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Pesticide Resistance: Strategies and Tactics for Management. 1986. National Academy Press, Washington, D.C. Genetics and Biochemistry of Insecticide Resistance in Arthropods Prospects for the Future . FREDERICK W. PLAPP, JR. Insecticide resistance in the housefly has a fairly simple genetic basis. There is one gene for decreased uptake of insecticides, one gene for target-site resistance to each insecticide type, and one major gene for metabolic resistance to all insecticides. The last interacts with minor genes located elsewhere in the genome. Based on limited data, resistance patterns are similar in other species. Evidence is presented that target-site resistance to pyrethroidsl DDT and to cyclodienes is controlled by changes in regulatory genes determining the number of receptor protein molecules synthesized. Resistance in both is recessive to susceptibility. The product of the major gene for metabolic resistance appears to be a receptor protein that recognizes and binds insecticides and then induces synthesis of appropriate detoxifying enzymes. Different types of enzymes, for example, oxidases, esterases, and glutathione transferases, are coordinately induced. The effect of the gene is qualitative, that is, it determines the specific form of detoxifying enzyme synthesized. Inheritance is codominant. Possible solutions to resistance include using synergists such as chlordimeform, which appear to act by increasing the binding of pyrethroid insecticides to their target-site proteins; using agonists, which successfully compete with insecticides for recognition by the receptor protein; and using either mixtures of insecticides or insec- ticides composed of multiple isomers. INTRODUCTION Resistance to insecticides in arthropods is widespread (Georghiou and Mellon, 1983), with at least 400 species resistant to one or more insecticides. 74

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INSECTICIDE RESISTANCE IN ARTHROPODS 75 In some species, populations are resistant to nearly every insecticide ever used to control them and, often, to related chemicals to which the population has never been exposed. Resistance, at least in the house fly, has a fairly simple and straightforward genetic basis. Extensive genetic studies in other species, most notably Lucilia cuprina and Drosophila melanogaster, have indicated a similar situation. The biochemistry of resistance is also compre- hensible, particularly when there is an adequate understanding of the genetics of resistant populations. GENETIC MECHANISMS CONFERRING RESISTANCE A very important question is, How many genes for resistance are there? Are there multiple genes for resistance, each conferring resistance to a narrow range of insecticides, or are there only a few genes, each conferring resistance to a wide array of insecticides? If there are numerous genes then cross- resistance associated with each gene should be limited, and new insecticides would solve the problem. Conversely, if a limited number of genetic mech- anisms is involved, then resistant populations should show resistance to insecticides to which they have never been exposed. The second hypothesis is more frequently true. Thus, developing new insecticides that are closely related to existing insecticides in either mode of action or pathways of me- tabolism will not solve the problem. If only a few major genes confer resistance to insecticides, it should be possible to characterize the mechanisms controlled by each gene. Once this is done, it may be possible to devise solutions and regain our ability to deal with populations recalcitrant to chemical control. Standard neo-Darwinian models (Moore, 1984) suggest that change occurs as a result of accumulation of multiple mutations, each mutation contributing a minute amount to the total; that is, insecticide resistance should be poly- genic, but it is not (Whitten and McKenzie, 1982~. In field populations resistance is almost invariably due to a single major gene. Therefore, standard evolutionary theory does not seem to apply to the development of resistance. A regulatory gene hypothesis is a more likely model to account for change, particularly at the population or subspecific level. Such genes, which control time and nature of expression of structural genes, are more likely to provide the genetic basis of adaptive variation such as the development of resistance (Levin, 19841. In my opinion, available data on resistance offer considerable support for Levin's hypothesis. In this paper I shall summarize both genetic and biochemical evidence that changes in regulatory genes are of major importance in insecticide resistance. Two types of regulatory genes seem to be present, and both differ in inheritance and biochemistry. One type exhibits all-or-none inheritance (fully dominant or recessive) and appears to involve changes in the amount of

