126.96.36.199.1. Metabolic resistance
The principal metabolic pathways by which pyre-throids are degraded in insects were mostly evaluated prior to 1985 and are summarized in Roberts and Hutson (1999). Metabolism is conveniently divided into two phases: the initial biotransformation of a pesticide is referred to as Phase I metabolism, comprising mainly oxidative, reductive, and hydrolytic processes; Phase II metabolism is biotransformation, in which the pesticide or Phase I metabolite is conjugated with a naturally occurring compound such as a sugar, sugar acid, glutathione, or an amino acid. In general, the major Phase I degradative routes in insects and other animals (mostly mammals and birds) are similar. It is only in the details of their Phase II reactions whereby polar conjugates with sugars or amino acids are formed that there are major qualitative differences in the nature of the metabolites. As degradation studies on insects are not required for the registration of insecticides, such studies are usually only undertaken in order to understand specific questions concerning structure-activity relationships or to evaluate problems associated with resistance caused by enhanced metabolic breakdown. It is the latter reason that has seen the majority of insect metabolism studies performed.
Since 1985 there has been a vast increase in the knowledge of insect molecular genetics. The publication of the draft genome sequences for the fruit fly Drosophila melanogaster in 2000 and the mosquito Anopheles gambiae in 2002 has greatly increased the knowledge of the enzymes involved in metabolic degradation, particularly cytochromes P450. Consequently, recent research is beginning to make inroads into understanding which of the isozymes of cytochrome P450 are responsible for metabolizing different substructures of the pyrethroid molecule. The situation with respect to the nature of the carboxyesterases responsible for catalysing the hydrolysis of the pyrethroid ester bond is less clear and the carboxyesterases that catalyze the hydrolysis of pyrethroids in different species of insects may be different and nonhomologous.
Insects detoxify pyrethroids at varying rates and this degradative metabolism is important in understanding the detailed toxicology. Indeed, part of the relatively modest insecticidal activity of the natural pyrethrins is attributable to their rapid metabolic breakdown. Consequently, household sprays are usually formulated with synergists that inhibit the enzymes that catalyze this metabolic degradation and thereby enhance the insecticidal activity. Selection of insect strains possessing elevated levels of catabolizing enzymes is also an important mechanism in the development of decreased sensitivity (resistance) toward pyrethroids. There are two principal routes of detoxification of pyrethroids in insects: de-esterification catalysed by both esterases and cytochromes P450, and the hydroxylation of aromatic rings or methyl groups by cytochromes P450, and these two mechanisms will be considered separately. The points of the chemical structure where a generalized 3-phenoxybenzyl pyrethroid
molecule is detoxified are shown in Figure 5. The width of the arrow indicates the approximate extent of metabolic attack. It is in the detailed enzymology and molecular biology of the enzymes responsible for pyrethroid metabolism where there have been the most important advances.
188.8.131.52.1.2. Metabolic pathways The most complete analysis of the metabolic pathways of pyre-throid degradation in insects has been by Shono (Shono et al., 1978; reviewed by Soderlund et al., 1983), who studied the metabolism of permethrin in American cockroaches (Periplaneta americana), houseflies (Musca domestica), and caterpillars (Trichoplusia ni). Forty-two metabolites (including conjugates) were identified, and the most important Phase 1 metabolites are shown in Figure 6. Metabolism occurred by ester cleavage to 3-phenoxybenzyl
alcohol (3-PBA) (45) and 3-(2,2-dichlorovinyl)-2,2-dimethylcyclopropanecarboxylic acid (DCVA) (46). Hydroxylation also occurred at the 40- and 6-positions of the phenoxybenzyl moiety and the cis or trans methyl groups of the DCVA moiety to give (47), (48), and (49) respectively. Hydroxylation of the trans-methyl group rather than the cis- was preferred. Ester cleavage of these hydroxylated metabolites gave the structures (50), (51), and (52) respectively. 3-PBA (50) was further oxidized to 3-phenoxybenzoic acid (53), as were the hydroxylated analogs of 3-PBA to their analogous benzoic acids.
