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Molecular biology of insecticide resistance
Toxicology Letters ELSEVIER
Toxicology Letters 82183 (1995) 83-90
Molecular
biology of insecticide resistance R. Feyereisen
Department of Entomology,
Forbes 410, University of Arizona,
Tucson, AZ 85721, USA
Abstract The widespread use of insecticides has amounted to a large scale ‘experiment’ in natural selection of insects by chemicals of toxicological importance to humans. Specific examples in which the molecular basis of insecticide resistance has been studied in detail are presented here. The biochemical/physiological mechanisms of resistance can be categorized as target site insensitivity, increased metabolic detoxification and sequestration or lowered availability of the toxicant. These are achieved at the molecular level by: point mutations in the ion channel portion of a GABA receptor subunit (cyclodiene insecticides); point mutations in the vicinity of the acetylcholinesterase (AChE) active site (organophosphorus and carbamate insecticide resistance); amplification of esterase genes (organophosphorus and carbamate insecticides); mutations linked genetically to a sodium channel gene (DDT and pyrethroid insecticides); and yet uncharacterized mutations leading to the up-regulation of detoxification enzymes,
such as cytochrome P450 and glutathione S-transferases (many classes of insecticides). In several cases, the selection of a precisely homologous mutation has been observed in different insect species. Keyword: Pesticides; GABA gated Cl--channek Acetylcholinesterase; Na’ -channel; Cytochrome P450; Glutathione S-transferases; Esterases
1. Introduction
Perhaps the most serious consequence of widespread, indiscriminate use of insecticides is the development of insecticide resistance. More than 500 species of insects and mites are reported to have developed resistance to one or more classes of insecticides [l]. A number of agricultural pests and disease vectors are so resistant in some areas of the world that chemical control has become extremely difficult. The list of effective insecticides for the control of crop pests and disease vectors is rapidly shrinking. Meanwhile, fewer new insecticides are being introduced to the market, largely because of the high costs associated with research, development and registra0378-4274/95/$09.50 @ 1995 SSDI 0378-4274(95)03470-6
tion, and the prognosis of a limited effective lifespan of the new insecticide. The impact of insecticide resistance in toxicology is 2-fold: (1) the insecticide ‘treadmill’ has lead to the introduction of a huge arsenal of sometimes highly toxic chemicals in the environment, and several have become problem pollutants over the last 50 years; (2) the immediate reaction to insecticide resistance by pest control practitioners has traditionally been an increase in the dosage, thus compounding the problems of resistance and environmental contamination. The study of insecticide resistance at the molecular level has uncovered a number of ways in which animals can survive in toxic environments. The nature of the mutational events that
Elsevier Science Ireland Ltd. All rights reserved
84
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-M
lead to resistance in insects should not be expected to be intrinsically different from those that are responsible for the genetic variation of human responses to xenobiotics. Indeed, insects and vertebrates share many of the favorite targets for ‘insecticidal’ action and they share many of the metabolic detoxification systems. Selectivity of insecticides is achieved either when insects and vertebrates do not share a target (e.g. the selectivity of juvenile hormone agonists) or when they follow different metabolic pathways (e.g. the low vertebrate toxicity of malathion). Resistance marks a genetic change in response to selection. Individuals carrying genetic traits for coping with the chemically hostile environment survive and reproduce, thereby passing on these traits to their progeny. Continued selection pressure exerted by the insecticide rapidly increases the frequency of the genetic trait (resistance) in the population. I present here a brief overview of molecular mechanisms (types of mutations) that lead to insecticide resistance.
2. Physiological/biochemical mechanisms of resistance
and molecular
Genetic and biochemical studies have contributed immensely to our understanding of resistance. There are 2 ways in which organisms can become resistant to xenobiotics such as pesticides: either by modifying the effective dose of the pesticide available at the target site or by modifying the target site itself. Thus, classically recognized mechanisms of resistance such as behavioral resistance, reduced penetration or absorption, sequestration and detoxification all contribute to decrease the dose of the pesticide, whereas a decreased target site sensitivity or a modification of target site number will contribute to render a dose of pesticide ineffective. In-depth discussions of these mechanisms are given in Roush and Tabashnik [2]. Molecular mechanisms of resistance are classified according to the type of genetic change selected by the insecticide pressure: point mutations in structural genes, or modifications in the number or activity of genes.
3. Point mutations in structural genes: cyclodiene resistance and acetylcholinesterase insensitivity Nucleotide deletions, additions, or substitutions in the DNA may alter the amino acid sequence of a protein. Point mutations in the target sites of several herbicides and fungicides are known to confer pesticide resistance. Two such examples are also known in insects, and illustrate the power of Drosophila as a model system for insecticide resistance in general [3].
