Insecticide resistance in insect vectors of human disease

Annu. Rev. Entomol. 2000. 45:371–391

INSECTICIDE RESISTANCE IN INSECT VECTORS

HUMAN DISEASE

Janet Hemingway and Hilary Ranson

School of Biosciences, University of Wales Cardiff, P.O. Box 915, Cardiff, Wales CF1

3TL; e-mail: sabjh@cardiff.ac.uk

Key Words insecticide, mosquito, esterases, monooxygenases, glutathione S-

transferases

Abstract Insecticide resistance is an increasing problem in many insect vectors

of disease. Our knowledge of the basic mechanisms underlying resistance to com-

monly used insecticides is well established. Molecular techniques have recently

allowed us to start and dissect most of these mechanisms at the DNA level. The next

major challenge will be to use this molecular understanding of resistance to develop

novel strategies with which we can truly manage resistance. State-of-the-art infor-

mation on resistance in insect vectors of disease is reviewed in this context.

INTRODUCTION

Insecticides play a central role in controlling major vectors of diseases such as

mosquitoes, sandﬂies, ﬂeas, lice, tsetse ﬂies, and triatomid bugs. In 1955 the

World Health Organization (WHO) assembly proposed the global eradication of

the most prevalent vector-borne human disease, malaria, by the use of residual

house-spraying of DDT. However, the insecticide euphoria soon ended and in

1976 WHO ofﬁcially reverted from malaria eradication to malaria control. This

marked shift from malaria eradication to primary health care was an emotive

issue, eliciting a rapid and complete change of rhetoric from WHO (12). Several

issues had prompted this switch, but a major cause of the change in policy was

the appearance of DDT resistance in a broad range of the mosquito vectors. In

1975 WHO reported that 256 million people were living in areas where DDT

and/or BHC resistance was undermining malaria control efforts. (This did not

include the African region, where 90% of malaria occurs and where DDT resis-

tance had already been noted in Anopheles gambiae, the major malaria vector.)

The resistance problems continued with the switch to newer insecticides such

as the organophosphates, carbamates and pyrethroids. Operationally, many con-

trol programs have switched from blanket spraying of house interiors to focal use

of insecticides on bednets. Focal spraying limits the insecticides of choice largely

to pyrethroids due to the speed of kill required to protect the occupant of the

0066-4170/00/0107-0371/$14.00 371

372 HEMINGWAY n RANSON

bednet and the safety margin needed for insecticides used in such close contact

with people. Today the major emphasis in resistance research is on the molecular

mechanisms of resistance and rational resistance management, with a view to

controlling the development and spread of resistant vector populations. In Africa,

WHO and the World Bank have instigated major new initiatives with other major

donors and the scientiﬁc community internationally to ‘‘roll back’’ malaria. One

major problem these initiatives are tackling is the presence of two developing

foci of pyrethroid resistance in the most important African malaria vector, An.

gambiae.

SCALE OF THE PROBLEM

The amount of resistance in insect vector populations is dependent both on the

volume and frequency of applications of insecticides used against them and the

inherent characteristics of the insect species involved. Tsetse ﬂies, for example,

were controlled by wide-scale spraying of DDT for many years, but DDT resis-

tance has never developed in this species. Another example of an insect vector

exhibiting little or no resistance to insecticides is the triatomid bug. In both cases

the major factor inﬂuencing insecticide resistance development is the life cycle

of the insect pest, in particular the long life cycles for the bugs, and the production

of very small numbers of young by the tsetses. In contrast, mosquitoes have all

the characteristics suited to rapid resistance development, including short life

cycles with abundant progeny.

Mosquito Resistance

The major mosquito vectors span the Culex, Aedes, and Anopheles genera. Culex

are the major vectors of ﬁlariasis and Japanese encephalitis, Aedes of dengue and

dengue hemorrhagic fever, and Anopheles of malaria. The range of many of these

species is not static. For example, several Aedes species recently extended their

range in Asia and Latin America, leading to an increased risk of dengue in these

areas.

DDT was ﬁrst introduced for mosquito control in 1946. In 1947 the ﬁrst cases

of DDT resistance occurred in Aedes tritaeniorhynchus and Ae. solicitans (15).