A similar pattern of Phase I metabolites was observed with both cis- and trans-permethrin, although the relative proportion of certain of the conjugated structures was influenced by the stereochemistry. All metabolites found in the cockroach were also found in houseflies but the series of metabolites arising from the 6-hydroxylation of the 3-PBA moiety was only found in flies. Metabolites consisting of the whole hydroxylated molecule (e.g., (47), (48), and (49)) were exclusively found as their glucosides, whereas the ester cleavage products were found both free and as their glucoside and amino acid conjugates. All three insects conjugated DCVA with one or more of the amino acids glycine, glutamic acid, glutamine, and serine, in addition to forming glucose esters. This study did not identify the 22-hydroxylated metabolite (54), a significant metabolite in mammalian studies. However, this metabolite and compounds derived from it were identified in a study of the metabolism of permethrin by the American bollworm Helicoverpa zea, the tobacco budworm Heliothis virescens (Lepidoptera, Noctuidae) (Bigley and Plapp, 1978) and the Colorado potato beetle, Leptinotarsa decemlineata (Soderlund et al., 1987), in which it was the principal metabolite. Other later studies have confirmed this overall pattern in different insects, although some studies have shown more differences in the pattern of metabolites between cis- and trans-permethrin isomers.
Holden (1979) showed that trans-permethrin was ester-cleaved at a higher rate than the cis-isomer in P. americana, an observation consistent with the findings that the trans-isomers of cyclopropane-containing pyrethroids are much better substrates for esterases than the cis-isomers (see below). This pattern of permethrin metabolism by insects can be taken as a template for the breakdown of other pyrethroids comprising esters of 2,2-dimethyl-3-(substituted)vinylcarboxylic acid with 3-PBA. Thus, the principal mechanisms of Phase I metabolism involve ester cleavage, both hydrolytic and oxidative, and aromatic substitution of one or
Figure 7 Metabolites from oxidative attack on the chry-santhemic acid moiety of allethrin (structures 55-57).
Figure 7 Metabolites from oxidative attack on the chry-santhemic acid moiety of allethrin (structures 55-57).
other of the 3-PBA rings (the 4-position is usually the major site) and the 2,2-dimethyl group of the acid moiety. Analogous reactions have been shown to take place with bifenthrin and deltamethrin in the bulb mite Rhizoglyphus robini to give the ester-cleaved products and the 40-hydroxy metabolites (Ruzo et al., 1988). Similarly, trans-cypermethrin gave trans-DCVA and both 22- and 40-hydroxy-cypermethrin in the cotton bollworm Helicoverpa armigera and in H. virescens (Lepidoptera) (Lee et al., 1989). Fenvalerate, which lacks a cyclopropyl group, was metabolized via ester cleavage and 4-hydroxylation in houseflies (Funaki et al., 1994). In pyrethroids containing the chrysanthemic acid moiety (rather than a 2,2-dimethyl-3-dihalovinylcarboxylic acid), such as allethrin, phenothrin, and tetramethrin, the methyl groups of the isobutylene group are also subject to oxidative attack. Hydroxylation of these groups in allethrin to the alcohol (55) (Figure 7), followed by successive oxidation to the aldehyde (56) and carboxylic acid (57), occurred in houseflies (Yamamoto et al., 1969). Tetramethrin was mainly metabolized via ester cleavage, with the production of the carboxyl derivative (57)of the trans-methyl group as a minor product (Yamamoto and Casida, 1968).