3.1. Point mutation in the Rdl gene of Drosophila [4] Elucidation of the molecular mechanisms of cyclodiene insecticide resistance in Drosophila followed a logical, ab initio approach that led to the cloning of the first invertebrate GABA receptor subunit. Field-collected populations of Drosophila were screened for dieldrin resistance. A homozygous strain with 4000 X resistance was isolated, and the resistance gene (Rdl) mapped genetically at 26 CM on the left arm of chromosome III. Deficiency mapping and irradiationinduced rearrangements allowed a more precise localization to the cytological region 66F. The resistance gene was then uncovered by P element-mediated transformation of flies with a cosmid clone obtained from a chromosomal walk in that region. The cosmid clone contained a susceptible copy of the Rdl gene, thus ‘rescuing’ susceptibility in resistant flies. Screening of a cDNA library with a fragment of this cosmid clone yielded a clone encoding a GABA receptor subunit. This confirmed earlier pharmacological evidence that had pointed to the picrotoxinin binding site on GABA, receptors as the likely neuronal target of cyclodiene insecticides. Expression of the cDNA in Xenopus oocytes demonstrated a functional, homo-oligomeric GABA-gated chloride channel. Sequencing of Rdl in resistant flies revealed a single common point mutation of Ala302 to Ser. This mutation is located in the second membrane spanning region of the channel subunit, lining the
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-90
chloride ion pore. Site-directed mutagenesis of the susceptible cDNA and expression in Xenopus oocytes showed that this mutation leads to channels insensitive to block by dieldrin. Armed with a molecular diagnostic, a number of dieldrin-resistant and susceptible strains were screened. The Ala”‘* to Ser mutation was absent in 122 susceptible strains, and present in 58 resistant strains of Drosophila melanogaster from around the world. The only observed fitness cost of this mutation is a temperature-sensitive phenotype, i.e. temporary paralysis at 38°C. Most remarkably perhaps, was the subsequent demonstration that an identical mutation is found in cyclodiene-resistant strains of a number of other insect species. In 2 species, the homologous Ala residue is mutated to Gly instead of Ser (Table 1). The molecular diagnostic for target site resistance to cyclodiene insecticides also allowed an accurate determination of resistance frequency in a relatively small samples of flies collected in the field. Resistance frequency was shown to be as high as 1%) even though wild populations of Drosophila in the US have apparently not been under cyclodiene selection pressure for many years [4]. Table 1 Conserved
M2 region
of the Rdl GABA
receptor
subunit
3.2. Point mutations in the Ace gene of Drosophila
Reduced sensitivity of acetylcholinesterase (AChE) is a well-characterized biochemical mechanism of insecticide resistance. Higher K,,, and lower k; values for organophosphorus and carbamate insecticides in resistant strains indicated structural alterations in the active site of the enzyme [5]. In turn, these alterations in protein structure may be attributed to changes in the amino acid sequence of the target protein. The structural gene, called Ace, coding for the AChE specifically expressed in the central nervous system of Drosophila melanogaster has been cloned, sequenced and extensively characterized. Because Ace is a single locus that was known a priori to be involved in resistance, a directed search for point mutations was justified. Comparisons of the coding sequences of the Ace gene of susceptible and resistant Drosophila strains revealed 5 point mutations at 4 positions in the resistant strains [6]. In one strain, mutations at all 4 positions are present in the same gene (Table 2). These point mutations were introduced in a minigene construct devoid of the carboxy terminus coding sequence. Thus expression of a soluble protein in Xenopus oocytes
in various
insect
species
Ala Arg Val Ala Leu Gly Val Thr Thr Residue
at the Ala’02 position
Drosophila melanogaster Drosophila simulans (2 alleles found) Aedes aegypti (mosquito) Periplaneta americana (cockroach) Musca domestica (house fly) Tribolium castaneum (flour beetle) Hypothenemus hampei (coffee berry borer) Bemisia tabaci (whitefly) Adapted
from
ffrench-Constant
[4].
x5
S-strain
R-strain
Ala (122 strains)
Ser (58 strains)
Ala
SeriGly
Ala
GUY
Ala
Ser
Ala
Ser
Ala
Ser
Ala
Ser
Ala
Ser
R. Feyereisen
86 Table 2 Point mutations
Drosophila S R Saltillo R Bygdea R Pierrefeu R MH19 Housefly R 77M R CH2 R 49R
in AChE
insensitive
I Toxicology Letters 82183 (1995) 83-90
to organophosphates/carbamates
Phe”‘
Ile”’
Ser
Val Val Thr
Gly _
“”
Ala Ala Ala _
Phe”’
Tyr _ _ Tyr
_ _ _ _
S Val”” Leu _ _
S, susceptible strain: R, resistant strain. Val”“, GlyZhZ, Phe3*’ and G~Y’~’ in the house fly correspond the Glu residue of the AChE catalytic triad. Adapted from Muter0 et al. [6]; Williamson [7].
to Val”“, Gly’“‘,
allowed a functional characterization of these mutations, either alone, or in some of the combinations found in resistant strains collected from the field. The results clearly indicated that multiple mutations caused higher levels of insensitivity to organophosphates and carbamates [6]. Multiple mutations in the same gene may result either from the accumulation of mutations, or from the recombination between genes carrying single mutations. This interesting problem has not been clearly resolved, but there are indications favoring the second possibility. Clearly, this would have important implications in resistance management, particularly with regard to migration, population size and fitness of the individual mutants in the absence of insecticide selection. Because the 3-dimensional structure of a model AChE, the electric organ enzyme from Torpedo californica, has been determined by Xray analysis, alignment of the Drosophila sequence has allowed a discussion of the possible influence of these mutations on substrate or inhibitor binding in the active site gorge of AChE. The enzyme has also been sequenced in other insect species, and recently the mutations conferring resistance in the house fly AChE have been determined [7]. Mutations at 4 sites were ob-
Gly”’ Ala Ala _
Phe”’ Tyr Tyr _
Gly”’ _ _ Ala
PhexhX and GIY~“~ m Drosophila. Gly”‘ is adjacent
to
served, 2 of which at residues homologous to those determined in Drosophila (Table 2). The evidence gathered to date on Rdl and Ace point mutations allows 2 interim conclusions. First, that Drosophila is a powerful and predictive model for major insect pest species. Second, that the ‘choice‘ of point mutations that lead to a modified sensitivity to xenobiotics upon selection is probably most limited in target molecules that are most conserved across taxonomical boundaries. This is also a conclusion that emerges from studies on herbicide resistance in photosystem II. 4. Gene amplification: to organophosphorus insecticides
esterases and resistance and carbamate
DNA amplification events create additional copies of chromosomal sequences (including functional genes) which can survive in either intra- or extrachromosomal forms. This phenomenon was first extensively documented in tumor cells and permanent cell lines, for instance methotrexate resistance resulting from the amplification of the target dihydrofolate reductase [8]. Initially regarded as abnormal events occurring in tissue cultures, DNA amplification is now amply documented in whole organisms where
