Genomic analysis of detoxification genes in the mosquito Aedes aegypti

ARTICLE IN PRESS Insect Biochemistry and Molecular Biology Insect Biochemistry and Molecular Biology 38 (2008) 113–123 www.elsevier.com/locate/ibmb

Genomic analysis of detoxification genes in the mosquito Aedes aegypti Clare Strodea, Charles S. Wondjia, Jean-Philippe Davidb, Nicola J. Hawkesa, Nongkran Lumjuanc, David R. Nelsond, David R. Dranee, S.H.P. Parakrama Karunaratnef, Janet Hemingwaya, William C. Black IVg, Hilary Ransona, a Vector Group, Liverpool School of Tropical Medicine, Liverpool L3 5QA, UK Laboratoire d’Ecologie Alpine, UMR 5553, Equipe Perturbations Environnementales et Xe´nobiotiques, BP 53, 38041 Grenoble Cedex 9, France c Research Institute for Health Sciences, Chiang Mai University, P.O. Box 80, Chiang Mai 50202, Thailand d Department of Molecular Sciences and Center of Excellence for Genomics and Bioinformatics, University of Tennessee, Memphis, TN 38163, USA e Aerospace Engineering, University of Tennessee, Knoxville, TN 37996, USA f Department of Zoology, University of Peradeniya, Peradeniya, Sri Lanka g Department of Microbiology, Immunology and Pathology, Colorado State University, Fort Collins, CO, USA b

Received 23 July 2007; received in revised form 18 September 2007; accepted 21 September 2007

Abstract Annotation of the recently determined genome sequence of the major dengue vector, Aedes aegypti, reveals an abundance of detoxification genes. Here, we report the presence of 235 members of the cytochrome P450, glutathione transferase and carboxy/ cholinesterase families in Ae. aegypti. This gene count represents an increase of 58% and 36% compared with the fruitfly, Drosophila melanogaster, and the malaria mosquito, Anopheles gambiae, respectively. The expansion is not uniform within the gene families. Secure orthologs can be found across the insect species for enzymes that have presumed or proven biosynthetic or housekeeping roles. In contrast, subsets of these gene families that are associated with general xenobiotic detoxification, in particular the CYP6, CYP9 and alpha esterase families, have expanded in Ae. aegypti. In order to identify detoxification genes associated with resistance to insecticides we constructed an array containing unique oligonucleotide probes for these genes and compared their expression level in insecticide resistant and susceptible strains. Several candidate genes were identified with the majority belonging to two gene families, the CYP9 P450s and the Epsilon GSTs. This ‘Ae. aegypti Detox Chip’ will facilitate the implementation of insecticide resistance management strategies for arboviral control programmes. r 2007 Elsevier Ltd. All rights reserved. Keywords: Aedes aegypti; P450; Glutathione transferases; Carboxylesterases; Detox chip; Microarray

1. Introduction The mosquito Aedes aegypti is the primary vector of urban arboviruses such as dengue and yellow fever. Twofifths of the world’s population are at risk from dengue, and each year there are an estimated 50–100 million dengue infections, resulting in 20,000–25,000 deaths from dengue haemorrhagic fever (WHO, 2002). Dengue control relies almost entirely on targeting the mosquito vectors by Abbreviations: CCE, carboxy/cholinesterases; P450, cytochrome P450 monooxygenases; GST, glutathione transferases Corresponding author. Tel.: +44 151 7053124; fax: +44 151 7053369. E-mail address: [email protected] (H. Ranson). 0965-1748/$ - see front matter r 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.ibmb.2007.09.007

environmental management, biological control and the use of chemical insecticides as either space sprays or larvicides. Unfortunately many mosquito control programmes are threatened by the emergence of resistance to existing chemicals. Insecticide resistant populations of Ae. aegypti have been detected throughout the geographical distribution of this species (Cui et al., 2006; da-Cunha et al., 2005; Jirakanjanakit et al., 2007; Rodriguez et al., 2001) and in some areas, the evolution of insecticide resistance has been linked to the failure of dengue control programs (Camacho et al., 2004; Coto et al., 2000; Focks et al., 2000). Effective management of insecticide resistance is only possible if the mechanisms underpinning the phenotype are understood. In general, there are two major resistance