184.108.40.206.1.3. Ester hydrolysis All pyrethroids with the exception of the nonester compounds, such as MTI 800 which has a hydrocarbon linkage between the ''acid'' and ''alcohol'' moieties, are subject to esterase-catalyzed breakdown. Even etofenprox, which has an ether linkage in this position, is cleaved via oxidation of the benzylic carbon and hydrolysis of the resultant ester in the rat (Roberts and Hutson, 1999). The stereochemistry of the cyclopropanecar-boxylic acid is important in determining the rate of esteratic detoxification, since the trans-isomers are very much better substrates for esterases than are the cis-isomers (Soderlund and Casida, 1977). Indeed, cis-pyrethroids possessing an a-cyano group (Type II
pyrethroids) and consequently a secondary ester are generally degraded via a microsomal P450 mechanism through oxidation of the a-carbon. Evidence from work with mammalian liver indicates that the esterases responsible for pyrethroid hydrolysis are also microsomal (Soderlund and Casida, 1977). This study also showed that the Type II pyrethroids cyper-methrin and deltamethrin were the least susceptible to both esterase-catalysed hydrolysis and oxidation. Generally, the major route of Type II pyrethroid metabolism in mammals is via de-esterification catalysed by microsomal oxidases (e.g., Crawford and Hutson, 1977). Studies on insects have also concluded that ester cleavage is often mainly by an oxidative mechanism (Casida and Ruzo, 1980; Funaki et al., 1994). The latter study concluded that the de-esterifi-cation of fenvalerate by pyrethroid-resistant house-flies was principally due to over-expression of cytochromes P450; only a small portion of the ester bond cleavage was caused by hydrolases.
Whether the ester cleavage is hydrolytic or oxi-dative is apparently largely dependent on the species. For example, T. ni (Ishaaya and Casida, 1980), S. littoralis, (Ishaaya et al., 1983), S. eridana (Abdelaal and Soderlund, 1980) (Lepidoptera), and B. tabaci (Jao and Casida, 1974; Ishaaya et al., 1987) (Hemiptera) all degraded trans-pyrethroids via a hydrolytic mechanism. Conversely, houseflies (Diptera) and the rust-red flour beetle Tribolium castaneum (Coleoptera) used an oxidative pathway (Ishaaya et al., 1987). A rather special case is that of the green lacewing Chrysoperla carnea agg. (Neuroptera), which is highly resistant to pyre-throids. This insect has been shown to contain high levels of a pyrethroid-hydrolyzing esterase that is able to catalyze the hydrolysis of cis-isomers, including those of Type II ester pyrethroids such as cyper-methrin (Ishaaya and Casida, 1981; Ishaaya, 1993). How the over-production of esterases induces resistance to cis-pyrethroids such as deltamethrin, which are poor substrates for the enzyme(s) from other species, has been the cause of some conjecture. Devonshire and Moores (1989) presented evidence that cis-pyrethroids bind tightly to the active site and are thus sequestered by the large amounts of the resistance-associated esterases (E4 and FE4) produced by resistant peach-potato aphids M. persicae. Consequently, deltamethrin acts as a competitive inhibitor of esterase activity but is removed by binding to the protein rather than by hydrolysis. Selection pressure, whether artificially or from frequent commercial use of pyrethroids, frequently leads to multifactorial mechanisms of insecticide resistance, including esterases, cytochromes P450, target-site (kdr and super-kdr) mechanisms, and sometimes reduced penetration (e.g., Anspaugh et al., 1994; Pap and Toth, 1995; Ottea et al., 2000; Liu and Pridgeon, 2002). Such insect strains have extremely high resistance and usually show some degree of cross-resistance to all pyrethroids.
Most insect esterases that catalyze the hydrolysis of pyrethroid esters are soluble nonspecific B-type carboxylesterases. These have a wide substrate-specificity and model substrates, such as 1-naphthyl acetate, have been used to visualize electrophoreti-cally distinct protein bands in B. tabaci (Byrne et al., 2000) and many other insect species. There is little information to date on the occurrence of microsomal esterases in insects analogous to those in mammals (e.g., Prabhakaran and Kamble, 1996). Most evidence for the involvement of esterases in pyrethroid breakdown is indirect and has used model substrates; however, direct evidence for pyrethroid hydrolysis has been obtained for esterases from several species, including the horn fly Haematobia irritans (Diptera) (Pruett et al., 2001). Concern has been raised of the validity of using model substrates to predict the breakdown of pyrethroids, and consequently some researchers have designed model substrates with pyrethroid-like structures. Thus, Shan and Hammock (2001) developed sensitive fluorogenic substrates based on DCVA coupled to the cyanohydrin of 6-methoxynaphthalene-2-carboxaldehyde. Hydrolysis of this ester and decomposition of the cyanohydrin regenerates this fluorescent aldehyde. An enzyme preparation that used this substrate from a cyper-methrin-selected resistant strain of H. virescens (Lepidoptera) gave better selectivity (x5) than did 1-naphthyl acetate (x1.4). A similar approach, but using 1-naphthyl esters of the four stereoisomers of DCVA with detection of released 1-naphthol by diazo-coupling to Fast Blue RR, has been reported (Moores et al., 2002). In these experiments, only the (1S)-trans-isomer acted as a substrate for aphid esterases, in agreement with other experiments on the stereospecificity of insect pyrethroid esterases. Activity staining of electrophoretic gels using this novel substrate showed that the pyrethroid-hydrolyzing activity was distinct from the resistance-associated esterases visualized with 1-naphthyl acetate, indicating that the main mechanism for resistance was binding/sequestration rather than hydrolysis.