R. Feyereisen
I Toxicology Letters 8.2183 (1995) 83-90
expression of the gene product must be faster than can be achieved by transcription from only a single copy of the gene [8]. At this point, it must be emphasized that DNA amplification per se does not bring about resistance. The amplified gene must be transcribed and translated (overexpressed) into functional protein products for expression of the resistance trait. Devonshire and Sawicki, working with organophosphorus-resistant populations of the aphid Myzus persicae, first invoked a simple model of ‘a succession of tandem duplications of the structural gene’ for an insecticide-associated esterase (esterase E4). In a series of progressively more resistant clones, progressive elevation of titers of E4 correlated with the degree of observed resistance. Esterase E4 (and the closely related FE4) confer broad resistance to insecticidal esters [9]. In vitro translation of poly(A)+RNA from 3 clones showed elevated levels of E4- or FECencoding mRNAs in the resistant clones. Direct evidence for an up to 64-fold amplification of the E4 esterase in M. persicae was obtained when a cDNA clone was used to probe the genomes of susceptible and various resistant aphid clones [9]. Reversion of resistance without loss of the amplified sequences involves in a still obscure way the DNA methylation status of the esterase genes. Evidence for DNA amplification of esterase genes was also obtained in a number of insecticide-resistant strains of the Culex pipienslquinquefasciatus complex. Resistant mosquitoes from around the world have been characterized over the last few years, revealing amplification of several closely related esterases (Table 3). Although migration was invoked as a major cause
Table 3 Amplified
esterases
in Culex pipienslquinquefasciatus
of the worldwide distribution of amplified esterase B2 [lo], it is equally likely that multiple amplification events have taken place. A molecular analysis of the amplification unit (or amplicon) of esterases in aphids and mosquitoes revealed that amplification of genomic sequences extends far beyond the esterase gene itself [9]. The Bl esterase of mosquitoes is a 2.8 kb gene found 250 times as an amplicon of approximately 30 kb, with a core of 25 kb. The E4 esterase of aphids is a 4.3 kb gene, amplified up to 64-fold as an amplicon of about 25 kb. This is consistent with observations on other xenobiotic-related amplification events in eukaryotes
PI. Because the amplified esterase can account for a very significant percentage of the total protein of the insect (up to 3% in most resistant aphids), is the amplified esterase merely a sponge that sequesters the insecticide? In other words, does resistance result from hydrolysis or sequestration of the insecticide? The kinetic constants (affinity, bimolecular rate of formation of the acylated enzyme, hydrolysis/ deacylation rate) for each esterase and each substrate need to be measured. A comparison of the aphid E4 and the mosquito A2/B2 esterases [12] indicated that hydrolysis was the major role of the aphid E4 esterase, whereas the mosquito A2/B2 esterases play a predominant sequestration role (Table 4). 5. Regulatory
changes in gene expression
DNA amplification is a mechanism whereby overabundance of a gene product is achieved by a multiplication of the gene itself, but there are molecular mechanisms responsible for increased
mosquitoes Chlorpyrifos
Origin
Esterase
Copies
Esterase
North America Africa. Asia, N. America France France Cyprus
Bl A2 B2 Al A4 B4 B5
250x 60x _
500X _
800x _
70x 50x 500x
100x 6.6X 95x
Adapted
from
[9-111.
25x 250x
87
levels
resistance
R. Feyereisen
88
Table 4 Esterase amplification in insects: sequestration pmol/insect
Mosquito A2/B2 Culex quinquefasciatus Aphid E4 Myzw persicae Adapted from Karunaratne
I Toxicology Letters 82183 (1995) 83-W
or metabolism?
(% total protein)
7.7
Paraoxon Sequestered (ng)
Hydrolysis/h (%)
2.1
5.6
2.5
33.2
(0.4)
et al. [12].
expression of non-amplified structural gene sequences. Conversely, gene expression can be decreased. Various types of mutations can lead to changes in gene expression and these can occur in cis (for instance disruption or deletion of an upstream regulatory element of the gene, whether this element is enhancing or repressing gene expression) or in trans (for instance disruption of a gene coding for a protein that binds to the above-mentioned cis elements). It is clear that evidence for regulatory changes in gene expression are more difficult to obtain and to describe accurately in molecular terms. Regulatory changes appear to be involved in cases of metabolic resistance, specifically in those instances where glutathione S-transferases (including DDT-dehydrochlorinase) and cytochrome P450 monooxygenases play a role in insecticide metabolism. Since both groups of enzymes are multigene families comprising many genes that have evolved by repeated duplication/ divergence events, there may be significant redundancy in the catalytic function of the gene products. This in turn may allow mutations affecting gene expression of one or more of these genes to arise and to be selected by insecticide pressure. Overexpression of glutathione S-transferases (GST) has been documented in resistant strains of several insect species. In the yellow fever mosquito, Aedes aegypti, overexpression of GST2 is controlled by a trans-acting regulatory locus
PAThe P450 gene CYP6Al cloned from a resistant strain of the house fly is overexpressed in a number of strains. Gene amplification as a pos-
sible mechanism for high constitutive expression in the resistant strain was ruled out by dilution dot blot analysis of equal amounts of genomic DNA from both strains. Genetic crossing experiments mapped high constitutive expression of CYPdAl and insecticide resistance to chromosome II. The structural gene for CYPdAl was mapped to chromosome V, however. The results suggested that the gene product of the resistance gene on chromosome II is a diffusible factor which differentially regulates transcription of structural genes in susceptible and resistant strains [14]. The exact nature of the difference, i.e. the resistance mutation has not yet been determined. The regulatory factor has a pleiotropic activity and must regulate P450 genes other than CYP6Al. This conclusion is based on the observation that metabolism of diazinon, increased in resistant strains, is not catalyzed by CYP6Al. Instead, CYP6Al expressed in E. coli, purified and reconstituted with house fly NADPH-cytochrome P450 reductase, catalyzes the epoxidation of aldrin, heptachlor and of some terpenoids [14]. Whether changes in gene expression are involved in other cases of metabolic resistance remains to be established. Preliminary evidence for 2 other P450 genes, the CYP6Dl gene in permethrin-resistant house flies and the Cypdu2 gene in DDT-resistant Drosophila also implicates up-regulation in trans. A regulatory gene mutation causing metabolic resistance to insecticides has been predicted on genetic and biochemical grounds by Plapp [15] and the characterization of this regulatory gene should prove to be rewarding.