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mechanisms: alterations in the target site and increased rates of insecticide metabolism. The latter mechanism is poorly elucidated and it is only in the post-genomic era that the full complexity of the insect detoxification enzyme families is being realised and the genetic basis of metabolic resistance to insecticides is beginning to be unravelled. Three enzyme families, the cytochrome P450 monooxygenases (P450), glutathione transferases (GST) and carboxy/cholinesterases (CCE) are implicated in insecticide metabolism. Each of these catalyse a wide range of detoxification reactions. They are the primary enzymatic defence against xenobiotics, are responsible for the removal of many by-products of metabolism, play essential roles in multiple biosynthetic pathways and are involved in chemical communication (Feyereisen, 2005; Oakeshott et al., 2005; Ranson and Hemingway, 2005). Some individual enzymes also have structural roles, instead of, or in addition to their catalytic activity. This diversity in the functions of each enzyme family is accomplished by a mixture of highly specialised enzymes, often with specific substrates and strictly regulated expression profiles, and more generalist, ubiquitously expressed enzymes. Many insect species show an amazing diversity of detoxification enzymes. As insect genomes have been sequenced, and the detoxification genes annotated, it has become apparent that these gene families are very rapidly evolving and each insect has a unique complement of detoxification genes, with very few orthologs across insect species (Claudianos et al., 2006; Ranson et al., 2002). The rapid expansion and diversification of detoxification genes likely facilitated the adaptation of insects to their particular ecological niches, and, on a more recent evolutionary timescale, has enabled them to survive various man-made xenobiotics, including insecticides. A small subset of the detoxification genes has been previously described in Ae. aegypti (David et al., 2006; Lumjuan et al., 2007; Sieglaff et al., 2005). In this study, we utilised the recently published genome sequence of Ae. aegypti (Nene et al., 2007) to fully characterise these complex gene families and developed a ‘detox chip’ to identify specific genes whose expression is elevated in insecticide resistant populations.

Institute for Genomic Research (TIGR; www.tigr.org/) was used as a reference to confirm our transcript predictions and our transcripts were compared to the automatic annotations in Vectorbase (www.vectorbase. org). The ClustalW algorithm (Thompson et al., 1994) was used to align protein sequences to further support annotation predictions. 2.2. Microarray construction Unique 70mer probes for 204 detoxification genes and six housekeeping genes were designed by the bioinformatics support group at Operon. Where possible the probes were designed towards the 30 end of the gene and any oligonucleotide that had 470% identity to another gene was discarded (an exception to this criterion was made for the 12 P450 genes, denoted v1 and v2 which have two near identical copies in the genome, see supplementary information for more detail). Two independent probes were designed for 35 genes to enable the specificity of the array to be assessed. The two probes were designated by the subscript a or b. 70-mer oligonucleotides were re-suspended in nuclease-free water. Each 70-mer oligo, plus 23 artificial spike-in control genes (Universal Lucidea Scorecard, G.E. Health Care, Bucks, UK) were spotted four times on each array. The array can be accessed at ArrayExpress (http://www.ebi.ac.uk/arrayexpress) (acc. No. A-MEXP-623). 2.3. Mosquito strains

2. Materials and methods

The Ae. aegypti New Orleans (NO), Rockefeller (RKF), Isla Mujeres (IM) and PMD-R strains were reared under standard conditions (28+2 1C, 80% RH) at the Liverpool School of Tropical Medicine. The NO and RKF are standard laboratory strains, susceptible to all insecticides. The PMD-R strain was selected from a parental PMD strain from Ban Pang Mai Dang, in Chiang Mai Province in northern Thailand (Lumjuan et al., 2005). The IM strain was colonised from Isla Mujeres, on the Atlantic coast of Mexico. This strain had been in colony for five generations when used and had been selected with adult exposure to 5 mg permethrin in a bottle bioassay (Brogdon and McAllister, 1998).

2.1. Gene identification and annotation

2.4. Target preparation and microarray hybridisations

To identify CCE, GST, and P450 genes in Ae. aegypti, we conducted a BLAST search of the Ae. aegypti (Liverpool strain) whole genome sequencing database at the Broad Institute (www.broad.mit.edu/annotation/disease_ vector/aedes_aegypti/) with known detoxification genes from Anopheles gambiae and Drosophila melanogaster (Ranson et al., 2002). When significant hits were observed in the genomic sequence, the region was manually annotated to identify the putative transcripts and translation products. The Ae. aegypti EST database at The

RNA extractions, cRNA synthesis and labelling reactions were performed independently for each biological replicate. Total RNA was extracted from batches of 10 1–day-old adult female NO, RKF and PMD-R mosquitoes or 30 IM or NO fourth instar larvae or 2–3 day adults using a PicoPureTM RNA isolation kit (Arcturus) or the RNeasy Midi-kit (Qiagen) according to manufacturer’s instructions. Total RNA quantity and quality were assessed using Nanodrop spectrophotometer (Nanodrop Technologies, UK) before further use. RNA was amplified