Work since 1985 has concentrated on understanding the nature of the esterases responsible for pyrethroid hydrolysis. In vivo studies have often yielded equivocal results and, using such tools as specific inhibitors of oxidative or hydrolytic metabolism, it has frequently been difficult to prove which mechanism is primarily responsible for pyrethroid catabolism. These problems have arisen in part because there is no such thing as a specific inhibitor and some compounds thought to be specific inhibitors of cytochromes P450 (e.g., piperonyl butox-ide) may also inhibit esterases (Gunning et al., 1998b; Moores et al., 2002).
Studies have included both in vivo work on whole insects or their tissues and in vitro studies with isolated enzymes. In resistant populations of many insect species, the mechanisms are most frequently due to both enhanced esterase and cyto-chrome P450 levels, so that dissection of the proportions of the different mechanisms is difficult. However, evidence that resistance ratios are reduced or abolished in the presence of reliably specific inhibitors such as organophosphates (Gunning et al., 1999; Corbel et al., 2003) can be taken as a good indication that enhanced esterase levels are responsible for metabolic resistance.
These are a class of Phase I detoxification enzymes that catalyse various NADPH- and ATP-dependent oxidations, dealkylations, and dehydrogenations. Both microsomal and mitochondrial forms occur in insects. They are probably responsible for the most frequent type of metabolism-based insecticide resistance (Oppenoorth, 1985; Mullin and Scott, 1992; Scott and Wen, 2001). They are also a major mechanism for pyrethroid catabolism (Tomita and Scott, 1995). Their occurrence and importance in insect xenobiotic metabolism has been reviewed by Scott and Wen (2001). The super-family of cytochrome P450 genes has probably evolved by gene duplication and adaptive diversification, and comprises 86 functional genes in D. melanogaster. The large number of substrates metabolized by P450s is due both to the multiple isoforms and to the fact that each P450 may have several substrates (Rendic and DiCarlo, 1997). Because these enzymes may have overlapping substrate specificities, it is difficult to ascribe the function to individual P450 enzymes. In insects, although the importance of oxygenases in the metabolism of many substrates is known, the particular P450 isoforms involved have rarely been identified. For a general review on insect P450 enzymes and particularly their regulation, see Chapter 4.1.
Several P450 iso-enzymes have been isolated or expressed from insect sources. Regarding pyrethroid metabolism, the best-characterized P450 isoform is CYP6D1. This was originally purified from a strain of highly resistant (ca. x5000) houseflies designated "Learn pyrethroid resistant'' (LPR) selected by the continuous usage of permethrin to control flies in a New York State dairy. A reduced-penetration mechanism and kdr were also present in the strain. CYP6D1 has been purified (Wheelock and Scott, 1989) and sequenced via the use of degenerate primers derived from known protein sequences and PCR amplification (Tomita and Scott, 1995). Overproduction of this P450 isozyme was found to be the major mechanism of deltamethrin detoxification in microsomes derived from the LPR flies (Wheelock and Scott, 1992). The enzyme requires cytochrome b5 as a co-factor and is specific in its action, because only the 4'-hydroxy metabolite was produced from cypermethrin (Zhang and Scott, 1996). CYP6D1 was found to be the major and possibly the only P450 isoform responsible for pyrethroid metabolism in this strain of houseflies; consequently, the resistance ratios are very much less for pyrethroids such as fenfluthrin that do not have the 3-phenox-ybenzyl group (Scott and Georghiou, 1986). The same mechanism was found to be responsible for PBO suppressible resistance to permethrin from a Georgia poultry farm in the USA (Kasai and Scott, 2000). In both these housefly strains, the mechanism was due to an increased (ca. x10) transcription of the gene, leading to increased levels of CYP6D1 mRNA and higher levels of the enzyme. CYP6D1 is expressed in the insect nervous system and has been shown to protect the tissue from the effects of cypermethrin (Korytko and Scott, 1998). Clearly, from the metabolic specificity of CYP6D1, other isoforms of cytochromes P450 must also be implicated in pyrethroid metabolism, although which reactions are catalyzed by which isoform has yet to be determined.