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-90
6. Linkage of DDT and pyrethroid insecticide resistance (knock-down resistance) to a Na+ channel locus
Knock-down resistance (kdr) has long been associated with the pharmacology of neuronal voltage-gated Na+-channels. Mutations in one Drosophila Na’-channel gene, para, were shown to have a weak resistant phenotype [16]. Recently, molecular probes for genes homologous to para have been obtained in the tobacco budworm, Heliothis virescens [17], in the house fly, and in the German cockroach, Blatella germanica. These probes were used to demonstrate genetic linkage between a polymorphism in the paru homolog of these species and permethrin (Heliothis) or DDT resistance. Linkage was very tight in the house fly and the cockroach, and less so in t.he tobacco budworm [17], possibly because metabolic resistance factors in addition to kdr may have been present in those insects. Thus, kdr mutants may be structural mutants of Na+ channel proteins. However, other mutations in Drosophila also affect para indirectly [16], and cause some degree of resistance to pyrethroids. For instance, a reduced number of nerve membrane Na’ channels may be involved in DDT and pyrethroid insecticide resistance as well. 7. Conclusion Great progress has been made in the last 5 years towards the elucidation of molecular mechanisms of insecticide resistance. But we have only a glimpse of the variety of mechanisms by which resistance is achieved. Work on the Met (methoprene tolerance) mutation in Drosophila which confers lOO-fold resistance to methoprene, a juvenile hormone analog, should soon yield exciting new insights into the mode of action of juvenile hormone. Resistance to the insecticidal toxins of Bacillus thuringiensis will need to be understood as transgenic crop plants expressing these toxins reach the farm. The demonstration of P450 gene amplification in humans (CYpZD6 in 2 Swedish families) led Meyer [18] to pose the question whether drugmetabolizing enzymes are preferred targets for
89
amplification in situations of xenobiotic or toxic exposure. The examples of insecticide resistance described above show that in such extreme situations eukaryotes can respond in very diverse ways, depending on the target and the chemical. This exploitation of genetic variation by insect pests will continue. We are challenged to integrate information on molecular mechanisms of resistance with population genetics with the aim of devising resistance management strategies. Acknowledgements
The brevity of this review has forced me to highlight and cite only some of the very nice research conducted on this subject. I acknowledge the fine contributions of all authors working on the molecular biology of insecticide resistance. References PI Georghiou,
G.P. (1990) Overview of insecticide resistance. ACS Symp. Ser. 421, 18-41. [21 Roush, R.T. and Tabashnik, B. (1990) Pesticide Resistance in Arthropods. Chapman & Hall. New York. 303 PP.
[31 ffrench-Constant,
R.H. (1992) Drosophila as a tool for investigating the molecular genetics of insecticide resistance. In: J. Oakeshott and M.J. Whitten (Eds.), Molecular Approaches to Fundamental and Applied Entomology, Springer, New York, pp. l-37. R.H. (1994) The molecular and popu141 ffrench-Constant, lation genetics of cyclodiene insecticide resistance. Insect Biochem. Mol. Biol. 24, 335-345. F.J. (1984) Biochemistry of insecticide 151 Oppenoorth, resistance. Pestic. Biochem. Physiol. 22, 187-193. 161 Mutero, A., Pralavorio, M., Bride, J.M. and Fournier, D. (1994) Resistance-associated point mutations in insecticide-insensitive acetylcholinesterase. Proc. Natl. Acad. Sci. USA 91, 5922-5926. at ACS special [71 Williamson, M. (1995) Data presented conference on pesticide resistance. PI Stark, G.R. and Wahl, G.M. (1984) Gene amplification. Ann. Rev. Biochem. 53. 447-491. A.L. and Field, L.M. (1991) Gene amplifi[91 Devonshire, cation and insecticide resistance. Ann. Rev. Entomol. 36. l-23. M., Callaghan, A., Fort, P. and Pasteur, N. t101 Raymond, (1992) Worldwide migration of amplified insecticide resistance genes in mosquitoes. Nature 350, 151-153. [Ill Poirie, M., Raymond, M. and Pasteur, N. (1992) Identification of two distinct amplifications of the esterase B
90
[12]
[13]
[14]
[15]
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-W
locus in Culex pipiens (L.) mosquitoes from Mediterranean countries. Biochem. Genet. 30, 13-26. Karunaratne, S.H.P.P., Jayawardena, K.G.I., Hemingway, J. and Ketterman, A.J. (1993) Characterization of a B-type esterase involved in insecticide resistance from the mosquito Culex quinquefasciatus. Biochem. J. 294, 575-579. Grant, D.F. and Hammock, B.D. (1992) Genetic and molecular evidence for a trans-acting regulatory locus controlling glutathione S-transferase-2 expression in Aedes aegypti. Mol. Gen. Genet. 234, 169-176. Feyereisen, R., Andersen, J.F., Carino, F.A., Cohen, M.B. and Koener, J.F. (1995) Cytochrome P450 in the house fly: structure, catalytic activity and regulation of expression in an insecticide-resistant strain. Pestic. Sci. 43, 233-239. 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. Pestic. Biothem. Physiol. 22,194-201. [16] Hall, L.M. and Kasbekar, D.P. (1989) Drosophila sodium channel mutations affect pyrethroid sensitivity. In: T. Narahashi and J.E. Chambers (Eds.), Insecticide Action. From Molecule to Organism, Plenum Press, New York, pp. 99-114. [17] Taylor, M.J.F., Heckel, D.G., Brown, T.M., Kreitman, M.E. and Black, B. (1993) Linkage of pyrethroid insecticide resistance to a sodium channel locus in the tobacco budworm. Insect Biochem. Mol. Biol. 23, 763775. [18] Meyer, U.A. (1994) Pharmacogenetics: the slow, the rapid and the ultrarapid. Proc. Natl. Acad. Sci. USA 91, 1983-1984.