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76 MECHANISMS OF RESISTANCE TO PESTICIDES protein synthesized. The second shows codominant (intermediate) inheritance and involves changes in the nature of proteins synthesized. Quantitative resistance (that type involving differences in amount of proteins synthesized) is similar in nature to certain bacterial operons. Resistance of this type apparently involves regulatory elements located adjacent to the structural genes in question. Change does not occur in the structural gene, but in an adjacent, distinct, genetic element. If it were in the structural gene, inheritance would be additive. Since it is not, the evidence is that a separate protein (i.e., the product of a distinct gene) must be the site of variation. Regulators of this type have been defined as "near" regulators (Paigen, 19791. The second type, qualitative resistance, appears to represent a mechanism allowing for production of altered forms of particular detoxifying enzymes in resistant as compared to susceptible insects. Genetic studies with the house fly (Plapp, 1984) show that change at a single genetic locus appears to control resistance associated with multiple detoxification enzymes. A similar mech- anism can be inferred from earlier studies with D. melanogaster (Kikkawa, 1964a,b). Since one locus appears to act on a variety of enzymes, the gene probably is not adjacent to the enzymes whose activity it regulates. Such regulators have been defined as "distant" regulators (Paigen, 1979), and such systems can be considered "regulons" (Plapp, 19841. According to Paigen, these systems are characterized by their codominant inheritance rather than the all-or-none type of similar bacterial systems. GENETICS OF RESISTANCE The number of major genes conferring resistance to insecticides in the house fly (and presumably other species) is limited. The list of known re- sistance genes includes: pen for decreased uptake of insecticides. This chromosome III gene is inherited as a simple recessive. By itself, pen confers little resistance to any insecticide, seldom more than two- to three-fold. It appears to be more important as a modifier of other resistance genes. In such cases pen may double resistance levels, for example, from 50- to 100-fold. kdr- for knockdown resistance to DDT and pyrethroids. This gene is a chromosome III recessive at a locus distinct from pen. It confers resistance to DOT and all analogs and to pyrethrins and all synthetic analogs. Low- level (kdr) and high-level (super kdr) alleles have been reported. The gene probably involves modifications at the target site of the insecticides. dld-r for resistance to dieldrin and all other cyclodienes. This is a chromosome IV gene whose inheritance is incompletely recessive. Resistance appears to involve change at the target site of these insecticides. AChE-R for altered acetylcholinesterase, the target site for organo

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INSECTICIDE RESISTANCE IN ARTHROPODS 77 phosphate (OP) and carbamate insecticides. The gene is located on chro- mosome II and is inherited as a codominant. Different alleles appear to confer different levels of resistance to multiple organophosphate and carbamate insecticides (Oppenoorth, 19821. The house fly's metabolic resistance to many types of insecticides, in- cluding OPs, carbamates, pyrethroids, DDT, and juvenile hormone analogs, is associated with a gene or genes on chromosome II. This type of resistance was long thought to be due primarily to mutations in structural genes for the specific enzymes. Earlier work had shown that resistance genes were located at a variety of loci on chromosome II (Hiroyoshi, 1977; Tsukamoto, 19831. More recent work (Wang and Plapp, 1980; Plapp and Wang, 1983) suggests that inversions or other rearrangements of the chromosome are present in many resistant strains and are of sufficient extent to explain the apparent differences in gene location on the chromosome, that is, only one gene seems to be present, but it is not always located at the same place relative to other genes on chromosome II Based on these results the idea of multiple structural genes for metabolic resistance on chromosome II becomes more tenuous, and the idea of a common resistance gene becomes more logical. Close linkage (and, therefore, possible allelism) exists among genes for metabolic resistance to insecticides in other insect species as well. Examples