It is characteristic of monooxygenases that they are inducible within an individual animal. The use of phenobarbitone to induce monooxygenase activity in rat liver is well known, and many other agents are capable of transiently up-regulating cyto-chromes P450. Phytophagous insects are exposed to many plant xenobiotics, for example monoter-penes which also induce P450 production. Such induction of P450s may incidentally induce an isoform also capable of metabolizing pyrethroids. For example, feeding larvae of H. armigera on mint (Mentha piperita) leaves induced a 4x resistance to pyrethroids compared with those fed on a semi-defined diet (Hoque, 1984; Terriere, 1984; Schuler, 1996; Scott et al., 1998). CYP6B2 mRNA, a P450 isoform also implicated in pyrethroid resistance, is inducible by peppermint oil and specifically a-pinene in larvae of H. armigera (Ranasinghe et al., 1997). This induction was rapid (ca. 4h) and disappeared within a similar period of removing the stimulus. Clearly, the mechanism for the transient induction of P450s (Ramana, 1998) is different from the situation with the LPR houseflies, in which CYPD1 is permanently up-regulated (Liu and Scott, 1998), although the precise mechanistic details still remain to be elucidated. In nonphytophagous arthropods for example the cattle tick Boophilus microplus which is not subjected to a barrage of allelochemicals in its diet, P450s were found to be of little importance in the induction of resistance to pyrethroids (Crampton et al., 1999), although the converse has been found for adults of the mosquito Culex quinquefasciatus (Kasai et al., 1998).
220.127.116.11.1.5. Model substrates When managing insecticide resistance, it is important that resistant alleles can be detected at low frequency in populations. Data from bioassays will only detect a quite high proportion of individuals with reduced sensitivity in the population. Consequently, it is useful to design biochemical or DNA (molecular) tests that can identify resistance in individual insects. Thus, model substrates commonly used to measure P450 levels in vitro need to be substrates of the relevant P450 isoforms that degrade the insecticide. Amongst such model substrates are the sensitive fluorogenic reagents 7-ethoxycoumarin, ethoxyresorufin, and methoxyresorufin, and the chromogenic substrate 4-nitroanisole. These compounds are dealkylated by monooxygenases to yield a fluorescent or colored product. Unfortunately, it has generally been found that these substrates are also isozyme specific, so that they may not be good indicators of P450-induced pyrethroid resistance. In the case of CYP6D1, methoxyresorufin was found to be a substrate, but ethoxyresorufin and 7-ethoxycoumarin were not (Wheelock and Scott, 1992). A similar variation in the activity of elevated oxygenases to 4-nitroanisole, benzo(a)pyrene, benzphetamine, and methoxyresorufin in the mid-gut of a multi-resistant (cypermethrin and thiodicarb) strain of H. virescens larvae was noted (Rose et al., 1995). In this strain, demethylation rates of both 4-nitroanisole and methoxyresorufin were useful as indicators of insecticide resistance; however, on such multi-resistant strains, several P450 isoforms are probably elevated in tandem making comparisons difficult. Consequently, the use of model substrates to estimate the levels of P450 monooxygenases in individual insects may only give equivocal information on levels of P450-derived metabolic resistance to pyrethroids.