Toxicology Letters 82183 (1995) 83-90
Molecular
biology of insecticide resistance R. Feyereisen
Department of Entomology,
Forbes 410, University of Arizona,
Tucson, AZ 85721, USA
Abstract The widespread use of insecticides has amounted to a large scale ‘experiment’ in natural selection of insects by chemicals of toxicological importance to humans. Specific examples in which the molecular basis of insecticide resistance has been studied in detail are presented here. The biochemical/physiological mechanisms of resistance can be categorized as target site insensitivity, increased metabolic detoxification and sequestration or lowered availability of the toxicant. These are achieved at the molecular level by: point mutations in the ion channel portion of a GABA receptor subunit (cyclodiene insecticides); point mutations in the vicinity of the acetylcholinesterase (AChE) active site (organophosphorus and carbamate insecticide resistance); amplification of esterase genes (organophosphorus and carbamate insecticides); mutations linked genetically to a sodium channel gene (DDT and pyrethroid insecticides); and yet uncharacterized mutations leading to the up-regulation of detoxification enzymes,
such as cytochrome P450 and glutathione S-transferases (many classes of insecticides). In several cases, the selection of a precisely homologous mutation has been observed in different insect species. Keyword: Pesticides; GABA gated Cl--channek Acetylcholinesterase; Na’ -channel; Cytochrome P450; Glutathione S-transferases; Esterases
1. Introduction
Perhaps the most serious consequence of widespread, indiscriminate use of insecticides is the development of insecticide resistance. More than 500 species of insects and mites are reported to have developed resistance to one or more classes of insecticides [l]. A number of agricultural pests and disease vectors are so resistant in some areas of the world that chemical control has become extremely difficult. The list of effective insecticides for the control of crop pests and disease vectors is rapidly shrinking. Meanwhile, fewer new insecticides are being introduced to the market, largely because of the high costs associated with research, development and registra0378-4274/95/$09.50 @ 1995 SSDI 0378-4274(95)03470-6
tion, and the prognosis of a limited effective lifespan of the new insecticide. The impact of insecticide resistance in toxicology is 2-fold: (1) the insecticide ‘treadmill’ has lead to the introduction of a huge arsenal of sometimes highly toxic chemicals in the environment, and several have become problem pollutants over the last 50 years; (2) the immediate reaction to insecticide resistance by pest control practitioners has traditionally been an increase in the dosage, thus compounding the problems of resistance and environmental contamination. The study of insecticide resistance at the molecular level has uncovered a number of ways in which animals can survive in toxic environments. The nature of the mutational events that
Elsevier Science Ireland Ltd. All rights reserved
84
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-M
lead to resistance in insects should not be expected to be intrinsically different from those that are responsible for the genetic variation of human responses to xenobiotics. Indeed, insects and vertebrates share many of the favorite targets for ‘insecticidal’ action and they share many of the metabolic detoxification systems. Selectivity of insecticides is achieved either when insects and vertebrates do not share a target (e.g. the selectivity of juvenile hormone agonists) or when they follow different metabolic pathways (e.g. the low vertebrate toxicity of malathion). Resistance marks a genetic change in response to selection. Individuals carrying genetic traits for coping with the chemically hostile environment survive and reproduce, thereby passing on these traits to their progeny. Continued selection pressure exerted by the insecticide rapidly increases the frequency of the genetic trait (resistance) in the population. I present here a brief overview of molecular mechanisms (types of mutations) that lead to insecticide resistance.
2. Physiological/biochemical mechanisms of resistance
and molecular
Genetic and biochemical studies have contributed immensely to our understanding of resistance. There are 2 ways in which organisms can become resistant to xenobiotics such as pesticides: either by modifying the effective dose of the pesticide available at the target site or by modifying the target site itself. Thus, classically recognized mechanisms of resistance such as behavioral resistance, reduced penetration or absorption, sequestration and detoxification all contribute to decrease the dose of the pesticide, whereas a decreased target site sensitivity or a modification of target site number will contribute to render a dose of pesticide ineffective. In-depth discussions of these mechanisms are given in Roush and Tabashnik [2]. Molecular mechanisms of resistance are classified according to the type of genetic change selected by the insecticide pressure: point mutations in structural genes, or modifications in the number or activity of genes.
3. Point mutations in structural genes: cyclodiene resistance and acetylcholinesterase insensitivity Nucleotide deletions, additions, or substitutions in the DNA may alter the amino acid sequence of a protein. Point mutations in the target sites of several herbicides and fungicides are known to confer pesticide resistance. Two such examples are also known in insects, and illustrate the power of Drosophila as a model system for insecticide resistance in general [3].