include the gene RI (for resistance to insecticides) located at 64.5-66 on chromosome II of Drosophila melanogaster, a locus conferring resistance to organophosphates, carbamates, and DDT (Kikkawa, 1964a,b), and major genes for metabolic resistance to diazinon and malathion in numerous pop- ulations of Lucilia cuprina (Hughes et al., 19841. Other evidence for allelism has been reported for malathion resistance in different populations of Tri- bolium castaneum (R. W. Beeman, U.S. Department of Agriculture, Man- hattan, Kansas, personal communication, 1983~. In fact, our knowledge of the genetics of resistance in insects other than dipterans is so inadequate that we can only guess as to the precise nature of the genetic mechanisms involved. Research has shown that resistance to different classes of insecticides is associated with a particular linkage group, but the number of genes involved is unknown. Genetically, the most feasible approach to this problem is to perform allelism tests. This method has demonstrated allelism of genes for reduced uptake of insecticides (pen) in American and European house fly populations (Sawicki, 1970) and for organophosphate resistance in spider mites (Ballantyne and Harrison, 19671. I have recently been doing such tests on several house fly strains with metabolic resistance to various organo- phosphates associated with chromosome II and with chromosome II resistance to DDT and organophosphates within a strain. All data indicate allelism of the genes. Although chromosome II has been shown to make a major contribution

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78 MECHANISMS OF RESISTANCE TO PESTICIDES to metabolic resistance in the house fly, minor genes on other chromosomes make additional contributions. An assay of total levels of resistance is made by crossing resistant strains with susceptible strains containing mutant mark- ers on multiple chromosomes. Recent work in my laboratory has shown that the contribution to metabolic resistance of chromosomes other than II is not expressed in the absence of chromosome II and is inherited as incomplete recessives. Such resistance is similar in inheritance to that described previ- ously for pen, kdr, and dId-r. Position also affects the expression of resistance associated with chro- mosome II. Strains showing a major (20 to 30 percent) reduction in recom- bination values between the resistance gene and the mutation carnation eye (car) have increased levels of resistance, compared with strains showing smaller reductions in recombination values (Plapp and Wang, 19831. Thus, the location of the gene on chromosome II is important in determining the level of resistance present. In summary four types of resistance, pen, kdr, dld-r, and metabolic, associated with chromosomes other than II, are inherited as incompletely or fully recessive characters. In contrast, altered acetycholinesterase resistance and metabolic resistance on chromosome II are inherited as codominants. The level of resistance associated with the major chromosome II gene for metabolic resistance varies with the location of the gene on the chromosome. BIOCHEMISTRY OF RESISTANCE This area has been intensively studied for the last 30 years. Earlier work concentrated on mechanisms associated with metabolic resistance and iden- tified a number of enzyme systems concerned with resistance (Tsukamoto, 1969; Oppenoorth, 1984~. Recent studies have dealt with mechanisms in- volved in nonmetabolic (target site) resistance. The availability of genetic stocks purified to contain individual mechanisms proved invaluable to these studies. High-affinity receptors for DDT and pyrethroids are present in insects (Chang and Plapp, 1983a,c). House flies possessing the gene kdr for target- site resistance bound less insecticide than susceptible flies. Resistant flies had fewer target-site receptors than susceptible flies (Chang and Plapp, 1983b). Further, binding affinity between preparations from R and S strains did not differ. Therefore, the major difference between strains was strictly quanti- tative, that is, in receptor numbers, and not qualitative, that is, in receptor affinity. Similar studies on cyclodiene mode of action/mechanism of resistance have been reported by Matsumura and coworkers. Kadous et al. (1983) reported that cyclodiene-resistant cockroaches were cross-resistant to the plant-derived neurotoxicant picrotoxinin and, further, that nerve components

