Identification and characterization of eleven glutathione S-transferase genes from the aquatic midge Chironomus tentans (Diptera: Chironomidae)

Insect Biochemistry and Molecular Biology 39 (2009) 745–754

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Identification and characterization of eleven glutathione S-transferase genes from the aquatic midge Chironomus tentans (Diptera: Chironomidae) Xiuwei Li a, b, Xin Zhang b, Jianzhen Zhang b, Xing Zhang a, Sharon R. Starkey b, Kun Yan Zhu b, * a b

R&D Center of Biorational Pesticides, Northwest A & F University, Yangling, Shaanxi 712100, China Department of Entomology, 123 Waters Hall, Kansas State University, Manhattan, KS 66506, USA

a r t i c l e i n f o

a b s t r a c t

Article history: Received 23 March 2009 Received in revised form 25 August 2009 Accepted 28 August 2009

Eleven cDNAs encoding glutathione S-transferases (GSTs) were sequenced and characterized in Chironomus tentans, an ecologically important aquatic midge. Phylogenetic analysis revealed seven GSTs in three different cytosolic classes including 4 in sigma (CtGSTs1, CtGSTs2, CtGSTs3, CtGSTs4), 2 in delta (CtGSTd1, CtGSTd2), and 1 in omega (CtGSTo1). The remaining four GSTs (CtGSTu1, CtGSTu2, CtGSTu3, CtGSTu4) were unclassified due to their low relatedness to currently known classes of insect GSTs. Reverse-transcription (RT)-PCR analysis of the 11 GST genes showed that CtGSTd1, CtGSTu2, CtGSTu4, CtGSTs1, CtGSTs2, CtGSTs3, CtGSTs4 and CtGSTo1 were expressed in all tissues examined, including salivary glands, hemolymph, midgut, Malpighian tubules, fatbodies and carcass, whereas CtGSTd2 and CtGSTu1 were expressed in a limited number of tissues. CtGSTs1 and CtGSTs4 appeared to be the only two genes, of which expressions can be detected in eggs, whereas all the 11 GST genes showed various expression patterns in the four larval instars. However, expressions of CtGSTd2, CtGSTu1 and CtGSTu2 were not detectable in pupal and adult stages. Real-time quantitative PCR confirmed that the herbicide alachlor increased CtGSTd1, CtGSTs2 and CtGSTs3 gene expression by 2.1-, 2.8- and 4.3-fold, respectively, when fourth-instar midges were exposed to alachlor at 1000 mg/L for 72 h. Such increased gene expressions were associated with 2.2- and 1.8-fold decreases of total GST activities in vivo when CDNB and DCNB were used as substrates, respectively. Further studies showed that 65.5 and 73.5% of GST activities were inhibited in vitro by alachlor at 100 and 1000 mg/L, respectively. Because alachlor has been known as an electrophilic substrate that can be conjugated by glutathione (GSH), rapid in vitro inhibition of GST activities by alachlor suggested that decreased GST activities were likely caused by the depletion of GSH. However, alachlor may regulate different GST genes, as found in other organisms, leading to significantly increased transcriptional levels of CtGSTd1, CtGSTs2 and CtGSTs3 in out of 11 GST genes examined in this study. Ó 2009 Elsevier Ltd. All rights reserved.

Keywords: Chironomus tentans Glutathione S-transferase Alachlor Gene expression Herbicide

1. Introduction Glutathione S-transferases (GSTs, EC 2.5.1.18) belong to a diverse family of dimeric enzymes that play important roles in phase II detoxification of both xenobiotics (drugs, insecticides, herbicides) and endogenous compounds in almost all living organisms (Ding et al., 2003; Enayati et al., 2005; Tu and Akgu¨l, 2005). These enzymes catalyze the conjugation reactions of nucleophilic sulfhydryl of the reduced form of glutathione (GSH), a tripeptide of g-glutamyl-cysteinyl-glycine, to electrophilic centers of lipophilic compounds via a nucleophilic substitution/addition reaction. Such catalytic reactions commonly result in water-soluble, less toxic substrates that can be rapidly excreted from the organism’s body

* Corresponding author. Tel.: þ1 785 532 4721; fax: þ1 785 532 6232. E-mail address: [email protected] (K.Y. Zhu). 0965-1748/$ – see front matter Ó 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.ibmb.2009.08.010