Since then more than 100 mosquito species are reported as resistant to one or

more insecticide, and more than 50 of these are anophelines (113). Insecticides

used for malaria control have included -BHC, organophosphorus, carbamate, and

pyrethroid insecticides, with the latter now taking increasing market share for

both indoor residual spraying and large-scale insecticide-impregnated bednet pro-

grams. Other insecticide groups, such as the benzylphenyl ureas and Bti, have

had limited use against mosquitoes. Resistance has tended to follow the switches

of insecticides. Resistance to -BHC/dieldrin is widespread despite the lack of use

of these insecticides for many years. Organophosphate (OP) resistance, either in

373 RESISTANCE IN INSECT VECTORS

the form of broad-spectrum OP resistance or malathion-speciﬁc resistance, occurs

in the major vectors An. culicifacies (59), An. stephensi (30, 44), An. albimanus

(3, 51), An. arabiensis (45) and An. sacharovi (54). An. culicifacies is recognized

as a species complex (101): In Sri Lanka malathion resistance occurs in An.

culicifacies species B, while in India resistance is in species B and C (59, 88).

Species B in Sri Lanka is resistant to fenitrothion, which is independent of the

malathion-speciﬁc resistance (58, 60), and is developing pyrethroid resistance

(SHPP Karunaratne, personal communication). Organophosphorus insecticide

resistance is widespread in all the major Culex vectors (53), and pyrethroid resis-

tance occurs in C. quinquefasciatus (1, 7, 19). Pyrethroid resistance has been

noted in An. albimanus (13), An. stephensi (107) and An. gambiae (18, 20, 112)

among others, while carbamate resistance is present in An. sacharovi and An.

albimanus (57). Pyrethroid resistance is widespread in Ae. aegypti (6, 48, 70) and

cases of OP and carbamate resistance have also been recorded in this species (72,

76).

The development of pyrethroid resistance in An. gambiae is particularly impor-

tant given the recent emphasis by the WHO and other organizations on the use

of pyrethroid-impregnated bednets for malaria control. Two cases of pyrethroid

resistance in An. gambiae, from the Ivory Coast and Kenya, are well documented

(18, 112). The west African focus appears to be larger and has higher levels of

resistance than that in east Africa.

Sandﬂy Resistance

The peridomestic vectors of Leishmania, Plebotomus papatasi, Lutzomyia lon-

gipalpis, and L. intermedia are controlled primarily by insecticides throughout

their range. The control of these sandﬂies is often a by-product of anti-malarial

house-spraying. The only insecticide resistance reported to date in sandﬂies is to

DDT in Indian P. papatasi (29).

Head and Body Louse Resistance

The body louse Pediculus humanus has developed widespread resistance to organ-

ochlorines (16), is malathion resistant in parts of Africa (113), and has low-level

resistance to pyrethroids in several regions (35). Resistance to organochlorine

insecticides, such as DDT and lindane, has been recorded in the human head

louse Pediculus capitis in Israel, Canada, Denmark, and Malaysia (16, 73, 113).

Permethrin has been extensively used for head louse control since the early 1980s

(77, 103). The ﬁrst reports of control failure with this insecticide were in the early

1990s in Israel (77), the Czech Republic (97), and France (21).

Simulium Resistance

Some cytospecies of the Simulium damnosum complex are vectors of onchocer-

ciasis. In 1974 the Onchocerciasis Control Programme in west Africa established

a long-term insecticide-based control program for this vector. Temephos resis-

374 HEMINGWAY n RANSON

tance occurred initially, prompting a switch to chlorphoxim, but resistance to this

insecticide occurred within a year (49). Resistance in this species is currently

being managed by a rotation of temephos, Bti, and permethrin, the insecticide

usage being determined by the rate at which water is ﬂowing in rivers forming

the major breeding sites of these vectors.

THE BIOCHEMISTRY OF RESISTANCE

Insecticide Metabolism

Three major enzyme groups are responsible for metabolically based resistance to

organochlorines, organophosphates, carbamates, and pyrethroids. DDT-dehydro-

chlorinase was ﬁrst recognized as a glutathione S-transferase in the house ﬂy,

Musca domestica (23). It has been shown to have this role commonly in anophe-

line and Aedes mosquitoes (40, 85). Esterases are often involved in organophos-

phate, carbamate, and to a lesser extent, pyrethroid resistance. Monooxygenases

are involved in the metabolism of pyrethroids, the activation and/or detoxication

of organophosphorus insecticides and, to a lesser extent, carbamate resistance.