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INSECTICIDE RESISTANCE IN ARTHROPODS 79 from resistant cockroaches bound significantly less [3H] a-dihydropicrotox- inin than similar preparations from susceptible insects. The receptor was sensitive to all cyclodiene insecticides (Tanaka et al., 19841. Similar studies with susceptible and cyclodiene-resistant house flies have shown reduced binding in resistant insects (K. Tanaka and F. Matsumura, Michigan State University, East Lansing, Michigan, personal communication, 1984), sug- gesting that the number of receptor binding sites is decreasing. Thus, quantitative decreases in numbers of target sites may be involved in target-site resistance to both DDT/pyrethroids and cyclodienes. At first glance it may appear contradictory for resistant insects to have fewer target- sites than susceptible insects. Decreased receptor numbers probably confer resistance by a needle-in-the-haystack approach (Lund and Narahashi, 1981a,b); the decrease in number may make it less likely for a toxicant to reach target- sites. Decreases in target-site numbers are consistent with the genetics of resis- tance to these insecticides. If the change were in the target-sites themselves, inheritance would be additive; R/S heterozygotes would be intermediate be- tween the parents in resistance. Inheritance being all-or-none agrees with the idea of quantitative change. The specific mutations conferring resistance are probably in genes coding for proteins that determine the number of target- site proteins synthesized. Here, heterozygotes would have the normal number of receptors since the diffusible protein product of the wild-type regulatory gene would act on both structural genes. Only the resistant homozygotes, those with two mutant genes, would produce fewer target-site receptor pro- teins than normal. This activity is an example of bans dominance; the protein product of a regulatory gene influences the expression of a specific structural gene on both members of a chromosome pair. The precise biochemical mechanism of the major gene for metabolic re- sistance to insecticides is not yet known with certainty, although a single gene locus is probably involved. Since all structural genes coding for de- toxification enzymes are probably not at the same site, a common controlling mechanism might be responsible. The key to metabolic resistance is induction. Induction of different de- toxifying enzymes is coordinate (Plapp, 1984~; that is, exposure to chemicals that induce one detoxifying enzyme induces several. Mixed-function oxi- dases, glutathione transferases, and DDT dehydrochlorinase are coordinately induced in the house fly (Plapp, 1984), as are oxidases and glutathione transferases in Spodoptera (Yu, 19841. When the products of several struc- tural genes (enzymes) respond to the same stimulus, they must be responding to the protein product of a separate gene, a genetic element that is distinct from the elements that define the enzymes themselves. The finding is not original. It comes from the research of Monod and Jacob on induction in E. coli. As reviewed by Judson the critical idea in

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80 MECHANISMS OF RESISTANCE TO PESTICIDES

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INSECTICIDE RESISTANCE IN ARTHROPODS Insecticides oPs Carban,~1es - \ Pyrethroids DOT JHAs Others ~\ Detoxifying Enzymes Resistance Gene MFOs /~ Single Gene, |_ Chromosome ~ Bomb Gene Product, a /~'0<, OCR for page 74
82 MECHANISMS OF RESISTANCE TO PESTICIDES are designed to counter. Target-site synergism may exist, however, and in at least one case the use of such a synergist has blocked the development of resistance. Several years ago we reported that the miticide-ovicide chlordimeform was found to be strongly synergistic with several hard-to-metabolize insecticides, including toxaphene and DDT, to which resistance was present in the tobacco budworm (Plapp, 1976; Plapp et al., 19761. Since then chIordimeform syn- ergism has been reported in the new, metabolically stable synthetic pyreth- roids (Plapp, 1979; Rajakulendren and Plapp, 19821. Many formamidines are synergistic with pyrethroids and other insecticides against several ar- thropod species (El-Sayed and Knowles, 1984a,b). The mechanism for this synergism may be that chlordimeform is acting as a target-site synergist (Chang and Plapp, 1983c). Chlordimeform may block pyrethroid resistance in Heliothis (Crowder, et al., 19841. Selection of H. virescens with permethrin resulted in 37-fold resistance within a few generations. Parallel selection with permethrin-chlor- dimeform combinations prevented resistance development. Therefore, limited data are available suggesting that chlordimeform may synergize insecticides against insects in cases of target-site resistance and block development of such resistance. Since the new synthetic pyrethroids