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using a RiboAmpTM RN amplification kit (Arcturus) according to the manufacturer’s instructions. Amplified RNAs were checked for quantity and quality by spectrophotometry and agarose gel electrophoresis. Amplified RNA was reverse transcribed into labelled cDNA and hybridised to the array as previously described (David et al., 2005). Each comparison was repeated three times with different biological samples. For each biological replicate, two hybridisations were performed in which the Cy3 and Cy5 labels were swapped between samples so a total of six hybridisations were performed for each comparison. Labelled cDNA from PMD-R or IM were co-hybridised with NO cDNA in pairwise comparisons. 2.5. Microarray data analysis Spots that failed to meet any of the following criteria in either channel were rejected: (1) an intensity value of 4300, (2) signal-to-noise ratio of 43 and (3) greater than 60% of pixel intensity superior to the median of the local background 72 S.D. Normalisation and statistical analyses of the data was performed using the Limma 1.9 software package for R 2.3.1, available from the CRAN repository (http://www.r-project.org). Background corrected intensities from the red, (R, Cy5), and the green, (G, Cy3), channel were transformed to intensity log-ratios, M ¼ log R/G, and their corresponding geometrical means, A ¼ (log R+log G)/2. Within each array M-values were normalised as a function of A using the Lowess (Clevel and Devlin, 1988) scatter plot smoothing function and scaled to equalise the median absolute value across all arrays to account for technical biases between replicate hybridisations. Mean expression ratios were submitted to a one-sample Student’s t-test against the baseline value of 1 (equal gene expression in both samples) with a multiple testing correction (Benjamini and Hochberg false discovery rate). Genes showing both t-test and p-valueso0.001, and X2-fold over or under expression were considered differentially expressed between comparisons. The expression data from these microarray experiments can be accessed at Vectorbase (http://www.vectorbase.org). 3. Results and discussion 3.1. Aedes aegypti detoxification gene families 3.1.1. Cytochrome P450s Ae. aegypti contains a total of 160 full-length, putatively catalytically active P450 genes. This represents an expansion of approximately 52% compared to An. gambiae, 86% compared to D. melanogaster and is 3-fold higher than the number of P450s found in the honeybee, Apis mellifera (Claudianos et al., 2006; Ranson et al., 2002). In addition to the full-length genes, we identified a number of truncated P450 sequences, usually within or flanking P450 gene clusters and frequently surrounded by transposon like sequences. Several large clusters of P450s are

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found in the Ae. aegypti genome, the largest being a cluster of 18 CYP6 genes on supercontig (sc) 1.371 and a cluster of 16 CYP9 genes on sc1.1188 (see Supplementary information for more details on the annotations of the P450 family). As in other Diptera, the majority of the P450s are represented by the CYP3 and CYP4 clades, which contain the insect specific families, CYP4, CYP6, CYP9 and CYP325 and include the majority of enzymes implicated in xenobiotic metabolism and insecticide resistance (Andersen et al., 1994; Daborn et al., 2002; Kasai and Scott, 2001; Maitra et al., 1996; Nikou et al., 2003; Pittendrigh et al., 1997; Tomita and Scott, 1995). Each of these families is expanded in Ae. aegypti compared to An. gambiae and D. melanogaster (Table 1) but this expansion is most pronounced in the CYP9 family (Fig. 1). Thirtyseven CYP9 genes are present in the Ae. aegypti genome compared with just nine in the malaria mosquito and five in the fruitfly. Why the CYP9 family is so abundant in Ae. aegypti is unclear but there are several very recent duplications in this gene family, several of which are over-expressed in one or more insecticide resistant strains

Table 1 Classification of detoxification genes in Drosophila melanogaster, Anopheles gambiae and Aedes aegypti D. melanogaster Glutathione transferases Delta Epsilon Omega Sigma Theta Zeta Others Total Cytochrome P450s CYP4 clade CYP3 clade CYP6 CYP9 Others CYP2 clade Mitochondrial clade Total Carboxy/cholinesterases Alpha esterases Hormone processsing Beta esterases Juvenile hormone esterases Others Glutactin Acetylcholinesterases Total

An. gambiae

Ae. aegypti

11 14 5 1 4 2 0

12 8 1 1 2 1 3

8 8 1 1 4 1 3

37

28

26

32

46

57

22 5 9 7 11

30 9 1 10 9

44 37 1 12 9

86

105

160

13

16

22

3 2

5 4

2 6

3 4 1

4 9 2

7 10 2

26

40

49

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C. Strode et al. / Insect Biochemistry and Molecular Biology 38 (2008) 113–123 CYP9 A E1 CYP9 B 2 CYP9 B 1 93 100 CYP9 H 1 97 CYP9 C1 CYP 9F2 CYP 9 K 1 100 84 CYP9 M 2 CYP9 M 1 100 100 CYP9 M 9 CYP9 M 8 CYP9 M 4 100 CYP9 M 5 CYP9 M 7 CYP9 M 6 100 CYP9 J3 CYP9 J1 5 100 CYP9 J27_ V 1 100 CYP9 J27_ V 2 100 CYP9 J2 6 99 100 CYP9 J3 0 CYP9 J2 9 100 CYP9 J3 1 CYP9 J6 CYP9 J5 83 CYP9 J3 2 100 100 CYP9 J22_ V 2 93 CYP9 J22_ V 1 CYP9 J2 3 100 CYP9 J2 8 CYP9 J2 4 98 CYP9 J7 CYP9 L 1 100 100 CYP9 L 3 79 CYP9 L 2 CYP9 J4 100 CYP919_ V 2 CYP9 J19_ V 1 CYP9 J2 92 100 CYP9 J18_ V 2 CYP9 J18_ V 1 100 CYP9 J1 7 CYP9 J1 6 100 CYP9 J10_ V 2 100 CYP9 J10_ V 1 100 CYP9 J9 _V 2 CYP9 J9 _V 1 100 CYP9 J8 _V 1 100 CYP9 J8 _V 2 100 CYP9 J20_ V 2 CYP9 J20_ V 1 100

