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Keywords:

  • insects;
  • insecticide resistance;
  • detoxification;
  • glutathione transferases

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

Glutathione transferases (GSTs) are a diverse family of enzymes found ubiquitously in aerobic organisms. They play a central role in the detoxification of both endogenous and xenobiotic compounds and are also involved in intracellular transport, biosynthesis of hormones and protection against oxidative stress. Interest in insect GSTs has primarily focused on their role in insecticide resistance. GSTs can metabolize insecticides by facilitating their reductive dehydrochlorination or by conjugation reactions with reduced glutathione, to produce water-soluble metabolites that are more readily excreted. In addition, they contribute to the removal of toxic oxygen free radical species produced through the action of pesticides. Annotation of the Anopheles gambiae and Drosophila melanogaster genomes has revealed the full extent of this enzyme family in insects. This mini review describes the insect GST enzyme family, focusing specifically on their role in conferring insecticide resistance.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

The glutathione transferases (GSTs) are a large family of multifunctional enzymes involved in the detoxification of a wide range of xenobiotics including insecticides (Salinas & Wong, 1999). GSTs primarily catalyse the conjugation of electrophilic compounds with the thiol group of reduced glutathione (GSH), generally making the resultant products more water soluble and excretable than the non-GSH conjugated substrates (Habig et al., 1974). In addition, some GSTs catalyse a dehydrochlorination reaction using reduced glutathione as a cofactor rather than a conjugate (Clark & Shamaan, 1984). GSTs are important in cancer epidemiology and drug resistance (Tew, 1994; Hayes & Pulford, 1995) and hence are well studied in mammals. The majority of studies on insect GSTs have focused on their role in detoxifying foreign compounds, in particular insecticides and plant allelochemicals and, more recently, their role in mediating oxidative stress responses. (Clark et al., 1986; Wang et al., 1991; Fournier et al., 1992; Ranson et al., 2001; Vontas et al., 2001; Sawicki et al., 2003).

Classification and nomenclature

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

There are at least two ubiquitously distributed distantly related groups of GSTs, classified according to their location within the cell: microsomal and cytosolic. A third group of GSTs, the Kappa class, are located in mammalian mitochondria and peroxisomes (Morel et al., 2004; Lander et al., 2004) and are structurally distinct from the microsomal and cytosolic GSTs (Robinson et al., 2004). To date, genome data mining has failed to detect any members of this GST class in insects.

A single microsomal GST gene is present in the genome of the fruitfly Drosophila melanogaster whereas the mosquito Anopheles gambiae has three microsomal GST genes (Toba & Aigaki, 2000; Ranson et al., 2002). The microsomal GSTs are trimeric, membrane-bound proteins. Although very different in structure and in origin, the microsomal GSTs catalyse similar reactions to the cytosolic GSTs (Gakuta & Toshiro, 2000; Prabhu et al., 2001). Microsomal GSTs have not been implicated in the metabolism of insecticides and will not be discussed further in this review.

Insect cytosolic GSTs were initially assigned numbers according to their order of elution from the various purification procedures employed or isoelectric points (Clark et al., 1985; Prapanthadara et al., 1993). Later two immunologically distinct classes of GSTs were recognized in houseflies and designated as class I and class II (Fournier et al., 1992). The class II insect GSTs are encoded by a single gene in all species studied to date (Beall et al., 1992; Reiss & James, 1993; Synder et al., 1995) although two distinct transcripts are produced by alternative splicing of the A. gambiae class II gene (Ding et al., 2003). The class I insect GSTs, in contrast, are encoded by a multigene family in Anopheles mosquitoes, D. melanogaster and Musca domestica (Toung et al., 1993; Zhou & Syvanen, 1997; Ranson et al., 2002).