(Grant and Matsumura, 1989; Hayes et al., 2005). GSTs can also bind to hydrophobic compounds that are not their substrates. Such binding is possibly associated with the sequestration, storage and transportation of drugs, hormones, and other metabolites (Hayes and Pulford, 1995; Lumjuan et al., 2007). Many studies have also demonstrated that GSTs play an important role in insecticide resistance by detoxifying insecticides (Chen et al., 2003; Enayati et al., 2005; Kristensen, 2005; Penilla et al., 2006; Yang et al., 2009; Che-Mendoza et al., 2009). GSTs can be divided into three major groups based on their distributions within the cell, which include cytosolic, microsomal and mitochondrial GSTs (Jakobsson et al., 1996; Sheehan et al., 2001; Robinson et al., 2004). Mammalian cytosolic GSTs have been classified to eight classes including alpha, kappa, mu, omega, pi, sigma, theta and zeta (Hayes and McLellan, 1999; Board et al., 2000; Sheehan et al., 2001; Pearson, 2005). In insects, mitochondrial GSTs have not been found to date, and most studies have focused on

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cytosolic GSTs. By extending the mammalian GST classification system to encompass insect GSTs, insect cytosolic GSTs have been assigned to six classes including delta, epsilon, omega, sigma, theta and zeta (Chelvanayagam et al., 2001; Ranson et al., 2001, 2002; Ding et al., 2003; Ranson and Hemingway, 2005; Tu and Akgu¨l, 2005; Yu et al., 2008). Among the six classes of insect GSTs, delta and epsilon are two predominant classes of GSTs in both Anopheles gambiae and Drosophila melanogaster (Enayati et al., 2005), and in Bombyx mori (Yu et al., 2008), all of which have had their genomes sequenced. The aquatic midge (Chironomus tentans) is a well recognized and widely used insect species for studying the impact of environmental pollutants in aquatic systems due to its significant role in food webs and occurrence in various aquatic habitats (US EPA, 1993; Anderson, 2006). Numerous studies have shown that although many triazine (e.g., atrazine) and chloroacetanilide (e.g., alachlor) herbicides at relatively high concentrations (e.g., 1000 mg/L or ppb) do not exhibit significant acute toxicity to the midge, they can modify metabolic pathways of insecticides (Belden and Lydy, 2000; Miota et al., 2000; ˜o Anderson and Lydy, 2001; Jin-Clark et al., 2002, 2008; London et al., 2004, 2007; Anderson and Zhu, 2004; Anderson et al., 2008). For example, alachlor can reduce GST activity, leading to increased susceptibility of the midge to organophosphates (Jin-Clark et al., 2008). However, very little has been known about the molecular mechanism of reduced GST activity in response to alachlor exposure in the midge. In fact, it is virtually unknown about the GST genes, their molecular characteristics, and developmental and tissuespecific expression patterns in the midge. To comprehensively characterize midge’s cellular and molecular responses to environmental pesticides and other toxic stressors, we have developed an expressed sequence tag (EST) database containing over 10,000 sequences from a C. tentans cDNA library. From the EST database, we have identified a total of 11 GST genes and carried out a relatively detailed analysis of these genes. Herein, we report: 1) molecular analysis of the 11 complete cDNAs encoding GSTs in the midge; 2) developmental stage- and tissue-specific expression patterns of the 11 GST genes; and 3) effect of alachlor on total GST activities and gene expressions. Our study provides the first insight into molecular characteristics of GSTs and their transcriptional response to alachlor exposures in C. tentans. 2. Materials and methods 2.1. Insect The aquatic midge (C. tentans) colonies were cultured in Insect Toxicology Laboratory at Kansas State University according to the standard procedures of U.S. Environmental Protection Agency (US EPA, 1993) with slight modifications. Specifically, the cultures were maintained with a mixture of all developmental stages (Jin-Clark et al., 2002; Anderson and Zhu, 2004; Rakotondravelo et al., 2006). 2.2. cDNA library construction, EST sequencing and analysis A midge cDNA library was constructed by American Gene C.T., LLC (Cranston, RI) using the mRNA purified from whole bodies of mixed second- to fourth-instar larvae of the midge, and lambda Uni-ZAPÒ XR vector and Gigapack III gold cloning kits according to the manufacturer’s instructions (Stratagene, La Jolla, CA). Recombinant plasmid within the lambda Uni-ZAPÒ XR vector was in vivo excised and recirculated to generate subclones in the pBluescript SK phagemid vector for DNA sequencing. The sequencing work was performed by the Roy J. Carver Biotechnology Center of the University of Illinois at UrbanaChampaign. Briefly, 10,368 cDNA clones (108 96-well plates) were