3.1. Point mutation in the Rdl gene of Drosophila [4] Elucidation of the molecular mechanisms of cyclodiene insecticide resistance in Drosophila followed a logical, ab initio approach that led to the cloning of the first invertebrate GABA receptor subunit. Field-collected populations of Drosophila were screened for dieldrin resistance. A homozygous strain with 4000 X resistance was isolated, and the resistance gene (Rdl) mapped genetically at 26 CM on the left arm of chromosome III. Deficiency mapping and irradiationinduced rearrangements allowed a more precise localization to the cytological region 66F. The resistance gene was then uncovered by P element-mediated transformation of flies with a cosmid clone obtained from a chromosomal walk in that region. The cosmid clone contained a susceptible copy of the Rdl gene, thus ‘rescuing’ susceptibility in resistant flies. Screening of a cDNA library with a fragment of this cosmid clone yielded a clone encoding a GABA receptor subunit. This confirmed earlier pharmacological evidence that had pointed to the picrotoxinin binding site on GABA, receptors as the likely neuronal target of cyclodiene insecticides. Expression of the cDNA in Xenopus oocytes demonstrated a functional, homo-oligomeric GABA-gated chloride channel. Sequencing of Rdl in resistant flies revealed a single common point mutation of Ala302 to Ser. This mutation is located in the second membrane spanning region of the channel subunit, lining the
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-90
chloride ion pore. Site-directed mutagenesis of the susceptible cDNA and expression in Xenopus oocytes showed that this mutation leads to channels insensitive to block by dieldrin. Armed with a molecular diagnostic, a number of dieldrin-resistant and susceptible strains were screened. The Ala”‘* to Ser mutation was absent in 122 susceptible strains, and present in 58 resistant strains of Drosophila melanogaster from around the world. The only observed fitness cost of this mutation is a temperature-sensitive phenotype, i.e. temporary paralysis at 38°C. Most remarkably perhaps, was the subsequent demonstration that an identical mutation is found in cyclodiene-resistant strains of a number of other insect species. In 2 species, the homologous Ala residue is mutated to Gly instead of Ser (Table 1). The molecular diagnostic for target site resistance to cyclodiene insecticides also allowed an accurate determination of resistance frequency in a relatively small samples of flies collected in the field. Resistance frequency was shown to be as high as 1%) even though wild populations of Drosophila in the US have apparently not been under cyclodiene selection pressure for many years [4]. Table 1 Conserved
M2 region
of the Rdl GABA
receptor
subunit
3.2. Point mutations in the Ace gene of Drosophila
Reduced sensitivity of acetylcholinesterase (AChE) is a well-characterized biochemical mechanism of insecticide resistance. Higher K,,, and lower k; values for organophosphorus and carbamate insecticides in resistant strains indicated structural alterations in the active site of the enzyme [5]. In turn, these alterations in protein structure may be attributed to changes in the amino acid sequence of the target protein. The structural gene, called Ace, coding for the AChE specifically expressed in the central nervous system of Drosophila melanogaster has been cloned, sequenced and extensively characterized. Because Ace is a single locus that was known a priori to be involved in resistance, a directed search for point mutations was justified. Comparisons of the coding sequences of the Ace gene of susceptible and resistant Drosophila strains revealed 5 point mutations at 4 positions in the resistant strains [6]. In one strain, mutations at all 4 positions are present in the same gene (Table 2). These point mutations were introduced in a minigene construct devoid of the carboxy terminus coding sequence. Thus expression of a soluble protein in Xenopus oocytes
in various
insect
species
Ala Arg Val Ala Leu Gly Val Thr Thr Residue
at the Ala’02 position
Drosophila melanogaster Drosophila simulans (2 alleles found) Aedes aegypti (mosquito) Periplaneta americana (cockroach) Musca domestica (house fly) Tribolium castaneum (flour beetle) Hypothenemus hampei (coffee berry borer) Bemisia tabaci (whitefly) Adapted
from
ffrench-Constant
[4].
x5
S-strain
R-strain
Ala (122 strains)
Ser (58 strains)
Ala
SeriGly
Ala
GUY
Ala
Ser
Ala
Ser
Ala
Ser
Ala
Ser
Ala
Ser
R. Feyereisen
86 Table 2 Point mutations
Drosophila S R Saltillo R Bygdea R Pierrefeu R MH19 Housefly R 77M R CH2 R 49R
in AChE
insensitive
I Toxicology Letters 82183 (1995) 83-90
to organophosphates/carbamates
Phe”‘
Ile”’
Ser
Val Val Thr
Gly _
“”
Ala Ala Ala _
Phe”’
Tyr _ _ Tyr
_ _ _ _
S Val”” Leu _ _
S, susceptible strain: R, resistant strain. Val”“, GlyZhZ, Phe3*’ and G~Y’~’ in the house fly correspond the Glu residue of the AChE catalytic triad. Adapted from Muter0 et al. [6]; Williamson [7].
to Val”“, Gly’“‘,
allowed a functional characterization of these mutations, either alone, or in some of the combinations found in resistant strains collected from the field. The results clearly indicated that multiple mutations caused higher levels of insensitivity to organophosphates and carbamates [6]. Multiple mutations in the same gene may result either from the accumulation of mutations, or from the recombination between genes carrying single mutations. This interesting problem has not been clearly resolved, but there are indications favoring the second possibility. Clearly, this would have important implications in resistance management, particularly with regard to migration, population size and fitness of the individual mutants in the absence of insecticide selection. Because the 3-dimensional structure of a model AChE, the electric organ enzyme from Torpedo californica, has been determined by Xray analysis, alignment of the Drosophila sequence has allowed a discussion of the possible influence of these mutations on substrate or inhibitor binding in the active site gorge of AChE. The enzyme has also been sequenced in other insect species, and recently the mutations conferring resistance in the house fly AChE have been determined [7]. Mutations at 4 sites were ob-
Gly”’ Ala Ala _
Phe”’ Tyr Tyr _
Gly”’ _ _ Ala
PhexhX and GIY~“~ m Drosophila. Gly”‘ is adjacent
to
served, 2 of which at residues homologous to those determined in Drosophila (Table 2). The evidence gathered to date on Rdl and Ace point mutations allows 2 interim conclusions. First, that Drosophila is a powerful and predictive model for major insect pest species. Second, that the ‘choice‘ of point mutations that lead to a modified sensitivity to xenobiotics upon selection is probably most limited in target molecules that are most conserved across taxonomical boundaries. This is also a conclusion that emerges from studies on herbicide resistance in photosystem II. 4. Gene amplification: to organophosphorus insecticides
esterases and resistance and carbamate
DNA amplification events create additional copies of chromosomal sequences (including functional genes) which can survive in either intra- or extrachromosomal forms. This phenomenon was first extensively documented in tumor cells and permanent cell lines, for instance methotrexate resistance resulting from the amplification of the target dihydrofolate reductase [8]. Initially regarded as abnormal events occurring in tissue cultures, DNA amplification is now amply documented in whole organisms where