As the volume of insect sequence data increased, additional GSTs were identified that did not clearly fit within class I or II and the mammalian system of GST nomenclature was adopted whereby classes are designated using Greek letters (Chelvanayagam et al., 2001). Phylogenetic comparison of insect and mammalian GST genes showed that the insect class II GSTs are orthologous to the Sigma GST class found in a diverse range of species from netmatodes to mammals (Agianian et al., 2003). In contrast, the class I GSTs are unique to insects and were re-named Delta GSTs. A second large class of GSTs, the Epsilon class, is also restricted to insects (Ranson et al., 2001). The Delta and Epsilon GST classes have expanded independently in D. melanogaster and A. gambiae, suggesting that these enzymes play important roles in the adaptation of these species to their specific environments (Ranson et al., 2002).

The majority of the remaining cytosolic insect GSTs are members of the Zeta, Theta and Omega classes (Board et al., 1997, 2000; Ranson et al., 2002). The relatively high degree of conservation of these GST genes across taxa suggests that they play essential steps in conserved physiological pathways. The complete inventory of A. gambiae cytosolic GSTs includes three genes, GSTu1, GSTu2 and GSTu3, that cannot be readily assigned to the existing classes. They share less than 40% amino acid idenitiy with other insect GSTs and are physically separate from other members of the GST gene family on the mosquito chromosomes (Ding et al., 2003). These may represent novel classes of insect GSTs but have been temporarily designated unclassified (u). The phylogenetic relationship between the different classes of GSTs found in insects is shown in Fig. 1.

image

Figure 1. Neighbor joining tree illustrating the relationship between the classes of GSTs found in insects. The amino acid sequence of all cytosolic GSTs from Anopheles gambiae and Drosophila melanogaster were aligned with GSTs from mammals, plants and nematodes from the Zeta, Sigma, Omega and Theta classes using ClustalW. For clarity, a single GST peptide was selected for each class from the non-insect phyla (accession numbers and wormbase protein IDs: HsZeta, O43708; CeZeta, WP:Ce25559; AtZeta, AF288182; CeSigma, WP:Ce04834; HsSigma, O60760; CeOmega, WP:Ce15815; HsOmega, P78417; AtTheta, AJ131580; HsTheta, S44358). The tree was constructed by the neighbor-joining method from a similarity matrix of pair-wise comparisons made using the Jukes–Cantor algorithm Bootsrap values (500 replicates) are shown at selected nodes. The nodes of highly supported insect GST classes have been collapsed. Those insect GSTs whose classifcation is unresolved are shown in bold. Ag = Anopheles gambiae; At = Arabidopsis thaliana; Ce = Caenorhabditis elegans; Dm = Drosophila melanogaster; Hs = Homo sapiens. See text for further details.

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By extending the mammalian GST classification system to encompass insect GSTs the nomenclature of individual insect GST genes and proteins has been clarified (Chelvanayagam et al., 2001). Individual GST subunits are now assigned names indicating the species from which they were isolated and the GST class. They are also given a number that may reflect the order of discovery or the genome organizations. For example, AgGSTd12 is the twelfth member of the A. gambiae Delta class of GST subunits to be identified. The distribution of different classes of GSTs in insects and other animals is shown in Table 1.

Table 1.  Summary of insect GST classes
GST classDistributionPutative transcript no. in A. gambiaePutative transcript no. in D. melanogasterExamples in other insect species
DeltaInsects only1511L. cuprina, M. domestica, N. lugens, C. variipennis
EpsilonInsects only 814P. xylotstella, M. sexta, M. domestica
OmegaNematodes, insects, mammals 1 5 
ThetaMammals, insects and plants 2 4 
SigmaMolluscs, helminths, nematodes, insects, mammals 1 1M. domestica, B. germanica, C. fumiferana, M. sexta
ZetaPlants, nematodes. Insects, mammals 1 2 