Esterase-Based Resistance

The esterase-based resistance mechanisms have been studied most extensively at

the biochemical and molecular level in Culex mosquitoes and the aphid Myzus

persicae. Work is in progress on related and distinct esterase resistance mecha-

nisms in a range of Anopheles and Aedes species. Broad-spectrum organophos-

phate resistance is conferred by the elevated esterases of Culex. All these esterases

act by rapidly binding and slowly turning over the insecticide: They sequester

rather than rapidly metabolize the pesticide (62).

Two common esterase loci, est

and estb, are involved alone or in combination

in this type of resistance in Culex (109). In C. quinquefasciatus the most common

elevated esterase phenotype involves two enzymes, esta2

and estb2

and B

on an earlier classiﬁcation) (110). The classiﬁcation of these esterases is based

on their preferences for a-or b-naphthyl acetate, their mobility on native poly-

acrylamide gels, and their nucleotide sequence (53). Smaller numbers of C. quin-

quefasciatus populations have elevated estb1 alone, elevated esta1 alone or

co-elevated estb1 and esta3 (27, 53). When puriﬁed esta and estb from the insec-

ticide-susceptible PelSS strain were compared to various enzymes puriﬁed from

resistant strains, up to 1000-fold differences among the inhibition-kinetic con-

stants occurred for the oxon analogues of various OPs (63).

The superiority of insecticide binding in enzymes from the resistant strains

suggests that there has been positive insecticide selection pressure to maintain

elevation of favorable alleles of the esterases in insecticide-resistant insects.

Although there are minor variations between the inhibition kinetics of the differ-

ent elevated alleles, the reason why the esta2

/estb2

phenotype is so common

375 RESISTANCE IN INSECT VECTORS

(in more than 90% of resistant populations) compared to the other elevated ester-

ase phenotypes is not obvious. This advantage may be linked to a third gene,

which is co-elevated with esterases esta2/estb2 but not with the other esterase

phenotypes (50).

Metabolic studies on Culex homogenates suggests that increased rates of ester-

ase-mediated metabolism plays little or no role in resistance. One exception to

this is C. tarsalis, where two resistance mechanisms co-exist: one involving ele-

vated sequestering esterases, the other involving non-elevated metabolically

active esterases (120). In contrast to the situation in Culex, a number of Anopheles

species have a non-elevated esterase mechanism that confers resistance speciﬁ-

cally to malathion through increased rates of metabolism (44–46) (11, 68). In An.

stephensi three esterases with malathion carboxylesterase activity have been iso-

lated and characterized (47, 52).

Glutathione S-Transferase-Based Resistance

Many studies have shown that insecticide-resistant insects have elevated levels

of glutathione S-transferase activity in crude homogenates, which suggests a role

for GSTs in resistance (37, 38). GSTs are dimeric multifunctional enzymes that

play a role in detoxiﬁcation of a large range of xenobiotics (86). The enzymes

catalyze the nucleophilic attack of reduced glutathione (GSH) on the electrophilic

centers of lipophilic compounds. Multiple forms of these enzymes have been

reported for mosquitoes, house ﬂy, Drosophila, sheep blow ﬂy, and grass grub

(22, 24, 105).

Two families of insect GST are recognized, and both appear to have a role in

insecticide resistance in insects. In Ae. aegypti at least two GSTs are elevated in

DDT-resistant insects (39, 41), while in An. gambiae a large number of different

GSTs are elevated, some of which are class I GSTs (84, 85). The Ae. aegypti and

An. gambiae GSTs in resistant insects are constitutively over-expressed. The GST-

2of Ae. aegypti is over-expressed in all tissues except the ovaries of resistant

insects (39).

Monooxygenase-Based Resistance

The monooxygenases are a complex family of enzymes found in most organisms,

including insects. These enzymes are involved in the metabolism of xenobiotics

and have a role in endogenous metabolism. The P450 monooxygenase are gen-

erally the rate-limiting enzyme step in the chain. These enzymes are important in

adaptation of insects to toxic chemicals in their host plants. P450 monooxygen-

ases are involved in the metabolism of virtually all insecticides, leading to acti-

vation of the molecule in the case of organophosphorus insecticides, or more

generally to detoxiﬁcation. P450 enzymes bind molecular oxygen and receive

electrons from NADPH to introduce an oxygen molecule into the substrate.

