance to pesticides. Phil. Trans. R. Soc. Lond. B. 353: 1787-95.

Huang, F., Buschman, L. L., Higgins, R. A., and McGaughey, W. H. 1999. Inheritance of resistance to Bacillus thuringiensis toxin (Dipel ES) in the European corn borer. Science 284: 965-967.

Iqbal, M., Verkerk, R. H. J., Furlong, M. J., Ong, P. C., Rahman, S. A., and Wright, D. J. 1996. Evidence for resistance to Bacillus thuringiensis (Bt) subsp. kurstaki HD-1, Bt subsp. aizawai and Abamectin in field populations of Plutella xylostella from Malaysia. Pestic. Sci. 48: 89-97.

Kegley, S.E. and Wise, L.J. 1998. Pesticides in fruit and vegetables. Sausalito, CA: University Science Books.

Kota, M., H. Daniell, S. Varma, S. F. Garczynski, F. Gould, and W. F. Moar. 1999. Overexpression of the Bacillus thuringiensis (Bt) Cry2Aa2 protein in chloroplasts confers resistance to plants against susceptible and Bt-resistant insects. Proc. Natl. Acad. Sci. USA 96: 1840-5.

Lereclus, D., Delecluse, A., and Lecadet, M.-M. 1993. Diversity of Bacillus thuringiensis toxins and genes. In: Bacillus thuringiensis, An Environmental Biopesticide: Theory and Practice, Entwistle, P. F., Cory, J. S., Bailey, M. J., and Higgs, S., Eds., John Wiley & Sons, Chichester, UK, 37-69.

Liu, Y. B. and Tabashnik, B. E. 1997. Experimental evidence that refuges delay insect adaptation to Bacillus thuringiensis. Proc. R. Soc. Lond B 264:605-10.

Liu, Y. B., Tabashnik, B. E., Dennehy, T. J., Patin, A. L., and Bartlett, A. C. 1999. Development time and resistance to Bt crops. Nature 400:519.

Mallet, J. and P. Porter. 1992. Preventing insect adaptation to insect-resistant crops: are seed mixtures or refugia the best strategy? Proc. R. Soc. Lond. B 250: 165-9.

MacIntosh, S. C., Stone, T. B., Jokerst, R. S., and Fuchs, R. L. 1991. Binding of Bacillus thuringiensis proteins to a laboratory-selected line of Heliothis virescens. Proc. Natl. Acad. Sci. USA 88: 8930-8933.

Marrone, P. G. and MacIntosh, S. C. 1993. Resistance to Bacillus thuringiensis and Resistance Management. In: Bacillus thuringiensis, An Environmental Biopesticide: Theory and Practice, Entwistle, P. F., Cory, J. S., Bailey, M. J., and Higgs, S., Eds., John Wiley & Sons, Chichester, UK, 221-235.

McGaughey, W. H. 1985. Insect resistance to the biological insecticide Bacillus thuringiensis. Science 229: 193-195.

McGaughey, W. H., Gould, F., and Gelernter, W. 1998. Bt resistance management. Nat. Biotech. 16: 144-6.

Michaud, D. 1997. Avoiding protease-mediated resistance in herbivorous pests. Trends Biotech. 15: 4-6.

Miller, G.T. 1998. Living in the environment (10th ed.). Belmont, CA: Wadsworth.

Monsanto. 2000. The whole plant. The whole season. Available at: http://www.monsanto.com/monsanto/agriculture/current_products/yieldgard_ipcorn/whole_plant_season.html.

Oppert, B., Kramer, K. J., Beeman, R. W., Johnson, D., and McGaughey, W. H. 1997. Proteinase-mediated insect resistance to Bacillus thuringiensis toxins. J. Biol. Chem. 272: 23473-23476.

Pimentel, D., and Burgess, M. 1985. Effects of Single Versus Combinations of Insecticides on the Development of Resistance. Environ. Entomol. 14: 582-9.