Enzyme structure

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

Cytosolic GSTs are hetero- or homo-dimeric proteins approximately 25 kDa in size. The polypeptide chain of each monomer folds into two domains joined by a variable linker region. The N-terminal domain (residues 1–80) consists of four beta sheets and three flanking alpha helices and adopts a conformation similar to the thioredoxin domain found in many proteins that bind GSH or cysteine (see reviews by Dirr et al., 1994; Wilce & Parker, 1994; Sheehan et al., 2001). This domain contains the majority of residues involved in the binding of glutathione (the G-site). The larger C-terminal domain consists of a variable number of alpha helices. The variable hydrophobic H-site, which interacts with the electrophilic substrates, is largely formed from residues in the C-terminal domain. Although each monomeric active site functions independently, their quaternary structure is essential for their activity (Danielson & Mannervik, 1985).

Mode of action

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

In a GST-catalysed conjugation reaction, one molecule of reduced GSH and one molecule of a second substrate are combined to form a thioester. The reaction proceeds via substrate binding, the activation of the thiol group of GSH and subsequent nucleophilic attack by the anionic GSH on the bound hydrophobic compound (Atkins et al., 1993). This conjugation neutralizes the electrophilic sites of the lipophilic substrate and protects the cellular components, especially the nucleophilic oxygen and nitrogen of DNA from electrophilic attack of nucleophiles. Conjugation also renders the product more water soluble and therefore more readily excretable from the cell.

GSTs have a high affinity towards GSH and because this tripiptide is present at high intracellular concentrations the GSH binding site of GST may always be occupied. The ‘active site residue’ in the N-terminal domain interacts with and activates the sulphydryl group of glutathione. In most mammalian GSTs the active site residue is a tyrosine (Karshikoff et al., 1993; Wilce & Parker, 1994) but in the Delta and Epsilon insect GST classes this role is performed by a serine residue (Board et al., 1995).

Gene organization

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

The diversity of reactions catalysed by GSTs is a function of the broad substrate specificities of many individual GST enzymes and the extensive nature of the GST supergene family. In insects, local gene duplications, particularly within the insect-specific Delta and Epsilon classes, have resulted in expansions of the GST family. Because substitution of a small number of amino acids can have dramatic effects on the substrate specificity of the enzymes (Ortelli et al., 2003), the process of gene duplication, diversification and selection can tailor the repetoire of reactions catalysed by GSTs to suit the particular ecological niche occupied by a species (Ranson et al., 2002).

Futher GST diversity is generated by alternative spicing in Anopheles mosquitoes and genetic rearrangements leading to gene fusions in Musca domestica. Two GST genes are alternatively spliced in A. gambiae: four distinct peptides with differing catalytic properties are generated from the Delta class AgGSTD1 gene (Ranson et al., 1998) and two transcripts, sharing two common 5′ exons but differing in their use of 3′ exons, are generated from the single Sigma GST gene in this species (Ding et al., 2003). The genome of M. domestica contains multiple intronless loci encoding Delta GSTs, some of which appear to have resulted from the fusion between the 5′ and 3′ ends of different Delta GST genes. It is not known whether all of these housefly GST genes encode functional proteins but it is possible that a rare recombination mechanism may be contributing to GST diversity in this species (Zhou & Syvanen, 1997).

Regulation of GST expression

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

In non-insect species, many GST enzymes are differentially regulated in response to various inducers or environmental signals or in a tissue- or developmental-specific manner. A similarly complex pattern of regulation is expected for insect GSTs. Two review articles have described the effect of various dietary compounds, insecticides and laboratory inducers on general GST expression (Clark, 1989; Yu, 1996). Now that the full extent of the GST family is known for two insect species, more specific studies can be conducted to determine the factors regulating expression of individual GST genes.