Elevated monooxygenase activity is associated with pyrethroid resistance in

An. stephensi, An. subpictus, An. gambiae (14, 55, 112), and C. quinquefasciatus

376 HEMINGWAY n RANSON

(65). Currently this enzyme system is poorly studied in insect vectors of disease.

The nomenclature of the P450 superfamily is based on amino acid sequence

homologies, with all families having the CYP preﬁx followed by a numeral for

the family, a letter for the subfamily, and a numeral for the individual gene. To

date insect P450s have been assigned to six families: ﬁve are insect-speciﬁc and

one, CYP4, has sequence homologies with families in other organisms (8).

Target-Site Resistance

The organophosphorus, carbamates, organochlorine, and pyrethroid insecticides

all target the nervous system. Newer classes of insecticides are available for vector

control, but the high cost of developing and registering new insecticides inevitably

means that insecticides are developed initially for the agricultural market and then

utilized for public health vector control, where their activities and safety proﬁle

are appropriate and where the market is sufﬁciently large to warrant the registra-

tion costs for public health use. Compounds targeting the nicotinic acetylcholine

receptor have recently made this transition from agriculture into public health.

Acetylcholinesterase

The organophosphates and carbamates target acetylcholinesterase (AChE). AChE

hydrolyzes the excitatory neurotransmitter acetylcholine on the post-synaptic

nerve membrane. Insect AChE has a substrate speciﬁcity intermediate between

vertebrate AChE and butyrylcholinesterase. The predominant molecular form in

insects is a globular amphiphilic dimer which is membrane-bound via a glycolipid

anchor. Alterations in AChE in organophosphate-and carbamate-resistant insects

result in a decreased sensitivity to inhibition of the enzyme by these insecticides

(5, 51). The organophosphorus insecticides are converted to their oxon analogues

via the action of monooxygenases before acting as AChE inhibitors. In C. pipiens,

AChE1 and AChE2 differ in their substrate speciﬁcity, inhibitor sensitivity, and

electrophoretic migration pattern (69). Only AChE1 appears to be involved in

conferring insecticide resistance.

GABA Receptors

Resistance to dieldrin was recorded in the 1950s, but the involvement of the

GABA receptors in this resistance was not elucidated until the 1990s. The GABA

receptor in insects is a heteromultimeric gated chloride-ion channel, a widespread

inhibitory neurotransmission channel in the insect’s central nervous system and

in neuromuscular junctions (9). The insect GABA receptor is implicated as a site

of action for pyrethroids and avermectins as well as cyclodienes. Studies showing

that cyclodiene-resistant insects are resistant to picrotoxin and phenylpyrazole

insecticides, and that the effect of ivermectin on cultured neurons can be reversed

by picrotoxin pretreatment, suggest that these insecticides exert their effect by

377 RESISTANCE IN INSECT VECTORS

interacting with the chloride ionophore associated with the insect GABA receptor

(10, 62).

Sodium Channels

The pharmacological effect of DDT and pyrethroids is to cause persistent acti-

vation of the sodium channels by delaying the normal voltage-dependent mech-

anism of inactivation (100). Insensitivity of the sodium channels to insecticide

inhibition was ﬁrst recorded in Musca domestica (32). In mosquitoes there have

been many reports of suspected ‘‘kdr’’-like resistance inferred from cross resis-

tance between DDT and pyrethroids, which act on the same site within the sodium

channel. These reports have been validated by electrophysiological measurements

in Ae. aegypti and An. stephensi (48, 107).

THE MOLECULAR BIOLOGY OF RESISTANCE

Metabolic Mechanisms

Over-expression of enzymes capable of detoxifying insecticides or amino acid

substitutions within these enzymes, which alter the afﬁnity of the enzyme for the

insecticide, can result in high levels of insecticide resistance. Increased expression

of the genes encoding the major xenobiotic metabolizing enzymes are the most

common cause of insecticide resistance in mosquitoes. These large enzyme fam-

ilies may contain multiple enzymes with broad overlapping substrate speciﬁcities,

and there is a high probability that at least one member of the family will be

capable of metabolizing one or more insecticides. Increased production of these

enzymes may have a lower associated ﬁtness cost than those associated with

alterations in the structural genes because the primary function of the enzyme is

not disrupted.