R. Feyereisen
I Toxicology Letters 8.2183 (1995) 83-90
expression of the gene product must be faster than can be achieved by transcription from only a single copy of the gene [8]. At this point, it must be emphasized that DNA amplification per se does not bring about resistance. The amplified gene must be transcribed and translated (overexpressed) into functional protein products for expression of the resistance trait. Devonshire and Sawicki, working with organophosphorus-resistant populations of the aphid Myzus persicae, first invoked a simple model of ‘a succession of tandem duplications of the structural gene’ for an insecticide-associated esterase (esterase E4). In a series of progressively more resistant clones, progressive elevation of titers of E4 correlated with the degree of observed resistance. Esterase E4 (and the closely related FE4) confer broad resistance to insecticidal esters [9]. In vitro translation of poly(A)+RNA from 3 clones showed elevated levels of E4- or FECencoding mRNAs in the resistant clones. Direct evidence for an up to 64-fold amplification of the E4 esterase in M. persicae was obtained when a cDNA clone was used to probe the genomes of susceptible and various resistant aphid clones [9]. Reversion of resistance without loss of the amplified sequences involves in a still obscure way the DNA methylation status of the esterase genes. Evidence for DNA amplification of esterase genes was also obtained in a number of insecticide-resistant strains of the Culex pipienslquinquefasciatus complex. Resistant mosquitoes from around the world have been characterized over the last few years, revealing amplification of several closely related esterases (Table 3). Although migration was invoked as a major cause
Table 3 Amplified
esterases
in Culex pipienslquinquefasciatus
of the worldwide distribution of amplified esterase B2 [lo], it is equally likely that multiple amplification events have taken place. A molecular analysis of the amplification unit (or amplicon) of esterases in aphids and mosquitoes revealed that amplification of genomic sequences extends far beyond the esterase gene itself [9]. The Bl esterase of mosquitoes is a 2.8 kb gene found 250 times as an amplicon of approximately 30 kb, with a core of 25 kb. The E4 esterase of aphids is a 4.3 kb gene, amplified up to 64-fold as an amplicon of about 25 kb. This is consistent with observations on other xenobiotic-related amplification events in eukaryotes
PI. Because the amplified esterase can account for a very significant percentage of the total protein of the insect (up to 3% in most resistant aphids), is the amplified esterase merely a sponge that sequesters the insecticide? In other words, does resistance result from hydrolysis or sequestration of the insecticide? The kinetic constants (affinity, bimolecular rate of formation of the acylated enzyme, hydrolysis/ deacylation rate) for each esterase and each substrate need to be measured. A comparison of the aphid E4 and the mosquito A2/B2 esterases [12] indicated that hydrolysis was the major role of the aphid E4 esterase, whereas the mosquito A2/B2 esterases play a predominant sequestration role (Table 4). 5. Regulatory
changes in gene expression
DNA amplification is a mechanism whereby overabundance of a gene product is achieved by a multiplication of the gene itself, but there are molecular mechanisms responsible for increased
mosquitoes Chlorpyrifos
Origin
Esterase
Copies
Esterase
North America Africa. Asia, N. America France France Cyprus
Bl A2 B2 Al A4 B4 B5
250x 60x _
500X _
800x _
70x 50x 500x
100x 6.6X 95x
Adapted
from
[9-111.
25x 250x
87
levels
resistance
R. Feyereisen
88
Table 4 Esterase amplification in insects: sequestration pmol/insect
Mosquito A2/B2 Culex quinquefasciatus Aphid E4 Myzw persicae Adapted from Karunaratne
I Toxicology Letters 82183 (1995) 83-W
or metabolism?
(% total protein)
7.7
Paraoxon Sequestered (ng)
Hydrolysis/h (%)
2.1
5.6
2.5
33.2
(0.4)
et al. [12].
expression of non-amplified structural gene sequences. Conversely, gene expression can be decreased. Various types of mutations can lead to changes in gene expression and these can occur in cis (for instance disruption or deletion of an upstream regulatory element of the gene, whether this element is enhancing or repressing gene expression) or in trans (for instance disruption of a gene coding for a protein that binds to the above-mentioned cis elements). It is clear that evidence for regulatory changes in gene expression are more difficult to obtain and to describe accurately in molecular terms. Regulatory changes appear to be involved in cases of metabolic resistance, specifically in those instances where glutathione S-transferases (including DDT-dehydrochlorinase) and cytochrome P450 monooxygenases play a role in insecticide metabolism. Since both groups of enzymes are multigene families comprising many genes that have evolved by repeated duplication/ divergence events, there may be significant redundancy in the catalytic function of the gene products. This in turn may allow mutations affecting gene expression of one or more of these genes to arise and to be selected by insecticide pressure. Overexpression of glutathione S-transferases (GST) has been documented in resistant strains of several insect species. In the yellow fever mosquito, Aedes aegypti, overexpression of GST2 is controlled by a trans-acting regulatory locus
PAThe P450 gene CYP6Al cloned from a resistant strain of the house fly is overexpressed in a number of strains. Gene amplification as a pos-
sible mechanism for high constitutive expression in the resistant strain was ruled out by dilution dot blot analysis of equal amounts of genomic DNA from both strains. Genetic crossing experiments mapped high constitutive expression of CYPdAl and insecticide resistance to chromosome II. The structural gene for CYPdAl was mapped to chromosome V, however. The results suggested that the gene product of the resistance gene on chromosome II is a diffusible factor which differentially regulates transcription of structural genes in susceptible and resistant strains [14]. The exact nature of the difference, i.e. the resistance mutation has not yet been determined. The regulatory factor has a pleiotropic activity and must regulate P450 genes other than CYP6Al. This conclusion is based on the observation that metabolism of diazinon, increased in resistant strains, is not catalyzed by CYP6Al. Instead, CYP6Al expressed in E. coli, purified and reconstituted with house fly NADPH-cytochrome P450 reductase, catalyzes the epoxidation of aldrin, heptachlor and of some terpenoids [14]. Whether changes in gene expression are involved in other cases of metabolic resistance remains to be established. Preliminary evidence for 2 other P450 genes, the CYP6Dl gene in permethrin-resistant house flies and the Cypdu2 gene in DDT-resistant Drosophila also implicates up-regulation in trans. A regulatory gene mutation causing metabolic resistance to insecticides has been predicted on genetic and biochemical grounds by Plapp [15] and the characterization of this regulatory gene should prove to be rewarding.