Levels of GST activity vary throughout the life stage of insects. For example, in Aedes aegypti, total GST activity measured with 1-chloro-2,4-dinitrobenzene (CDNB) and 1,2-dichloronitrobenzene (DCNB) increases during larval development, reaching its peak in the pupal stage and declining in adults as they age (Hazelton & Laing, 1983). In a preliminary investigation of the expression profiles of A. gambiae GSTs, transcripts were detectable for all but one of the genes in 1-day-old adults (Ding et al., 2003). No attempt was made to quantify the expression level in different developmental stages in A. gambiae but it is apparent from studies in other insects that the levels of individual enzymes can fluctuate widely during the lifespan of an insect. For example, a Sigma GST from the spruce budworm, Choristoneura fumiferana, is expressed at very low levels in feeding larvae but high levels in diapausing larvae (Feng et al., 1999).

Variations in the level of GST activity in different insect tissues have been reported in several species. In cases where the variation in activity is attributed to individual enzymes, such studies can provide valuable insights into the functions of different GSTs. Thus the finding that Sigma GSTs from housefly and Drosophila were predominately located in the indirect flight muscles, in association with troponin H, suggested that the role of this GST class was structural rather than catalytic (subsequently, however, these GSTs have been found to play a very important role in protection against oxidative stress) (Franciosa & Berge, 1995; Singh et al., 2001). Very high levels of GST activity have been reported in the fat body and midguts of insects. Both these tissues are important sites for the detoxification of xenobiotics.

GSTs and insecticide resistance

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

Elevated GST activity has been associated with resistance to all the major classes of insecticides (Prapanthadara et al., 1993; Huang et al., 1998; Vontas et al., 2001). In many cases, the individual GST enzyme(s) involved in resistance have not been identified and GSTs have been implicated by association only (i.e. an increase in GST activity, detected using model substrates, in insecticide-resistant strains of insects vs. their susceptible counterparts). In cases that have been studied in more detail, resistance has been attributed to increases in the amount of one or more GST enzymes, either as a result of gene amplification or more commonly through increases in transcriptional rate, rather than qualitative changes in individual enzymes (Grant & Hammock, 1992; Ranson et al., 2001).

Dehydrochlorination is an important mechanism for DDT detoxification. This reaction is catalysed by GSTs (Clark & Shamaan, 1984). The proposed mechanism is shown in Fig. 2. A GSH conjugate of DDT has never been identified, but this tripeptide is an essential cofactor in the reaction. The thiolate anion generated in the active site of the GST acts as a general base and abstracts a hydrogen atom from DDT resulting in the elimination of chlorine to generate DDE. Other organochlorine insecticides, e.g. lindane, are detoxified by conjugation to glutathione.

image

Figure 2. Model of DDT dehydrochlorinase activity of GSTs (after Matsumura, 1985).

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Increased rates of DDT dehydrochlorination confer resistance to DDT in many insect species including houseflies and the mosquitoes Ae. aegypti, A. gambiae and A. dirus (Prapanthadara et al., 1993, 1996; Grant et al., 1991). In Ae. aegypti, two immunologically distinct forms of GST are overproduced in a DDT-resistant strain (Grant et al., 1991). Overexpression of at least one of these is thought to be controlled by a mutation in a trans-acting repressor in the DDT-resistant strain (Grant & Hammock, 1992) but the nature of this repressor is not known. Multiple Epsilon class GSTs from A. gambiae are overexpressed in a DDT-resistant strain and one of these, GSTe2, encodes an enzyme that has the highest levels of DDT dehydrochlorinase activity reported for any GST (Ortelli et al., 2003). Genetic mapping of the major loci conferring DDT resistance in A. gambiae implicate both cis- and trans-acting factors in the overexpression of the Epsilon class GSTs (Ranson et al., 2000).

GSTs are responsible for many cases of organophosphate resistance (Hayes & Wolf, 1988). The conjugation of glutathione to organophosphate insecticides results in their detoxification via two distinct pathways. In O-dealkylation, glutathione is conjugated with the alkyl portion of the insecticide, e.g. the demethylation of tetrachlorvinphos in resistant houseflies (Oppenoorth et al., 1979). In the second mechanism, O-dearylation, the glutathione reacts with the leaving group, e.g. the detoxification of parathion and methyl parathion in the diamondback moth Plutella xylostella (Chiang & Sun, 1993). Recombinant GST enzymes from the diamondback moth and housefly have verified the role of these enzymes in the metabolism of organophosphate insecticides (Huang et al., 1998; Wei et al., 2001).