Mutations in Structural Genes

In many cases of resistance caused by increased metabolism of the insecticide the

exact genetic mechanism is not known. As yet no validated reports exist of muta-

tions within detoxifying enzymes leading to resistance to insecticides in disease

vectors. Two examples have been reported in non-vector species: because both

of these mechanisms may be present in disease vectors, we describe them here.

Resistance to the organophosphate insecticide malathion is caused by a single

amino acid substitution (Trp

251

-Leu) within the E3 esterase of the sheep blow ﬂy,

Lucilia cuprina (17). Malathion-resistant strains of L. cuprina have very low

levels of activity with aliphatic esters that are conventionally used as stains for

esterase activity (80). A similar phenotype has been observed in malathion-

resistant strains of An. stephensi, An. arabiensis, and An. culicifacies. At least

three enzymes are able to metabolize malathion in An. stephensi but it is not yet

378 HEMINGWAY n RANSON

known whether a point mutation similar to that described in Lucilia is responsible

for malathion resistance in Anopheles (51).

A second distinct amino acid substitution (Gly

137

-Asp) within the active site

of the Lucilia E3 isozyme confers broad cross resistance to many organophos-

phorus insecticides but not to malathion (79). This same mutation is present in

OP-resistant strains of house ﬂy.

Gene Ampliﬁcation

The metabolic resistance mechanism studied in most detail in insect disease vec-

tors is the elevated esterase-based system in Culex (110, 111). In the three Culex

species studied to date at the molecular level the homologous estb gene is ampli-

ﬁed in resistant insects (64, 111, 114, 115). Insecticide resistance via ampliﬁcation

of genes involved in their detoxiﬁcation is common in several insects. The most

common ampliﬁed esterase-based mechanism in Culex involves the co-ampliﬁ-

cation of two esterases, est

and estb2

in C. quinquefasciatus and other mem-

bers of the C. pipiens complex worldwide (109). Other strains of C.

quinquefasciatus have est

3 and estb1 co-ampliﬁed (27), while the TEM-R strain

has ampliﬁed estb1 alone (75). Similarly, C. tritaeniorhynchus has a single ampli-

ﬁed estb, Ctrestb1 (64).

The est

and estb genes have arisen as the result of a gene duplication event,

which must have occurred prior to speciation within the Culex genus. The genes

occur in a head-to-head arrangement approximately 1.7 Kb apart in susceptible

insects (109). In resistant insects with esta2

/estb2

the ampliﬁed genes are 2.7

Kb apart, the difference being accounted for by expansion with three indels in

the intergenic spacer (109). The indels may have introduced further regulatory

elements to the amplicon intergenic spacer (52).

The identical RFLP patterns of the est

and estb2

loci in resistant C. pipiens

complex populations worldwide suggest that the ampliﬁcation of these alleles

occurred once and has since spread by migration (92). Other alleles of the esterase

A and B loci are ampliﬁed in some Culex strains. For example, a strain of C.

pipiens from Cyprus has an estimated 40 to 60 copies of the est

and estb5

genes (42) whereas in the TEM-R strain from California ampliﬁcation is found

only at the B locus. The chromosomal region containing these esterase genes

presumably represents an ampliﬁcation hot spot, a theory supported by the ampli-

ﬁcation of the homologous esterase B gene in a distinct Culex species, C. tritae-

niorhynchus (64).

An extensive study examining the spread and ﬁtness of insects containing

different esterase amplicons has been undertaken in southern France (93).

Transcriptional Regulation Esterases

Elevated levels of esterases may not always be the result of gene ampliﬁcation.

The expression of esta1 in the Barriol strain of C. pipiens from southern France

is thought to be increased due to changes in an unidentiﬁed regulatory element

379 RESISTANCE IN INSECT VECTORS

rather than underlying ampliﬁcation of the esta gene (42). Ampliﬁed esterases

can also be expressed at different levels. For example, there is fourfold more estb

than esta in resistant C. quinquefasciatus, although the genes are present in a 1:1

ratio. This difference in expression is reﬂected at the protein and mRNA level

(63, 81a).