*

*

* *

primary role in metabolism of endogenous compounds rather than xenobiotics (Maibeche-Coisne et al., 2000). However, another member of the CYP4G family, CYP4G20 is absent from brains but expressed at very high levels in the olfactory apparatus of the cabbage armyworm Mamestra brassicae and is suggested to be important in odorant metabolism (Maibeche-Coisne et al., 2005). In contrast, the mitochondrial and CYP2 clades are fairly well conserved across the Diptera with only a moderate expansion in Ae. aegypti (Table 1) and close sequence similarities between species (Fig. 2). These two clades include the D. melanogaster Halloween genes involved in ecdysteroid biosynthesis, all of which have 1:1:1 orthologies within the Diptera and also in the honeybee, and the juvenile hormone epoxidase, CYP15B1.

*

* * * *

*

* *

* *

* * * *

Fig. 1. Unrooted distance neighbour joining tree showing phylogeny of Dipteran CYP9 P450s. Sequences in green are from Ae. aegypti, blue sequences are from An. gambiae and red are from D. melanogaster. Sequences were aligned using ClustalW. Nodes with 470% bootstrap support (500 pseudoreplicates) are indicated. P450s were named by the P450 nomenclature committee (http://drnelson.utmem.edu/CytochromeP450. html). Genes with v1 and v2 designation are very recent duplications and, have not yet been assigned individual gene names. Genes showing a significant differential expression in the microarray experiments (Fig. 4) are marked with an asterisk.

and hence these recent expansions may partially reflect an adaptation to insecticide exposure. Just two of the 160 Ae. aegypti genes in the CYP3 and CYP4 clades have orthologs in the other two Dipteran species. CYP4G35 is the ortholog of D. melanogaster CYP4G1 and An. gambiae CYP4G17, and CYP4G36 is the ortholog of D. melanogaster CYP4G15 and An. gambiae CYP4G16. Together, these six P450s form a monophyletic cluster. The function of these CYP4s is unknown but these genes are amongst the most highly expressed of the cytochrome P450 family (Maibeche-Coisne et al., 2005, 2000; and this study) and one, CYP4G15 has been localised to the insect’s nervous system, suggesting that they have a

3.1.2. Carboxy/choline esterases The CCE family is characterised by the presence of the a/b hydrolase fold and includes both catalytically active enzymes and non-catalytic proteins which are thought to be primarily involved in neurone signalling and development (Oakeshott et al., 2005). In the current discussion, we focus on the catalytic enzymes but we have included the glutactin clade in our gene-count. Although the only characterised insect member of the glutactin clade is catalytically inactive and thought to be involved in cell adhesion in the nervous system (Oakeshott et al., 2005), other members of this clade are putatively catalytically active, as predicted by the presence of a conserved catalytic triad. Ten glutactins are present in the Ae. aegypti genome (Table 1), and all but two of these are predicted to be enzymatically active, although their substrates are unknown. We have identified 49 CCE genes in the Ae. aegypti genome. The vast majority of these are full length, although for five genes the start methionine could not be unambiguously determined. In addition, we identified nine partial sequences with high similarities to CCEs. Seven of these are on very short contigs and are probably a result of assembly errors but two (CCEae2Ap and CCEaejhe5Fp on supercontigs 1.142 and 1.145, respectively) are probably CCE pseudogenes. The classification system described in (Oakeshott et al., 2005) was used to designate the clades in the CCE phylogeny and this is partially reproduced in Fig. 3. Each clade was well supported, although clade E, representing the b esterases, was split into two monophyletic groups. The number of catalytically active enzymes in Ae. aegypti exceeds that found in An. gambiae or D. melanogaster (Table 1). Most of this expansion occurs in the a-esterase clade, clade B. There are three clusters of five or six enzymes, which share 38–91% amino acid identity within the cluster. The high level of identity between several of the pairwise comparisons, together with the lack of clear orthologies between Anopheles and Aedes a-esterases (with two exceptions) suggest that this is a rapidly evolving