GSTs have not yet been implicated in the direct metabolism of pyrethroid insecticides. Nevertheless, they may play an important role in conferring resistance to this insecticide class by detoxifying lipid peroxidation products induced by pyrethroids (Vontas et al., 2001). A Delta class GST has been cloned from a pyrethroid resistant strain of Nilaparvata lugens. The recombinant protein produced from this gene has high peroxidase activity and it has been proposed that this enzyme is involved in the prevention or repair of oxidative damage induced by insecticide exposure (Vontas et al., 2002). Elevated GSTs in other resistant insects may play a similar role. GSTs may also protect against pyrethroid toxicity in insects by sequestering the insecticide (Kostaropoulos et al., 2001).

Summary

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

Advances in genetics and biochemistry have revealed the complexity of the insect GST family. Specific functions of some individual GSTs, particularly their role in detoxifying xenobiotics, have been identified but much remains to be learnt about the endogenous substrates of insect GSTs. Deciphering the precise expression profile of each GST gene may provide clues to function that can subsequently be elucidated in in vitro systems.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References

This work was funded in part by a Royal Society Dorothy Hodgkin Fellowship (to H.R.) and a Wellcome Trust Project Grant (to J.H. and H.R.).

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Classification and nomenclature
  5. Enzyme structure
  6. Mode of action
  7. Gene organization
  8. Regulation of GST expression
  9. GSTs and insecticide resistance
  10. Summary
  11. Acknowledgements
  12. References
  • Agianian, B., Tucker, P.A., Schouten, A., Leonard, K., Bullard, B. and Gros, P. (2003) Structure of a Drosophila Sigma class glutathione S-transferase reveals a novel active site topography suited for lipid peroxidation products. J Mol Biol 326: 151165.
  • Atkins, W.M., Wang, R.W., Bird, A.W., Newton, D.J. and Lu, A.H.W. (1993) The catalytic mechanism of glutathione S-transferase (GST): spectroscopic determination of the pKa of Tyr-9 in rat alpha 1–1 GST. J Biol Chem 268: 1918811991.
  • Beall, C., Fyrberg, C., Song, S. and Fyrberg, E. (1992) Isolation of a Drosophila gene encoding glutathione S-transferase. Biochem Genet 30: 515527.
  • Board, P.G., Baker, R.T., Chelvanayagam, G. and Jermiin, L.S. (1997) Zeta, a novel class of glutathione transferase in a range of species from plants to humans. Biochem J 328: 929935.
  • Board, P.G., Coggan, M., Chelvanayagam, G., Easteal, S., Jermiin, L.S., Schulte, G.K., Danley, D.E., Hoth, L.R., Griffor, M.C., Kamath, A.V., Rosner, M.H., Chrunyk, B.A., Perregaux, D.E., Gabel, C.A., Geoghegan, K.F. and Pandit, J. (2000) Identification, characterization, and crystal structure of the Omega class glutathione transferases. J Biol Chem 275: 2479824806.
  • Board, P.G., Coggan, M., Wilce, M.C. and Parker, M.W. (1995) Evidence for an essential serine residue in the active site of the Theta class glutathione transferases. Biochem J 311: 247250.
  • Chelvanayagam, G., Parker, M.W. and Board, P.G. (2001) Fly fishing for GSTs: a unified nomenclature for mammalian and insect glutathione transferases. Chem Biol Interact 133: 256260.