Glutathione S-Transferases

The primary role of GSTs in mosquito insecticide resistance is in the metabolism

of DDT to nontoxic products, although they also have a secondary role in organ-

ophosphate resistance (54). GST-based DDT resistance is common in a number

of anopheline species, reﬂecting the heavy use of this insecticide for malaria

control over several decades. Molecular characterization of GSTs is most devel-

oped in An. gambiae, although work on An. dirus from Thailand suggests that a

similar arrangement of GSTs occurs. However, there is evidence for more limited

allelic diversity in this species (52, 86, 87).

Two classes of insect GSTs have been recognized and members of both classes

are important in the metabolism of insecticides in mosquitoes and other insects.

The sequence of a single class II An. gambiae GST has been published (94), and

we have cloned and sequenced the major class II GST from Ae. aegypti (D Grant,

M Wajidi, H Ranson, and J Hemingway, unpublished data). This Aedes GST,

GST-2, is over-expressed in the DDT resistant GG strain of Ae. aegypti. In this

species the resistance mutation is thought to lead to disruption of a transacting

repressor. The mutation prevents the normal function of the repressor leading to

elevated levels of GST-2 enzyme in resistant mosquitoes (39).

The insect class I GSTs are encoded by a large gene family in An. gambiae,

M. domestica, and D. melanogaster. The genomic organization of this GST class

in these three insect species is strikingly different (89). In D. melanogaster eight

divergent intronless genes are found within a 14 Kb DNA segment (106). In An.

gambiae multiple class I GST genes are also clustered in a single location. At

least one member of this family is intronless (91) but other class I GSTs in An.

gambiae contain one or more introns (Figure 1). One of these genes, aggst1a,is

alternatively spliced to produce four distinct mRNA transcripts, each of which

shares a common 58 exon with a different 38 exon (89). The products of these

spliced genes differ in their ability to metabolize DDT (91) and some of these

metabolically active GSTs are upregulated in resistant mosquitoes (90).

The organization of this class I GST gene family in insecticide-resistant and

insecticide-susceptible An. gambiae is very similar. Hence the actual GST-based

resistance mechanism is probably caused by a trans-acting regulator. The devel-

opment of a ﬁne-scale microsatellite map (119) and bacterial artiﬁcial chromo-

some (BAC) library for An. gambiae have now made this species amenable to a

positional cloning approach. Such an approach is being used to deﬁne the regu-

lator responsible for this GST resistance.

Figure 1 The class I glutathione S-transferase family of Anopheles gambiae, which includes an intronless gene aggst1-2, an alternatively

spliced gene aggst1a with four distinct viable transcripts, and the aggst1b gene with two introns.

380

381 RESISTANCE IN INSECT VECTORS

Monooxygenases

Insect microsomal P450 monooxygenases belong to six families. Increased tran-

scription of genes belonging to the CYP4, CYP6, and CYP9 families has been

observed in insecticide-resistant strains in different insect species. As yet, it is

not known which enzymes are responsible for insecticide metabolism in mos-

quitoes. Seventeen partial cDNAs encoding CYP4 P450s have been identiﬁed in

An. albimanus (98) and a similar level of diversity is present in An. gambiae (H

Ranson & F Collins, unpublished) but the role, if any, that this family plays in

resistance in the mosquito is not known. Studies on the Australian cotton boll-

worm, Helicoverpa armigera, show that the resistance-associated P450s can vary

between different strains; CYP6B2 is over-expressed in one pyrethroid-resistant

strain (118) whereas in another, CYP4G8 is over-expressed (83). Increased

expression of CYP9A1, a member of a third family, is found in the related species

Heliothis virescens (95).

There is strong evidence to suggest that P450 monooxygenase-based resistance

in M. domestica and D. melanogaster is mediated by mutations in trans-acting

regulatory genes. CYP6A8 is highly expressed in the DDT-resistant 91-R strain

of D. melanogaster but not detectable in the uninduced 91-C susceptible strain

(28, 67). Hybrids between the two strains show low levels of expression, sug-

gesting that the 91-C strain carries a repressor that suppresses transcription of

CYP6A8. A mutation in this repressor is thought to be responsible for the high

level of expression of CYP6A8 in 91-R. In the house ﬂy, CYP6D1-mediated

metabolic resistance to pyrethroids is controlled by a nearly completely dominant

cis-factor on autosome 1 and an incompletely recessive trans-factor on autosome

2 (66).