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100

CYP305A4 CYP305A3 CYP305A2 100 CYP305A5 100 CYP305A6 CYP305A1 76 100 CYP15B1 CYP15B1 CYP303A1 100 86 100 CYP303A1 CYP303A1 CYP304C1 100 CYP304C1 100 CYP304A1 81 CYP304B3 100 CYP304B1 99 CYP304B2 CYP18A 1 100 CYP18A1 100 CYP306A1 100 phn CYP306A 1 100 CYP306A 1 100 CYP307A1 100 CYP307A1 CYP307A1 100 CYP307B1 100 CYP307B1_V1 100 CYP307B1_V2 CYP315A1 100 CYP315A1 99 sad CYP315A1 CYP314A1 100 100 CYP314A1 shd CYP314A1 CYP302A1 100 dib 100 CYP302A1v1 CYP302A1 CYP301A1 100 100 CYP301A1 CYP301A1 100 CYP49A 1 100 CYP49A 1 100 CYP49A 1 CYP12A 5 100 100 89 CYP12A 4 98 CYP12C1 CYP12E1 CYP12D 2 100 100 CYP12D 1 CYP12B2 CYP12F6 100 CYP12F5 100 CYP12F8 100 100 CYP12F7 CYP12F4 CYP12F1 89 CYP12F3 CYP12F2 70 100

77

77

78

CYP2 clan

spo

mitochondrial clan

Fig. 2. Unrooted distance neighbour joining tree showing phylogeny of Dipteran cytochrome P450s from the mitochondrial clade and the CYP2 clade. Sequences in green are from Ae. aegypti, blue sequences are from An. gambiae and red are from D. melanogaster. Sequences were aligned using ClustalW. Nodes with 470% bootstrap support (500 pseudoreplicates) are indicated. 1:1:1 orthologies among the three Dipteran species are designated with a dotted line, and the Halloween genes (phn, spo, sad, sho, dib, spok and spookiest) involved in ecdysteroid biosynthesis are labelled. Note that two identical copies of CYP307B1 are found in the current assembly of the Ae. aegypti genome.

enzyme group. Alpha esterases in other species are involved in metabolic resistance to insecticides. These include the Lucilia cuprina a E7 (Newcomb et al., 1997) and the Culex quinquefasciatus ‘a’ and ‘b’ genes (named according to substrate specificity as opposed to phylogenetic relationships) (reviewed in Hemingway and Karunaratne, 1998). These two Culex genes are arranged in a head-to-head orientation and amplification of one or both of these genes

is a major cause of organophosphate (OP) resistance in Culex populations worldwide (Mouches et al., 1986; Vaughan et al., 1997). Interestingly, in contrast to the rapid radiation of other a-esterases, these two genes are well conserved across Culex, Anopheles and Aedes (CCEae1D and 2D; Fig. 3) and the head-to-head orientation is maintained in all species. It remains to be seen whether OP resistance in Ae. aegypti, reported by Mourya

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CCEa e2F CCEa e2D CCEa e1F 100 100 CCEa e1D CG1082 85 CG1108 CG1257 74 CG1131 95 CG1128 86 CG1121 CG1112 CG1089 CG2505 10 0 CG1031 83 75 CG6018 99 CCEa e3D CCEa e2D CCEa e3B CCEa e1B CCEa e5B 71 CCEa e6B CCEa e4B 99 CCEa e2B CCEa e1D 100 CCEae7 G 93 CCEae5 G 100 CCEae6 G CCEa e6A CCEa e1o 100 CCEae2 G 93 100 CCEae1 G CCEa e3o CCEae3 G 99 91 CCEa e5A 94 CCEa e3A CCEa e1A 99 CCEae4 G CCEa e4A CG9858 100 CCEa e5o 100 CCEa e2o CG10175 CCEa e6o 100 99 CCEa e2A CCEae6 C 71 CCEa e1A 100 99 CCEae5 C CCEae4 C 98 CCEae1 C 100 CCEae3 C CCEae2 C 100 CCE15o CCEunk4o 100 CCEunk6o 100 CCEunk5o 98 CCEunk3 o 81 CCEunk2o 70 CCEunk1o CCE13o 97 100 CCE14o CCEunkn 100 CCEbe2o 100 CCEbe1o CG6414 CG4382 100 CCEunk7o 100 CG4757 CG3841 100 CCEbe4C CCEbe3 C CCEbe2C 100 75 CCEbe1C 100 CCEbe2o 100 CG17148 CG6917 96 CG8424 CG8425 100 CCEj he1E 87 CCEj hei E 100 he4E CCEj 86 CCEj he3E CCEj he5E 89 CCEj he2E CCEj he1F CCEj he3F 100 CCEj he4F 100 100 CCEj he1o CCEj he2F 100

84

100

C

B

G

E D

E

F

Fig. 3. Unrooted distance neighbour joining tree showing phylogeny of catalytic classes of Dipteran carboxyl/cholinesterases. Sequences in green are from Ae. aegypti, blue sequences are from An. gambiae and red are from D. melanogaster. Sequences were aligned using ClustalW. Nodes with 470% bootstrap support (500 pseudoreplicates) are indicated. Glutactins and acetylcholinesterases were omitted for clarity. Clades were designated according to the nomenclature adopted by Oakeshott et al., 2005.