  • Chiang, F.-M. and Sun, C.-N. (1993) Glutathione transferase isozymes of diamondback moth larvae and their role in the degradation of some organophosphorus insecticides. Pest Biochem Physiol 45: 714.
  • Clark, A.G. (1989) The comparative enzymology of the glutathione S-transferases from non-vertabrate organisms. Comp Biochem Physiol 92: 419446.
  • Clark, A.G., Dick, G.L., Martindale, S.M. and Smith, J.N. (1985) Glutathione S-transferases from the New Zealand grass grub, Costelytra zealandica. Insect Biochem 15: 3544.
  • Clark, A.G. and Shamaan, N.A. (1984) Evidence that DDT-dehydrochlorinase from the house fly is a glutathione S-transferase. Pest Biochem Physiol 22: 249261.
  • Clark, A.G., Shamaan, N.A., Sinclair, M.D. and Dauterman, W.C. (1986) Insecticide metabolism by multiple glutathione S-transferases in two strains of the house fly, Musca domestica (L.). Pest Biochem Physiol 25: 169175.
  • Danielson, U.H. and Mannervik, B. (1985) Kinetic independence of the subunits of cytosolic glutathione transferase from the rat. Biochem J 231: 263267.
  • Ding, Y., Ortelli, F., Rossiter, L.C., Hemingway, J. and Ranson, H. (2003) The Anopheles gambiae glutathione transferase supergene family: annotation, phylogeny and expression profiles. BMC Genomics 4: 35.
  • Dirr, H., Reinemer, P. and Huber, R. (1994) X-ray crystal structures of cytosolic glutathione S-transferases. Implications for protein architecture, substrate recognition and catalytic function. Eur J Biochem 220: 645661.
  • Feng, Q.L., Davey, K.G., Pang, A.S.D., Primavera, M., Ladd, T.R., Zheng, S.C., Sohi, S.S., Retnakaran, A. and Palli, S.R. (1999) Glutathione S-transferase from the spruce budworm, Chorstoneura fumiferana: identification, characterization, localization, cDNA cloning and expression. Insect Biochem Mol Biol 29: 779793.
  • Fournier, D., Bride, J.M., Poire, M., Berge, J.B. and Plapp, F.W. (1992) Insect glutathione S-transferases. Biochemical characteristics of the major forms from houseflies susceptible and resistant to insecticides. J Biol Chem 267: 18401845.
  • Franciosa, H. and Berge, J.B. (1995) Glutathione S-transferases in housefly (Musca domestica): location of GST-1 and GST-2 families. Insect Biochem Mol Biol 25: 311317.
  • Gakuta, T. and Toshiro, A. (2000) Disruption of the microsomal glutathione S-transferase-like gene reduces life span of Drosophila melanogaster. Gene 253: 179187.
  • Grant, D.F., Dietze, E.C. and Hammock, B.D. (1991) Glutathione S-transferase isozymes in Aedes aegypti: purification, characterization, and isozyme specific regulation. Insect Biochem 4: 421433.
  • 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: 169176.
  • Habig, W.H., Pabst, M.J. and Jakoby, W.B. (1974) Glutathione S-transferases. J Biol Chem 249: 71307139.
  • Hayes, J.D. and Pulford, D.J. (1995) The glutathione S-transferase supergene family – Regulation of GST and the contribution of the isoenzymes to cancer chemoprotection and drug resistance. Crit Rev Biochem Mol Biol 30: 445600.
  • Hayes, J.D. and Wolf, C.R. (1988) Role of glutathione transferase in drug resistance. In Glutathione Conjugation: Mechanisms and Biological Significance (Sies, H. and Ketterer, B., eds), pp. 315355. Academic Press Ltd, London.
  • Hazelton, G.A. and Laing, C.A. (1983) Glutathione S-transferase activities in the yellow-fever mosquito [Aedes aegypti (Loisville)] during growth and aging. Biochem J 210: 281287.