18.104.22.168.1.6. Glutathione-S-transferases (GSTs) GSTs are important in the detoxification of organo-phosphorus insecticides and other electrophilic compounds, which are dealkylated and conjugated with glutathione (see Chapter 5.11). Pyrethroids are not electrophilic and would not be expected to be detoxified by this mechanism. However, there are several reports that have correlated enhanced levels of GSTs with pyrethroid resistance in a number of species, S. littoralis (Lagadic et al., 1993), T. casta-neum (Reidy et al., 1990), and Aedes aegypti (Grant and Matsumura, 1989). Additionally, pyrethroids have been shown to induce production of GSTs in the honeybee Apis mellifera (Yu et al., 1984), fall armyworm Spodoptera frugiperda (Punzo, 1993), and German cockroach Blatella germanica (Hemingway et al., 1993). The role of the glutathi-one-S-transferase system as a mechanism of defence against pyrethroids is not fully understood, but it is thought that GST proteins sequester the pyrethroids (Kostaropoulos et al., 2001) or possibly protect the tissues from pyrethroid-induced lipid peroxidation (Vontas et al., 2001).
22.214.171.124.2. Cuticle penetration As a resistance mechanism, reduced penetration of the insect cuticle has been studied less and has generally been considered of subordinate importance to enhanced detoxification and target-site mutations. Where detected, it is usually found with other mechanisms, e.g., the ''LPR strain'' of houseflies referred to above. Reduced cuticular penetration of pyrethroids has been detected in resistant strains of a number of other species, including H. armigera (Gunning et al., 1991), H. zea (AbdElghafar and Knowles, 1996), H. virescens (Little et al., 1989), S. exigua (Delorme etal., 1988), the diamond-back moth Plutella xylostella (Noppun et al., 1989), B. germanica (Wu et al., 1998), and B. microplus (Schnitzerling et al., 1983). In all these cases, reduced cuticular penetration was found in addition to other resistance mechanisms, enhanced metabolism, and/or target-site resistance (kdr). Mechanisms involving reduced uptake appear not to give significant resistance to the lethal effects of insecticides but provide a more than additive effect when combined with other mechanisms. Thus, decreased penetration, although a minor factor on its own, can, when coupled with other mechanisms, increase resistance many-fold (Ahmad and McCaffery, 1999).
126.96.36.199.3. Target-site resistance Target-site resistance is the most important mechanism of pyre-throid resistance and its selection and spread have compromised the use of pyrethroid insecticides on many insect pests. It is a particularly important mechanism of pyrethroid resistance, as it confers a degree of loss of sensitivity to all members of the class. It is characterized by a reduction in the sensitivity of the insect nervous system and has been termed ''knockdown resistance'' or kdr (Sawicki, 1985). The kdr trait was first identified in the early 1950s in houseflies (Busvine, 1951). It causes a loss of sensitivity to DDT and its analogs, and to pyrethrins and pyrethroids, which all owe their activity to interaction with the para-type voltage-gated sodium channel in nerve membranes. This loss of activity is characterized by a reduction in the binding of these insecticides to the sodium channel (Pauron et al., 1989). An enhanced form of this resistance termed super-kdr has also been characterized in houseflies (Sawicki, 1978). Both the kdr and super-kdr traits were mapped to chromosome 3 and found to occupy the same allele, the para-type sodium channel. This has been confirmed by molecular cloning studies of these channels in kdr and super-kdr houseflies (Williamson et al., 1996) and kdr B. germanica (Miyazaki et al., 1996; Dong, 1997). The super-kdr resistance in houseflies is due to a methionine to threonine (M918T) point mutation in the gene encoding the para sodium channel. Both mutations were located with domain II of the ion channel. The L1014F mutation in IIS6 was found in both housefly and cockroach strains, and confers kdr resistance. To date, the M918T mutation has not been detected as a single substitution in any housefly strain, but only occurs in conjunction with L1014F. Mammals are intrinsically much less susceptible to pyrethroids and DDT. Significantly, mammalian neuronal sodium channels have an iso-leucine rather than methionine in the position (874) that corresponds to the housefly super-kdr site (918). Site-directed mutation of this residue to methionine gives rise to a channel with >100x increased sensitivity to deltamethrin, suggesting that differential pyrethroid sensitivity between mammals and insects may be due in part to structural differences between the mammalian and insect sodium channels (Vais et al., 2000). In some insect species, the existence of kdr-type target-site resistance has often been masked by efficient metabolic resistance mechanisms; an example is M. persicae, in which resistance-associated esterase is responsible for much of the reduced sensitivity towards organophosphates, carbamates, and pyrethroids. Any inference that target-site resistance might also be a factor has usually been based on the sensitivity of the insects in the presence of synergists such as PBO or DEF S,S,S,-tributyl phosphorotrithiolate, and the assumption that any residual decreased sensitivity is due to kdr-type target-site resistance.