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et al. (1993), is associated with amplification of this genomic locus. The other clade of esterases associated with OP resistance is the actetylcholinesterases (ace). These are the target site of both OP and carbamate insecticides and the toxins act as irreversible inhibitors of the enzyme, blocking hydrolysis of the neurotransmitter acetylcholine. Mutations in ace can reduce the binding of insecticides resulting in resistance. As in Anopheles, Ae. aegypti contains two ace genes. Mutations in ace-1 have been associated with resistance to insecticides in other mosquitoes (Weill et al., 2004) but no such mutations have been reported in Ae. aegypti to date. One esterase lineage, the b-esterases (clade E in Fig. 3), is greatly pared down in Aedes compared to Anopheles. This clade includes the Drosophila Est-6 and 7 enzymes, which are important in reproductive physiology (Meikle et al., 1990; Saad et al., 1994). Six juvenile hormone esterase (JHE) genes are present in Ae. aegypti, two of which, CCEjhe1F and 4F, contain the GQSAG motif around the catalytic site, typical of JHEs (Oakeshott et al., 2005) although neither of these has clear orthology with the D. melanogaster JHE (CG8425). Finally, a clade of unknown function but phylogenetically related to lepidopteran JHEs, clade G is well represented in mosquitoes (Fig. 3). 3.1.3. Glutathione transferases In contrast to the other two enzyme families, there has been no expansion of the GST family in Ae. aegypti. Indeed mosquitoes have considerably fewer GST genes than Drosophila although this deficit is partially rectified by alternative splicing of two mosquito GST genes (Ding et al., 2003; Lumjuan et al., 2007) which increases the number of Ae. aegypti GST transcripts by 3 to 29. Each of the GST classes found in An. gambiae is represented in Ae. aegypti, including the two classes (Xi and Iota in Figure S1) which have so far been found uniquely in mosquitoes. Over half of the GSTs belong to two insect specific classes, the Delta and Epsilon classes, which include the vast majority of the GST enzymes with a defined role in insecticide metabolism (Ranson et al., 2001). The Ae. aegypti GST family has recently been described in detail (Lumjuan et al., 2007) and only the major findings are summarised below. As with the CCEs and P450s very few 1:1:1 orthologies can be found within the Dipteran GSTs (Figure S1). Indeed, within the insect specific GST classes, only the Delta GST, GSTD7 and the Epsilon GST, GSTE8 have clear 1:1:1 orthology across the three species. An ortholog of GSTD7 has also been identified in Bombyx mori (accession no. AJ006502) and Manduca sexta (Rogers et al., 1999). It is possible that these two conserved genes represent the ancestral members of the Delta and Epsilon classes from which other members have evolved through a process of local gene duplication and diversification. The expansion of these two classes presumably occurred after

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the split between the nematocera and brachycera suborders as both the gene structure and arrangement of the Epsilon and Delta GSTs differ between the mosquito and fruitflies. Whether the radiation of these two GST classes occurred before or after the split between the culicine and anopheline subfamilies is difficult to predict with only two full mosquito genome sequences available at present. There are several probable orthologs within the Delta and Epsilon GST classes of An. gambiae and Ae. aegypti but also other examples of apparent independent radiation (Figure S1). The intron positions within the orthologs are conserved and there is also conservation of alternative splicing of the Sigma GST, GSTS1 and the Delta GSTD1 between An. gambiae and Ae. aegypti although the GSTD1 gene produces four distinct transcripts in Anopheles as opposed to the three putative transcripts of the Ae. aegypti GSTD1 gene. Why Ae. aegypti has such an abundance of detoxification genes compared to other insect species is currently mysterious. There are aspects of a mosquito’s life history, such as host seeking, haematophagy and pathogen transmission, that distinguish them from other insect species with fully characterised detoxification gene families (D. melanogaster and A. mellifera) and detoxification enzymes may play important roles in each of these behaviours. But these can neither account for the approximately third higher gene count in Ae. aegypti than An. gambiae nor can this difference be readily accounted for by differential exposure to xenobiotics. Both species have a preference for breeding in clean water (as opposed to Culex mosquitoes which readily breed in water heavily contaminated with organic material), both are highly anthropophilic and hence exposed to man-made pollutants and both are frequently targeted with insecticides. Perhaps the more generalist ecological niche occupied by Ae. aegypti has exerted a greater distribution of selective pressure, resulting in an increased repertoire of detoxification enzymes in this species? The geographic range of Ae. aegypti is also much wider than that of An. gambiae, which is restricted to sub-Saharan Africa, and the dengue vector may have historically encountered a wider range of toxins, food sources, environmental and ecological stresses. It will be necessary to examine the distributions and numbers of detoxification genes in the tribe Culicini (basal to the subfamily Culicinae) and in the basal members of Aedini (e.g. genus Psorophora) before it can be determined whether expansion of detoxification gene families occurred early in the evolution of Culicinae or is more recently derived. 3.2. Identifying detoxification genes associated with insecticide resistance As a consequence of the rapid expansion of the detoxification supergene families, which has likely resulted in a degree of genetic redundancy, and the exceptionally high rate of diversification, which has left very few secure