  • Huang, H.S., Hu, N.T., Yao, Y.E., Wu, C.Y., Chiang, S.W. and Sun, C.N. (1998) Molecular cloning and heterologous expression of a glutathione S- transferase involved in insecticide resistance from the diamondback moth, Plutella xylostella. Insect Biochem Mol Biol 28: 651658.
  • Karshikoff, A., Reinemer, P., Huber, R. and Ladenstein, R. (1993) Electrostatic evidence for the activation of the glutathione thiol by Tyr7 in pi-class glutathione transferases. Eur J Biochem 215: 663670.
  • Kostaropoulos, I., Papadopoulos, A.I., Metaxakis, A., Boukouvala, E. and Papadopoulou-Mourkidou, E. (2001) Glutathione S-transferase in the defence against pyrethroids in insects. Insect Biochem Mol Biol 31: 313319.
  • Lander, J.E., Parsons, J.F., Rife, C.L., Gilliland, G.L. and Armstrong, R.N. (2004) Parallel evolutionary pathways for glutathione transferases: structure and mechanims of the mitochondrial class Kappa enzyme rGSTK1–1. Biochemistry 43: 352261.
  • Matsumura, F. (1985) Toxicology of Insecticides. Plenum Press, New York.
  • Morel, F., Rauch, C., Petit, E., Piton, A., Theret, N., Coles, B. and Guillouzo, A. (2004) Gene and protein characterization of the human glutathione S-transferase kappa and evidence for a peroxisomal localization. J Biol Chem 279: 1624616253.
  • Oppenoorth, F.J., Van der Pas, L.J.T. and Houx, N.W.H. (1979) Glutathione S-transferase and hydrolytic activity in a tetrachlorvinphos-resistant strain of housefly and their influence on resistance. Pest Biochem Physiol 11: 176188.
  • Ortelli, F., Rossiter, L.C., Vontas, J., Ranson, H. and Hemingway, J. (2003) Heterologous expression of four glutathione transferase genes genetically linked to a major insecticide-resistance locus from the malaria vector Anopheles gambiae. Biochem J 373: 957963.
  • Prabhu, K.S., Reddy, P.V., Gmpricht, G., Hildenbrandt, G.R., Scholz, R.W., Sordillo, L.M. and Reddy, C.C. (2001) Microsomal glutathione S-transferase A1–1 with glutathione peroxidase activity from sheep liver: molecular cloning, expression and characterization. Biochem J 360: 345354.
  • Prapanthadara, L., Hemingway, J. and Ketterman, A.J. (1993) Partial purification and characterization of glutathione S-transferase involved in DDT resistance from the mosquito Anopheles gambiae. Pest Biochem Physiol 47: 119133.
  • Prapanthadara, L., Koottathep, S., Promtet, N., Hemingway, J. and Ketterman, A.J. (1996) Purification and characterization of a major glutathione S-transferase from the mosquito Anopheles dirus (Species B). Insect Biochem Mol Biol 26: 277285.
  • Ranson, H., Claudianos, C., Ortelli, F., Abgrall, C., Hemingway, J., Sharakhova, M.V., Unger, M.F., Collins, F.H. and Feyereisen, R. (2002) Evolution of supergene families associated with insecticide resistance. Science 298: 179181.
  • Ranson, H., Collins, F. and Hemingway, J. (1998) The role of alternative mRNA splicing in generating heterogeneity within the Anopheles gambiae class I glutathione S-transferase family. Proc Natl Acad Sci USA 95: 1428414289.
  • Ranson, H., Jensen, B., Wang, X., Prapanthadara, L., Hemingway, J. and Collins, F.H. (2000) Genetic mapping of two loci affecting DDT resistance in the malaria vector, Anopheles gambiae. Insect Mol Biol 9: 499507.
  • Ranson, H., Rossiter, L., Ortelli, F., Jensen, B., Wang, X., Roth, C.W., Collins, F.H. and Hemingway, J. (2001) Identification of a novel class of insect glutathione S-transferases involved in resistance to DDT in the malaria vector Anopheles gambiae. Biochem J 359: 295304.