will probably be subject to kdr-type resistance, the use of such combinations offers a possible way to manage the problem. Metabolic resistance has been attacked by a variety of approaches, pri- marily the use of synergists designed to poison the enzymes involved in detoxification. Since the work described in this paper indicates that a single gene is of primary importance in this resistance, different approaches may be possible. Rather than poisoning the detoxifying enzymes, it may be pos- sible to affect the receptor protein by using agonists that compete with in- secticides for recognition sites on xenobiotic receptor proteins. This idea may already have been demonstrated. Ranasinghe and Georghiou (1979) selected an organophosphate-resistant mosquito population with three regimens. These were temephos only, temephos plus the antioxidant synergist piperonyl butoxide, and temephos plus DEF. DEF, S,S,S-tributyl phospho- rotrithioate, is a plant defoliant that inhibits oxidases and esterases. I suggest that it is a receptor agonist. Selection with temephos resulted in the rapid development of a high level of resistance. The same thing occurred, but slightly slower, with temephos plus piperonyl butoxide. Selection with te- mephos plus DEF quickly restored a near-normal level of susceptibility to the test population. The authors were unable to offer an explanation for the results of the temephos/DEF selection. I believe that DEF has a high affinity for the receptor protein, which recognizes temephos as a xenobiotic. With the temephos/DEF selection the receptor protein increased its ability to recognize and bind DEF

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INSECTICIDE RESISTANCE IN ARTHROPODS 83 and simultaneously lost its ability to recognize, bind, and, thus, respond to temephos. Other work with DEF as a synergist has been done with Lucilia (Hughes, 19821. Preexposure to DEF significantly synergized diazinon, while simul- taneous exposure to DEF and diazinon was much less effective. Again the results agree with a receptor-level effect for DEF. Another approach to overcoming metabolic resistance involves using in- secticides composed of two isomers. The major example of this effect in- volves phenylphosphonates of the EPN series. These insecticides have four different substituents attached to the central phosphorus atom. They exist as plus and minus isomers. Insects with metabolic resistance to the more typical dialkyl phenyl phosphorothioates show little or no cross-resistance to the phenylphosphonates. The single gene hypothesis for metabolic resistance offers an explanation. If only one receptor gene is of primary importance in metabolic resistance, its protein product can recognize either the plus or the minus isomer, but not both at once. If this is so, then synthesis of enzymes of high specific activity toward only one isomer will be induced. An example of the use of two isomer organophosphates to circumvent resistance involves profenofos to control multiresistant populations of Spodoptera littoralis in Egypt (Dittrich et al., 19791. I have confirmed these findings of lack of resistance to the two isomer OPs in fly strains with metabolic resistance to single isomer OPs. It may be a general phenomenon. This idea may not be practical, however, because of the delayed neurotoxicity syndrome associated with at least some of these organophosphates (Metcalf and Metcalf, 19841. A final approach involves using multiple isomers of an insecticide. The idea is that the two will compete for the receptor protein just as the plus and minus isomers of the phenylphosphonates compete. I tested this idea by comparing the toxicity of dimethyl and diisopropyl isomers of parathion, alone and in combination, to susceptible and resistant house flies. Toxicities of the mixture were additive to susceptible flies, but synergistic with resistant flies. These results suggest that using mixed alkyl isomers of dialkyl phen- ylphosphates and phosphorothioates might prove quite effective for over- coming resistance. Again the mechanism responsible may be the lack of ability of a single resistance gene to handle multiple chemicals simulta- neously. CONCLUSION Resistance genetics in the house fly is comparatively simple. The studies described here would not have been possible without the availability of mutant stocks to identify different chromosomes and to map resistance gene locations on specific chromosomes. Such studies are currently not feasible with most resistant species, due to lack of mutant markers. Nevertheless, what is true