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orthologs even within the mosquito family, comparative genomics can provide few clues as to the genes responsible for insecticide resistance. We therefore constructed a smallscale microarray, the ‘Aedes Detox Chip’, containing unique 70mer probes for 204 of the 235 Ae. aegypti detoxification genes and used this to compare expression profiles in susceptible and resistant populations. We selected two populations of Ae. aegypti each colonised from sites where pyrethroid resistance had been detected in the field, and then further selected with permethrin in the laboratory, and compared the transcript levels in these strains to the laboratory susceptible NO strain. The PMD-R strain originated from Chiang Mai

district in Northern Thailand and the IM strain is from Isla Mujeres, off the Atlantic coast, near Cancun Mexico (see Section 2 for further strain details). Ideally, the comparisons would have been performed between a susceptible and resistant population selected from the same panmictic population. However, resistance to insecticides is now so widespread in Ae. aegypti populations that it is very difficult, if not impossible, to find a fully insecticide susceptible population in the field. We therefore conducted a series of preliminary microarray experiments comparing the NO strain with a second laboratory susceptible strain (RKF) with a different geographical origin (Rodriguez et al., 2001). Using cut-off values42-fold change in

25

25

20

20

AaGSTe2 CYP9J27

CCEbe2 o CYP9J 8

CYP9J2 8 AaGSTe3 CYP12F6

AaGSTs1-2

CYP9J2 6 CYP9J10

15

CYP6AG4

CYP9J27

CYP9J 8

CYP304C1 CYP9J32 CYP6Z9

CYP6Z6_b

10

-log10 (pvalue)

-log10 (pvalue)

CYP6CB 1 CYP6AG3

CYP329B 1

AaGSTe2

15

CYP9J3 2 CYP9J 10

CYP9J 24

CYP6Z9

AaGSTe4 CYP9J1 9

10

AaGSTe7

5

0 -20

5

CYP9M6

-16

-12

-8 -4 0 4 8 Fold change (PMDR/NO)

upregulated NO

12

16

20

0 -20

-16

-12

-8

-4

0

4

8

12

16

20

Fold change (IM/NO)

upregulated PMDR

upregulated NO

upregulated IM

25

-log10 (pvalue)

20

15 CYP9J32

10

AaGSTe3 CYP9J28 AaGSTe2

5

AaGSTe6 CYP18A 1

0 -20

-16

-12

CYP4J15

-8 -4 0 4 8 Fold change (IM/NO Larvae)

upregulated NO

12

16

20

upregulated IM

Fig. 4. Differential expression of Ae. aegypti detoxification genes in insecticide resistant and susceptible strains. Differences are indicated as a function of both expression ratio (X-axis) and significance, expressed as the negative log10 scale of the p-value of the t-test of the fold change between the groups (Y-axis). Vertical lines indicate two-fold expression differences in either direction. The horizontal line indicates the significance threshold of po0.001 adopted for the one sample t-test. CYP9 P450s are shown in green, Epsilon GSTs in red. Selected genes are named. The entire data set is available at ArrayExpress (accession no. A-MEXP-623). Panel (A) shows constitutive expression levels in adults from the PMD-R strain compared to the susceptible NO strain, panel (B) shows the adult IM versus NO comparison and panel (C) shows the results from the IM versus NO larval comparison.