Target-Site Resistance

Non-silent point mutations within structural genes are the most common cause

of target-site resistance. For selection of the mutations to occur, the resultant

amino acid change must reduce the binding of the insecticide without causing a

loss of primary function of the target site. Therefore the number of possible amino

acid substitutions is very limited. Hence, identical resistance-associatedmutations

are commonly found across highly diverged taxa. The degree to which function

is impaired by the resistance mutation is reﬂected in the ﬁtness of resistant indi-

viduals in the absence of insecticide selection. This ﬁtness cost has important

implications for the persistence of resistance in the ﬁeld.

The main sodium and GABA channel genes in insects have been cloned and

their sequences compared in resistant and susceptible insects. Acetylcholinester-

ase-based resistance has been well characterized in Drosophila (36), but the elu-

cidation of this mechanism at the molecular level in mosquitoes has proved more

difﬁcult.

382 HEMINGWAY n RANSON

GABA Receptor Changes

The GABA receptors belong to a superfamily of neurotransmitter receptors that

also includes the nicotinic acetylcholine receptors. These receptors are formed by

the oligomerization of ﬁve subunits around a central transmitter-gated ion chan-

nel. Five different subunits have been cloned from vertebrates. To date only three

subunits have been cloned from Drosophila melanogaster, but these do not ﬁt

readily into the vertebrate GABA subunit classiﬁcation (61).

An alanine-to-serine substitution in the putative channel-lining domain of the

GABA receptor confers resistance to cyclodienes such as dieldrin (c- HCH) (34).

The mutation was ﬁrst identiﬁed in Drosophila but has since been shown to occur

in a broad range of dieldrin-resistant insects, including Ae. aegypti (104). The

only variation in resistant insects is that glycine rather than serine can sometimes

be the substituted amino acid residue. Despite the widespread switch away from

the use of cyclodiene insecticides for agricultural and public health use the resis-

tance allele is still found at relatively high frequencies in insect ﬁeld populations

(4).

Sodium Channels

A reduction in the sensitivity of the insect’s voltage-gated sodium channels to the

binding of insecticides causes the resistance phenotype known as ‘‘kdr.’’ Changes

associated with pyrethroid/DDT resistance in the sodium channels of insects are

more variable than those seen in the GABA receptors but still appear to be limited

to a small number of regions on this large channel protein.

The para sodium channel of houseﬂies contains 2108 amino acids, which fold

into 4 hydrophobic repeat domains (I-IV) separated by hydrophilic linkers. The

ﬁrst mutation to be characterized in kdr insects was a leucine to phenylalanine

point mutation in the S6 transmembrane segment of domain II in the sodium

channel sequence of M. domestica (116, 117) which produces 10- to 20-fold

resistance to DDT and pyrethroids. In ‘‘super-kdr’’ houseﬂies, this mutation also

occurs with a second methionine to threonine substitution further upstream in the

same domain, resulting in more than 500-fold resistance (117). Analysis of the

domain region of the para-sodium channel gene in pyrethroid-resistant An. gam-

biae from the Ivory Coast showed an identical Leu to Phe mutation in this species

(71).

A PCR-based diagnostic test discriminates between homozygous-susceptible,

homozygous-resistant, and heterozygous individuals with the Leu to Phe mutation

(71). Because kdr is semi-recessive or fully recessive (81), the ability to detect

heterozygotes is of paramount importance in the early detection and management

of resistance in the ﬁeld.

The limited number of changes associated with kdr-type resistance may be

constrained by the number of modiﬁcations that can inﬂuence pyrethroid/DDT

binding to the sodium channels. However, a note of caution should be added:

383 RESISTANCE IN INSECT VECTORS

There is already a tendency to investigate pyrethroid-resistant insects with a PCR

approach conﬁned to regions where a kdr mutation has already been seen. Hence

consistent resistance-associated changes in other parts of the sodium-channel gene

could be missed. A different approach to isolating kdr-type mutants has been used

in Drosophila, utilizing the relative ease with which large numbers of mutants in

the para sodium channel gene can be isolated based on their temperature sensi-

tivity. Two classes of these mutations are in positions equivalent to the kdr and

super-kdr mutations in different domains. The third class is in a novel position

(Figure 2) (34).