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84 MECHANISMS OF RESISTANCE TO PESTICIDES for house flies and other higher Diptera in the way of resistance genetics is probably true for other insects; that is, the genetic mechanisms involved are probably ubiquitous rather than specific. Based on the genetics, it is possible to develop a comprehensive theory of resistance. Resistance is best understood as being due to changes in reg- ulatory genes controlling the amount or nature of target proteins or enzymes synthesized. From this understanding, approaches to solving the problem become feasible, at least for metabolic resistance. Solutions involve using mixtures of insecticides or using insecticides composed of several isomers. The mixture approach will work because change at only a single locus is involved. Not all components of an insecticide need to be toxic; some may work primarily as receptor agonists rather than enzyme inhibitors. Nothing in the foregoing should be interpreted, however, as an opinion that resistance is subject to perfect and/or complete suppression via chemical means. I have no doubt that, in the long term, life will always overcome chemistry and find ways to persevere. The best that can be said is that if we are lucky, we should be able to suppress resistance to such an extent that we can live with it. REFERENCES Ballantyne, G. H., and R. A. Harrison. 1967. Genetic and biochemical comparisons of organo- phosphate resistance between strains of spider mites (Tetranychus species). Entomol. Exp. Appl. 10:231-239. Britten, R. J., and E. H. Davidson. 1969. Gene regulation for higher cells: A theory. Science 169:349-357. Chang, C. P., and F. W. Plapp, Jr. 1983a. DDT and pyrethroids: Receptor binding and mode of action in the house fly. Pestic. Biochem. Physiol. 20:76-85. Chang, C. P., and F. W. Plapp, Jr. 1983b. DDT and pyrethroids: Receptor binding and mechanism of knockdown resistance (kdr) in the house fly. Pestic. Biochem. Physiol. 20:86-91. Chang, C. P., and F. W. Plapp, Jr. 1983c. DDT and synthetic pyrethroids: Mode of action, selectivity, and mechanism of synergism in the tobacco budworm, Heliothis virescens (F.), and a predator Chrysopa carnea Stephens. J. Econ. Entomol. 76:1206-1210. Crowder, L. A., M. P. Jensen, and T. F. Watson. 1984. Permethrin resistance in the tobacco budworm, Heliothis virescens. Pp. 223-224 in Proc. Beltwide Cotton Conf., Atlanta, Gal, January 9-12, 1984. Dittrich, V., N. Luetkemeier, and G. Voss. 1979. Monocrotophos and profenofos: Two organo- phosphates with a different mechanism of action in resistant races of the Egyptian cotton leafworm Spodoptera littoralis. J. Econ. Entomol. 72:380-384. El-Sayed, G. N., and C. O. Knowles. 1984a. Formamidine synergism of pyrethroid toxicity to two- spotted spider mites (Acari: Tetranychidae). J. Econ. Entomol. 77:23-30. El-Sayed, G. N., and C. O. Knowles. 1984b. Synergism of insecticide activity to Heliothis zea (Boddie) by formanilides and formamidines. J. Econ. Entomol. 77:872-875. Georghiou, G. P., and R. B. Mellon. 1983. Pesticide resistance in time and space. Pp. 1-46 in Pest Resistance to Pesticides, G. P. Georghiou and T. Saito, eds. New York: Plenum. Hallstrom, I. P. 1984. Cytochrome P4so in Drosophila melanogaster: Activity, Genetic Variation and Regulation. Ph.D. dissertation. University of Stockholm, Sweden.

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INSECTICIDE RESISTANCE IN ARTHROPODS 85 Hiroyoshi, T. 1977. Some new mutants and revised linkage maps of the house fly, Musca domestica L. Jpn. J. Genet. 52:275-288. Hughes, P. B. 1982. Organophosphorus resistance in the sheep blowfly, Lucilia cuprina (Wiede- mann) (Oiptera: Calliphoridae): A genetic study incorporating synergists. Bull. Entomol. Res. 72:573-582. Hughes, P. B., P. E. Green, and K. G. Reichmann. 1984. A specific resistance to malathion in laboratory and field populations of the Australian sheep blowfly, Lucilia cuprina. J. Econ. En- tomol. 77:1400-1404. Judson, H. F. 1979. The Eighth Day of Creation. New York: Simon and Schuster. Kadous, A. A., F. Matsumura, J. G. Scott, and K. Tanaka. 1983. Difference in the picrotoxinin receptor between the cyclodiene-resistant and susceptible strains of the German cockroach. Pestic. Biochem. Physiol. 19:157-166. Kikkawa, H. 1964a. Genetical analysis on the resistance to parathion in Drosophila melanogaster. II. Induction of a resistance gene from its susceptible allele. Botyu-Kagaku 2:37-41. Kikkawa, H. 1964b. Genetical studies on the resistance to Sevin in Drosophila melanogaster. Botyu- Kagaku 29:42-46. Levin, B. R. 1984. Science as a way of knowing Molecular evolution. Am. Zool. 24:451-464. Lund, A. E., and T. Narahashi. 1981a. Modification of sodium channel kinetics by the insecticide tetramethrin in crayfish giant axons. Neurotoxicology 2:213-229. Lund, A. E., and T. Narahashi. 