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expression and a p-valueo0.001, expression of the detoxification genes was indistinguishable between the populations (Figure S2) In contrast, using these same cut-off values, 25 genes were over-expressed in the PMD-R relative to NO and 14 over-expressed in IM versus NO (Fig. 4 and Table S3). Five of these genes, three CYP9 P450s and two Epsilon GSTs, were over expressed in both resistant strains. For the IM strain, we also compared the basal gene expression in fourth instar larvae with age-matched larvae from the NO strain. The resistance phenotype is manifested in both larval and adult stages but previous studies had suggested that the genes responsible may differ between different developmental stages (Nikou et al., 2003). In the present study, we observed a similar profile of differential gene expression between the IM and NO strains in both life stages, although the magnitude of the expression ratios was generally lower in the larval comparisons and four of the CYP9 P450s over expressed in IM adults showed no differential expression in larvae (Fig. 4 and Table S3). The most striking result from this microarray data is the over-representation of two gene families in the subset of genes expressed at higher levels in resistant populations. Of the 33 genes showing constitutively higher expression in one or both of the resistant strains 18 (54%) belong to either the CYP9 family of P450s or the Epsilon GST class. If the threshold is raised to include only genes showing 43-fold change in expression, the percentage of the upregulated genes belonging to these two families increases to 67%. As a percentage of the genes represented on the chip the CYP9 P450s and Epsilon GSTs represent less than 20%, so even considering the massive expansion that has occurred in the CYP9 family in Ae. aegypti, these two groups of enzymes are disproportionately represented amongst the candidate set of genes associated with insecticide resistance. Further experimental validation including enzyme characterisation is necessary to conclusively incriminate these candidate genes in insecticide metabolism and hence in conferring resistance in these strains. 4. Conclusions Our preliminary microarray analysis of the detoxification genes in Ae. aegypti has identified two enzyme families as candidates for conferring insecticide resistance in this species. CYP9 P450s have been implicated in pyrethroid resistance in other species (Pittendrigh et al., 1997) and elevated expression of Epsilon GSTs have been shown to cause DDT and OP resistance in mosquitoes and houseflies (Motoyama and Dauterman, 1974; Ranson et al., 2001) although the role of GSTs in pyrethroid metabolism is less clear and clearly warrants further investigation. Further experiments with additional resistant populations will help refine this candidate gene list. Moreover, the ability to store Aedes eggs in a desiccated state greatly facilitates the maintenance of multiple lines and will enable microarray experiments to be conducted on progeny collected directly

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from the field thereby eliminating any confounding effects of colonisation. Thus, the sequencing of the Ae. aegypti genome, and the development of a robust microarray platform for detoxification genes, will greatly accelerate research into the genetics of insecticide resistance. By utilising this knowledge to develop molecular markers for resistance alleles, we will be able to survey any given population to predict its resistance profile and identify suitable classes of insecticide for control. This will greatly facilitate the monitoring and management of insecticidebased Aedes control programmes and will enhance our ability to control this major disease vector in future generations. Acknowledgements We are grateful to the Aedes aegypti genome community for their efforts in determining the genome sequence and to the staff of VectorBase, in particular Dan Lawson and Bob MacCallum. We also thank Drs. Pradya Samboon and La-aied Prapanthadara for the initial colonisation and selection of the PMD-R Aedes aegypti strain and we wish to acknowledge the technical support of Dr. Margaret Hughes in the printing of the microarray slides. Funding was provided by the Innovative Vector Control Consortium and the Wellcome Trust. Appendix A. Supplementary Materials Supplementary data associated with this article can be found in the online version at doi:10.1016/j.ibmb. 2007.09.007. References Andersen, J.F., Utermohlen, J.G., Feyereisen, R., 1994. Expression of house fly CYP6A1 and NADPH-cytochrome P450 reductase in Escherichia coli and reconstitution of an insecticide-metabolizing P450 system. Biochemistry 33, 2171–2177. Brogdon, W.G., McAllister, J.C., 1998. Simplification of adult mosquito bioassays through use of time-mortality determinations in glass bottles. J. Am. Mosq. Control Assoc. 14, 159–164. Camacho, T., de la Hoz, F., Cardenas, V., Sanchez, C., de Calderon, L., Perez, L., Bermudez, A., 2004. Incomplete surveillance of a dengue-2 epidemic in Ibague, Colombia, 1995–1997. Biomedica 24, 174–182. Claudianos, C., Ranson, H., Johnson, R.M., Biswas, S., Schuler, M.A., Berenbaum, M.R., Feyereisen, R., Oakeshott, J.G., 2006. A deficit of detoxification enzymes: pesticide sensitivity and environmental response in the honeybee. Insect Mol. Biol. 15, 615–636. Clevel, W.S., Devlin, S.J., 1988. Locally weighted regression: an approach to regression analysis by local fitting. J. Am. Stat. Assoc. 83, 596–610. Coto, M.M., Lazcano, J.A., de Fernandez, D.M., Soca, A., 2000. Malathion resistance in Aedes aegypti and Culex quinquefasciatus after its use in Aedes aegypti control programs. J. Am. Mosq. Control Assoc. 16, 324–330. Cui, F., Raymond, M., Qiao, C.L., 2006. Insecticide resistance in vector mosquitoes in China. Pest Manag. Sci. 62, 1013–1022. da-Cunha, M.P., Lima, J.B., Brogdon, W.G., Moya, G.E., Valle, D., 2005. Monitoring of resistance to the pyrethroid cypermethrin in Brazilian Aedes aegypti (Diptera: Culicidae) populations collected between 2001 and 2003. Mem. Inst. Oswaldo Cruz 100, 441–444.

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