Tabashnik, B. E. 1994a. Delaying insect adaptation to transgenic plants: seed mixtures and refugia reconsidered. Proc. R. Soc. Lond. B 255: 7-12.

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

Tabashnik, B. E. 1997. Seeking the root of insect resistance to transgenic plants. Proc. Natl. Sci. USA 94: 3488-3490.

Tabashnik, B. E., Liu, Y.-B., Finson, N., Masson, L., and Heckel, D. G. 1997. One gene in diamondback moth confers resistance to four Bacillus thuringiensis toxins. Proc. Natl. Acad. Sci. USA 94: 1640-1644.

Tabashnik, B. E., R. T. Roush, E. D. Earle, and A. M. Shelton. 2000. Resistance to Bt toxins. Science 287: 42.

Tabashnik, B. E., Finson, N., Groeters, F. R., Moar, W. J., Johnson, M. W., Luo, K., and Adang, M. J. 1994. Reversal of resistance to Bacillus thuringiensis in Plutella xylostella. Proc. Natl. Acad. Sci. USA 91: 4120-4.

Tabashnik, B. E., Liu, Y.-B., Malvar, T., Heckel, D. G., Masson, L., Ballester, V., Granero, F., Mensua, J. L., and Ferre, J. 1997. Global variation in the genetic and biochemical basis of diamondback moth resistance to Bacillus thuringiensis. Proc. Natl. Acad. Sci. USA 94: 12780-12785.

United States Environmental Protection Agency. 1999a. EPA and USDA position paper on insect resistance management in Bt crops. Available at: http://www.epa.gov/oppbppd1/biopesticides/otherdocs/bt_position_paper_618.htm.

United States Environmental Protection Agency. 1999b. Meeting summary: EPA-USDA Bt crop insect resistance management workshop. Available at: http://www.epa.gov/oppbppd1/biopesticides/summary826.htm.

Van Frankenhuyzen, K. 1993. The challenge of Bacillus thuringiensis. In: Bacillus thuringiensis, An Environmental Biopesticide: Theory and Practice, Entwistle, P. E., Cory, J. S., Bailey, M. J., and Higgs, S., Eds., John Wiley & Sons, Chichester, UK, 1-35.

Van Rie, J., McGaughey, W. H., Johnson, D. E., Barnett, B. D., and van Mellaert, H. 1990. Mechanism of insect resistance to the microbial insecticide Bacillus thuringiensis. Science 247: 72-74.

Van Rie, J., Van Mellaert, H., and Peferoen, M. 1992. Mechanism of insect resistance to Bacillus thuringiensis in Plodia interpunctella and Plutella xylostella. In: Molecular mechanisms of insecticide resistance: diversity among insects, Mullin, C. A. and Scott, J. G., Eds., American Chemical Society, Washington, DC.

Wearing, C. H. and Hokkanen, H. M. T. 1995. Pest resistance to Bacillus thuringiensis: ecological crop assessment for Bt gene incorporation and strategies of management. In: Biological control: benefits and risks, Hokkanen, H. M. T. and Lynch, J. M., Eds., Cambridge University Press, Cambridge, UK, 236-252.

Whalon, M. E. and W. H. McGaughey. 1998. Bacillus thuringiensis: Use and Resistance management. In: Insecticides with novel modes of action: mechanism and application, Ishaaya, I. and Degheele, D., Eds., Springer, Berlin, 106-137.

Wirth, M. C., Georghiou, G. P., and Federici, B. A. 1997. CytA enables CryIV endotoxins of Bacillus thuringiensis to overcome high levels of CryIV resistance in the mosquito, Culex quinquefasciatus. Proc. Natl. Acad. Sci. USA 94: 10536-40.

Wood, R. J. 1981. Strategies for conserving susceptibility to insecticides. Parasitology 82: 69-80.

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This paper was written as a thesis for the completion of the degree of Bachelor of Arts in Environmental Studies at the University of Chicago.

My thanks to Profs. Ted Steck and Joy Bergelson for guiding me through the Environmental Studies Program and advising the creation of this paper, respectively.
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The Environmental Studies Program

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