1981b. Kinetics of sodium channel modification by the insecticide tetramethrin in squid axon membranes. Pharmacol. Exp. Ther. 219:464-473. Metcalf, R. L., and R. A. Metcalf. 1984. Steric, electronic, and polar parameters that affect the toxic actions of O-alkyl, O-phenyl phosphorothionate insecticides. Pestic. Biochem. Physiol. 22:169-177. Moldenke, A. F., and L. C. Terriere. 1981. Cytochrome P450 in insects. 3. Increase in substrate binding by microsomes from phenobarbital-induced houseflies. Pestic. Biochem. Physiol. 16:222- 230. Moore, J. A. 1984. Science as a way of knowing Evolutionary biology. Am. Zool. 24:467-534. Nebert, D. W., M. Negishi, M. A. Lang, L. M. Hjelmeland, and J. J. Eisen. 1982. The Ah locus, a multigene family necessary for survival in a chemically adverse environment: Comparison with the immune system. Adv. Genet. 21:1-52. Oppenoorth, F. J. 1982. Two different paraoxon-resistant acetylcholinesterase mutants in the house fly. Pestic. Biochem. Physiol. 18:26-27. Oppenoorth, F. J. 1984. Biochemistry of insecticide resistance. Pestic. Biochem. Physiol. 22:187- 193. Ottea, J. A., and F. W. Plapp, Jr. 1981. Induction of glutathione S-aryl transferase by phenobarbital in the house fly. Pestic. Biochem. Physiol. 15:10-13. Ottea, J. A., and F. W. Plapp, Jr. 1984. Glutathione S-transferase in the house fly: Biochemical and genetic changes associated with induction and insecticide resistance. Pestic. Biochem. Physiol. 22:203-208. Paigen, K. 1979. Acid hydrolases as models of genetic control. Annul Rev. Genet. 13:417-466. Phillips, I. R., E. A. Shephard, B. R. Rabin, R. M. Bayney, S. F. Pike, A. Ashworth, and M. R. Estall. 1983. Factors controlling the expression of genes coding for drug-metabolizing enzymes. Biochem. Soc. Trans. 11 :460-463. Plapp, F. W., Jr. 1976. Chlordimeform as a synergist for insecticides against the tobacco budworm. J. Econ. Entomol. 69:91-92. Plapp, F. W., Jr. 1979. Synergism of pyrethroid insecticides by formamidines. J. Econ. Entomol. 72:667-670. Plapp, F. W., Jr. 1984. The genetic basis of insecticide resistance in the house fly: Evidence that a single locus plays a major role in metabolic resistance to insecticides. Pestic. Biochem. Physiol. 22:194-201.

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86 MECHANISMS OF RESISTANCE TO PESTICIDES Plapp, F. W., Jr., and T. C. Wang. 1983. Genetic origins of insecticide resistance. Pp. 47-70 in Pest Resistance to Pesticides, G. P. Georghiou and T. Saito, eds. New York: Plenum. Plapp, F. W., Jr., L. G. Tale, and E. Hodgson. 1976. Biochemical genetics of oxidative resistance to diazinon in the house fly. Pestic. Biochem. Physiol. 6:175-182. Rajakulendran, S. V., and F. W. Plapp, Jr. 1982. Synergism of five synthetic pyrethroids by chlordimeform against the tobacco budworm and a predator, Chrysopa carnea. J. Econ. Entomol. 75: 1089-1092. Ranasinghe, L. E., and G. P. Georghiou. 1979. Comparative modification of insecticide-resistance spectrum of Culex pipiens fatigans Wied. by selection with temephos and temephos/synergist combinations. Pestic. Sci. 10:502-508. Sawicki, R. M. 1970. Interaction between the factor delaying penetration of insecticides and the desethylation mechanism of resistance in organophosphorus-resistant house flies. Pestic. Sci. 1 :84- 87. Tanaka, K., J. G. Scott, and F. Matsumura. 1984. Picrotoxinin receptor in the central nervous system of the American cockroach: Its role in the action of cyclodiene-type insecticides. Pestic. Biochem. Physiol. 22:117-127. Tsukamoto, M. 1969. Biochemical genetics of insecticide resistance in the house fly. Residue Rev. 25:289-314. Tsukamoto, M. 1983. Methods of genetic analysis of insecticide resistance. Pp. 71-98 in Pest Resistance to Pesticides, G. P. Georghiou and T. Saito, eds. New York: Plenum. Wang, T. C., and F. W. Plapp, Jr. 1980. Genetic studies on the location of a chromosome II gene conferring resistance to parathion in the house fly. J. Econ. Entomol. 73:200-203. Whitten, M. J., and J. A. McKenzie. 1982. The genetic basis for pesticide resistance. Pp. 1016 in Proc. 3rd Australas. Conf. Grassl. Invert. Ecol., K. E. Lee, ed. Adelaide, Australia: S.A. Government Printer. Yamamoto, I., Y. Takahashi, and N. Kyomura. 1983. Suppression of altered acetylcholinesterase of the green rice leafhopper by N-propyl and N-methyl carbamate combinations. Pp. 579-594 in Pest Resistance to Pesticides, G. P. Georghiou and T. Saito, eds. New York: Plenum. Yu, S. J. 1984. Interactions of allelochemicals with detoxification enzymes of insecticide-susceptible and resistant fall armyworm. Pestic. Biochem. Physiol. 22:60-68.