Acetylcholinesterase

Vertebrates have two cholinesterases: acetylcholinesterase and butyrylcholines-

terase. In D. melanogaster only a single cholinesterase gene, Ace, coding for

acetycholinesterase has been cloned, based on a knowledge of its location via

Figure 2 The location of amino acid substitutions occurring in the para-sodium channel

gene of resistant strains of various insects. Mutations that have been documented in ﬁeld

strains of medical vectors are shown in A. B shows replacements in Drosophila isolated

on the basis of temperature sensitivity paralytic phenotypes which confer DDT/pyrethroid

resistance (adapted from 34).

384 HEMINGWAY n RANSON

isolation of a range of different mutants (43). A range of different amino acid

substitutions in the Ace genes of Drosophila and the house ﬂy M. domestica

putatively cause resistance (reviewed in 33). Many of these resistance-associated

residues are predicted to lie close to or within the acetylcholinesterase active site

gorge (78). In C. pipiens at least two cholinesterase genes occur, both with ace-

tylcholinesterase-like activity (69). A similar organization occurs in Amphioxus.

To date the only acetylcholinesterase to have been cloned in C. pipiens is AChE2,

which is not involved in insecticide resistance (69). The AChE2 gene is sex

linked, unlike the resistance-conferring AChE1, which is autosomal. AChE genes

have been cloned from the mosquitoes Ae. aegypti and An. stephensi, but both of

these genes are also sex linked (2, 99).

As yet, there is no recorded AChE-based resistance mechanism in An. ste-

phensi, but to date none of the acetylcholinesterase resistances recorded in insects

have been sex linked, suggesting that these genes do not represent the insecticidal

target site. However, detailed analysis of inhibition proﬁles for acetylcholinester-

ase from Ae. aegypti suggests that there is only one AChE locus in this species.

If such is the case, altered acetylcholinesterase-based resistance in this species

should be sex linked. At least one case of altered AChE has been reported in Ae.

aegypti from Trinidad (108).

Five point mutations associated with resistance to organophosphorus and car-

bamate insecticides have been identiﬁed within the D. melanogaster acetylcho-

linesterase gene (78) and site-directed mutagenesis of the sex-linked AChE from

Ae. aegypti has demonstrated that these same mutations also confer resistance in

the mosquito enzyme (74). However, none of these mutations have been identiﬁed

in ﬁeld-collected or laboratory-selected strains of mosquitoes.

Insensitive AChE has a severe ﬁtness cost in C. pipiens populations in southern

France (31), which is probably caused by a reduction in the AChE activity of the

mutated enzyme compared to the wild type.

MANAGEMENT OF INSECTICIDE-RESISTANT VECTOR

POPULATIONS

The practice of using an insecticide until resistance becomes a limiting factor is

rapidly eroding the number of suitable insecticides for vector control. Rotations,

mosaics, and mixtures have all been proposed as resistance management tools

(25, 26, 96). Numerous mathematical models have been produced to estimate

how these tools should be optimally used (102). However, these models have

rarely been tested under ﬁeld conditions for insect vectors, due to the practical

difﬁculties in estimating changes in resistance gene frequencies in large samples

of insects (56). With the advent of different biochemical and molecular techniques

for resistance-gene frequency estimation, ﬁeld trials of resistance management

strategies have become more feasible.

385 RESISTANCE IN INSECT VECTORS

A large-scale trial of the use of rotations or mosaics of insecticides compared

to single use of DDT or a pyrethroid is currently underway in Mexico (56, 82).

Changes in resistance gene frequencies in An. albimanus are being monitored

over a four-year period (82). Information resulting from such large-scale trials

may allow us to establish rational strategies for long-term insecticide use in dis-

ease control programs.

As our ability to manipulate the insect genome improves and our understanding

of the regulation of insecticide resistance mechanisms increase, new strategies

should be devised for incorporating this new knowledge into these control

programs.

Visit the Annual Reviews home page at www.AnnualReviews.org.

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