Use of degenerate DNA primers for the para-type sodium channel and sequencing of the gene have resulted in the unequivocal identification of resistance-inducing mutations in the trans-membrane domain II (Martinez-Torres ) of the channel in M. persicae, in which a leucine to phenyl-alanine (L1014F) mutation associated with kdr was identified. Insects containing this mutation could also be identified by the use of a discriminating dose of DDT, as DDT is unaffected by the enhanced-esterase mechanisms also present in most of the aphid clones. Indeed, of 58 aphid clones analysed for both kdr- and esterase-based mechanisms, only four contained an esterase (E4) mutation and not kdr. Estimates for resistance factors in aphids containing both mechanisms are 150-540-fold, in comparison to 3-4-fold (FE4 esterase alone) or 35-fold for kdr alone. Consequently, these dualresistance mechanisms afford a level of decreased sensitivity whereby insects become totally immune to field dosages of pyrethroids. Similar kdr mutations have also been detected in cockroaches (Miyazaki et al., 1996; Dong, 1997), H. irritans (Guerrero et al., 1997), P. xylostella (Schuler et al., 1998), and An. gambiae (Martinez-Torres et al., 1998). In a pyrethroid-resistant strain of the tobacco budworm H. virescens, the same locus was mutated to histidine rather than phenylalanine (Park and Taylor, 1997)
As with metabolic resistance mechanisms, it is important to establish methods that can identify sodium-channel kdr-type mechanisms in single insects so that it is possible to adjust insect-control methods. The kdr-mutation of nerve insensitivity was originally identified by electrophysiology, and this method still remains a fundamental way of confirming nerve insensitivity. However, it is a specialized and rather cumbersome technique that is out of the question when attempting to test large numbers of an agricultural pest species. The DDT bioassay using a discriminating dose remains a useful method but may not completely discriminate between homozygous and heterozygous individuals. The most useful technique has been direct diagnosis of the mutation(s) based on PCR amplification and sequencing. The identification of the L1014F mutation in knockdown-resistant housefly strains has led to the development of several diagnostic assays for its occurrence in other species, including H. irritans (Guerrero et al., 1997), the mosquitoes An. gambiae (Martinez-Torres et al., 1998), and Culex pipiens (Martinez-Torres et al., 1999a), as well as M. persi-cae. However, this technique will only identify known mutations and the test designed to detect L1014F in An. gambiae (Ranson et al., 2000) did not detect the additional L1014S.
The molecular biology of knockdown resistance to pyrethroids has been reviewed (Soderlund and Knipple, 2003). Sequencing of the para-sodium channel gene from several arthropod species has led to the discovery of a number of amino-acid polymorphisms in the protein. Of the 20 amino-acid polymorphisms each uniquely associated with pyre-throid resistance, those occurring at four sites have so far been found as single mutations in resistant populations. These are valine 410 (V410M in H. vir-escens; Park et al., 1997), methionine 918 (M918V in B. tabaci; Morin et al., 2002), leucine 1014 (L1014F in several species; L1014H in H. virescens; Park and Taylor, 1997; L1014S in C. pipiens; Martinez-Torres et al., 1999a), An. gambiae (Ranson et al., 2000), and phenylalanine 1538 (F538I, in B. microplus; He et al., 1999). All these sites have been located on trans-membrane regions of the channel and it is inferred that they are close to the binding site(s) of pyrethroids and DDT. Additionally, other polymorphisms have been found that only occur in the presence of L1014F and appear to act in a similar way to M918T in houseflies, causing enhanced resistance.
Was this article helpful?