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-90
6. Linkage of DDT and pyrethroid insecticide resistance (knock-down resistance) to a Na+ channel locus
Knock-down resistance (kdr) has long been associated with the pharmacology of neuronal voltage-gated Na+-channels. Mutations in one Drosophila Na’-channel gene, para, were shown to have a weak resistant phenotype [16]. Recently, molecular probes for genes homologous to para have been obtained in the tobacco budworm, Heliothis virescens [17], in the house fly, and in the German cockroach, Blatella germanica. These probes were used to demonstrate genetic linkage between a polymorphism in the paru homolog of these species and permethrin (Heliothis) or DDT resistance. Linkage was very tight in the house fly and the cockroach, and less so in t.he tobacco budworm [17], possibly because metabolic resistance factors in addition to kdr may have been present in those insects. Thus, kdr mutants may be structural mutants of Na+ channel proteins. However, other mutations in Drosophila also affect para indirectly [16], and cause some degree of resistance to pyrethroids. For instance, a reduced number of nerve membrane Na’ channels may be involved in DDT and pyrethroid insecticide resistance as well. 7. Conclusion Great progress has been made in the last 5 years towards the elucidation of molecular mechanisms of insecticide resistance. But we have only a glimpse of the variety of mechanisms by which resistance is achieved. Work on the Met (methoprene tolerance) mutation in Drosophila which confers lOO-fold resistance to methoprene, a juvenile hormone analog, should soon yield exciting new insights into the mode of action of juvenile hormone. Resistance to the insecticidal toxins of Bacillus thuringiensis will need to be understood as transgenic crop plants expressing these toxins reach the farm. The demonstration of P450 gene amplification in humans (CYpZD6 in 2 Swedish families) led Meyer [18] to pose the question whether drugmetabolizing enzymes are preferred targets for
89
amplification in situations of xenobiotic or toxic exposure. The examples of insecticide resistance described above show that in such extreme situations eukaryotes can respond in very diverse ways, depending on the target and the chemical. This exploitation of genetic variation by insect pests will continue. We are challenged to integrate information on molecular mechanisms of resistance with population genetics with the aim of devising resistance management strategies. Acknowledgements
The brevity of this review has forced me to highlight and cite only some of the very nice research conducted on this subject. I acknowledge the fine contributions of all authors working on the molecular biology of insecticide resistance. References PI Georghiou,
G.P. (1990) Overview of insecticide resistance. ACS Symp. Ser. 421, 18-41. [21 Roush, R.T. and Tabashnik, B. (1990) Pesticide Resistance in Arthropods. Chapman & Hall. New York. 303 PP.
[31 ffrench-Constant,
R.H. (1992) Drosophila as a tool for investigating the molecular genetics of insecticide resistance. In: J. Oakeshott and M.J. Whitten (Eds.), Molecular Approaches to Fundamental and Applied Entomology, Springer, New York, pp. l-37. R.H. (1994) The molecular and popu141 ffrench-Constant, lation genetics of cyclodiene insecticide resistance. Insect Biochem. Mol. Biol. 24, 335-345. F.J. (1984) Biochemistry of insecticide 151 Oppenoorth, resistance. Pestic. Biochem. Physiol. 22, 187-193. 161 Mutero, A., Pralavorio, M., Bride, J.M. and Fournier, D. (1994) Resistance-associated point mutations in insecticide-insensitive acetylcholinesterase. Proc. Natl. Acad. Sci. USA 91, 5922-5926. at ACS special [71 Williamson, M. (1995) Data presented conference on pesticide resistance. PI Stark, G.R. and Wahl, G.M. (1984) Gene amplification. Ann. Rev. Biochem. 53. 447-491. A.L. and Field, L.M. (1991) Gene amplifi[91 Devonshire, cation and insecticide resistance. Ann. Rev. Entomol. 36. l-23. M., Callaghan, A., Fort, P. and Pasteur, N. t101 Raymond, (1992) Worldwide migration of amplified insecticide resistance genes in mosquitoes. Nature 350, 151-153. [Ill Poirie, M., Raymond, M. and Pasteur, N. (1992) Identification of two distinct amplifications of the esterase B
90
[12]
[13]
[14]
[15]
R. Feyereisen
I Toxicology Letters 82183 (1995) 83-W
locus in Culex pipiens (L.) mosquitoes from Mediterranean countries. Biochem. Genet. 30, 13-26. Karunaratne, S.H.P.P., Jayawardena, K.G.I., Hemingway, J. and Ketterman, A.J. (1993) Characterization of a B-type esterase involved in insecticide resistance from the mosquito Culex quinquefasciatus. Biochem. J. 294, 575-579. Grant, D.F. and Hammock, B.D. (1992) Genetic and molecular evidence for a trans-acting regulatory locus controlling glutathione S-transferase-2 expression in Aedes aegypti. Mol. Gen. Genet. 234, 169-176. Feyereisen, R., Andersen, J.F., Carino, F.A., Cohen, M.B. and Koener, J.F. (1995) Cytochrome P450 in the house fly: structure, catalytic activity and regulation of expression in an insecticide-resistant strain. Pestic. Sci. 43, 233-239. 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. Pestic. Biothem. Physiol. 22,194-201. [16] Hall, L.M. and Kasbekar, D.P. (1989) Drosophila sodium channel mutations affect pyrethroid sensitivity. In: T. Narahashi and J.E. Chambers (Eds.), Insecticide Action. From Molecule to Organism, Plenum Press, New York, pp. 99-114. [17] Taylor, M.J.F., Heckel, D.G., Brown, T.M., Kreitman, M.E. and Black, B. (1993) Linkage of pyrethroid insecticide resistance to a sodium channel locus in the tobacco budworm. Insect Biochem. Mol. Biol. 23, 763775. [18] Meyer, U.A. (1994) Pharmacogenetics: the slow, the rapid and the ultrarapid. Proc. Natl. Acad. Sci. USA 91, 1983-1984.