  • Reiss, R.A. and James, A.A. (1993) A glutathione S-transferase gene of the vector mosquito, Anopheles gambiae. Insect Mol Biol 2: 2532.
  • Robinson, A., Huttley, G.A., Booth, H.S. and Board, P.G. (2004) Modelling and bioinformatics studies of the human Kappa-class glutathione transferase predict a novel third glutathione transferase family with similarity to prokaryotic 2-hydroxychromene-2-carboxylate isomerases. Biochem J 379: 541552.
  • Salinas, A.E. and Wong, M.G. (1999) Glutathione S-transferases – A review. Current Med Chem 6: 279309.
  • Sawicki, R., Singh, S.P., Mondal, A.K., Benes, H. and Zimniak, P. (2003) Cloning, expression and biochemical characterization of one Epsilon-class (GST-3) and ten Delta-class (GST-1) glutathione S-transferases from Drosophila melanogaster, and identification of additional nine members of the Epsilon class. Biochem J 370: 661669.
  • Sheehan, D., Meade, G., Foley, V.M. and Dowd, C.A. (2001) Structure, function and evolution of glutathione transferases: implications for classification of non-mammalian members of an ancient enzyme superfamily. Biochem J 360: 116.
  • Singh, S.P., Coronella, J.A., Benes, H., Cochrane, B.J. and Zimniak, P. (2001) Catalytic function of Drosophila melanogaster glutathione S-transferase DmGSTS1–1 (GST-2) in conjugation of lipid peroxidation end products. Eur J Biochem 268: 29122923.
  • Synder, M.J., Walding, J.K. and Feyereisen, R. (1995) Glutathione S-transferases from larval Manduca sexta midgut: sequense of two cDNAs and enzyme induction. Insect Biochem Mol Biol 25: 455465.
  • Tew, K.D. (1994) Glutathione-associated enzymes in anticancer drug resistance. Cancer Res 54: 43134320.
  • Toba, G. and Aigaki, T. (2000) Disruption of the microsomal glutathione S-transferase-like gene reduces life span of Drosophila melanogaster. Gene 253: 179187.
  • Toung, Y.-P.S., Hsieh, T. and Tu, C.-P.D. (1993) The glutathione S-transferase D genes: a divergently organized, intronless gene family in Drosophila melanogaster. J Biol Chem 268: 97379746.
  • Vontas, J.G., Small, G.J. and Hemingway, J. (2001) Glutathione S-transferases as antioxidant defence agents confer pyrethroid resistance in Nilaparvata lugens. Biochem J 357: 6572.
  • Vontas, J.G., Small, G.J., Nikou, D.C., Ranson, H. and Hemingway, J. (2002) Purification, molecular cloning and heterologous expression of a glutathione S-transferase involved in insecticide resistance from the rice brown planthopper, Nilaparvata lugens. Biochem J 362: 329337.
  • Wang, J.Y., McCommas, S. and Syvanen, M. (1991) Molecular cloning of a glutathione S-transferase overproduced in an insecticide-resistant strain of the housefly (Musca domestica). Mol Gen Genet 227: 260266.
  • Wei, S.H., Clark, A.G. and Syvanen, M. (2001) Identification and cloning of a key insecticide-metabolizing glutathione S-transferase (MdGST-6A) from a hyper insecticide-resistant strain of the housefly Musca domestica. Insect Biochem Mol Biol 31: 11451153.
  • Wilce, M.C.J. and Parker, M.W. (1994) Structure and function of glutathione S-transferases. Biochem Biophys Acta 1205: 118.
  • Yu, S.J. (1996) Insect gutathione S-transferases. Zool Studies 35: 919.
  • Zhou, Z.H. and Syvanen, M. (1997) A complex glutathione transferase gene family in the housefly Musca domestica. Mol Gen Genet 256: 187194.