robotically picked up from the plated plasmid library by the QPix robot (Genetix, Ltd., Hampshire, UK) and transferred into 96-well plates. The plasmids containing cDNA inserts from midges were prepared by heat lysis and sequenced by single-pass using the ABI3730x1 capillary sequencer (Applied Biosystems, Foster City, CA). 2.3. Identification and analysis of GST cDNAs DNA sequences were processed, clustered and assembled using the online program EGassembler (Masoudi-Nejad et al., 2006). Blast search, mapping and annotation were performed using the online program Blast2GO (http://www.blast2go.de). Each EST putatively encoding a GST retrieved from Blast2GO was further searched using BLASTX against the non-redundant database at NCBI (http://www.ncbi.nlm.nih.gov/) to confirm its identity as a GST gene. If a particular EST did not have a complete open reading frame (ORF) for its GST, additional sequencing was performed for the same cDNA template using the same or new sequencing primers. The sequence accuracy of each GST cDNA was verified by sequencing the same cDNA template from both 30 and 50 directions. 2.4. Phylogenetic analysis The amino acid sequences of the GSTs were deduced from their cDNAs and aligned using ClustalW (Thompson et al., 1994). Alignments were converted to meg files using MEGA software (http://www.megasoftware.net/m_computing.html). The phylogenetic tree was constructed by the neighbour-joining method (Saitou and Nei, 1987) with TREECON by using MEGA2 (van de Peer and De Wachter, 1994). 2.5. Reverse-transcription PCR (RT-PCR) analysis For stage-specific gene expression studies, total RNA was isolated from the whole bodies of each of seven different developmental stages, including egg; first-, second-, third- and fourthinstar larva; pupa; and adult, using TRIzol Total RNA Isolation kit (Invitrogen). For tissue-specific gene expression studies, total RNA was isolated from each of six different tissue samples, including salivary glands, hemolymph, midgut, Malpighian tubules, fatbodies and carcass (i.e., the insect body after its digestive canal is removed), dissected from fourth-instar larvae (Zhu, 2009). Total RNA was then treated with DNase using DNase I kit (Fermentas, Glen Burnie, MD) and cDNA was synthesized using the First Strand cDNA Synthesis kit (Fermentas). Beacon Designer software from Primer Biosoft (http://www.premierbiosoft.com) was used to design all the PCR primers (Supplementary material). RT-PCR was performed using the thermal cycle program consisting of an initial denaturation at 94  C for 2 min followed by 28 cycles of 94  C for 30 s, 55  C for 30 s and 72  C for 45 s, and a final extension at 72  C for 10 min. The midge ribosomal protein S3 (CtRPS3) transcript was used as a reference for RT-PCR analysis. RT-PCR was repeated at least three times, each with a new preparation of total RNA, for each GST gene for each developmental stage and each tissue type. 2.6. Real-time quantitative PCR (qPCR) analysis The exposures of the midges to different concentrations of alachlor were carried out by adding 100 ml of the herbicide solution prepared in acetone to 1 L of reconstituted water containing 15 fourth-instar larvae and 10 ml of fine silica sand as a substrate to avoid cannibalism (Jin-Clark et al., 2008). The final concentrations of alachlor were 1, 10, 100, and 1000 mg/L. The same procedure was used to treat the midges with a corresponding concentration of acetone (100 mg/L) in water as a control. The treated midges were

X. Li et al. / Insect Biochemistry and Molecular Biology 39 (2009) 745–754

maintained in a growth chamber at 25  1  C with a 16:8-h light:dark photoperiod for 72 h. The mortality of midges under such conditions was typically <10% (Jin-Clark et al., 2008). All surviving midges from each replication of a treatment were collected for total RNA isolation. The qPCR was performed using iCycler iQ real-time PCR detection system (Bio-Rad, Hercules, CA) and Maxima SYBR Green qPCR Master Mix kit (Fermentas). A melting curve was established for each sample by using 80 cycles consisting of 95  C for 1 min, 55  C for 1 min and 55  C for 10 s. The relative expression of each GST gene was determined using cycling parameters of an initial denaturation at 95  C for 5 min followed by 40 cycles of 95  C for 15 s, 55  C for 30 s and 70  C for 30 s. The threshold cycle (Ct) value for each dilution was then plotted against the log of its concentration, and Ct values for the experimental samples were plotted onto this dilution series standard curve. Target quantities were calculated from separate standard curves generated for each experiment. Relative expression values (REVs) were then determined by dividing the quantities of the target sequence of interest with the quantity obtained for CtPRS3 as an internal reference gene. The qPCR was repeated four times (n ¼ 4) for each gene. Each replication was performed based on an independent RNA sample preparation and consisted of two technical replications. 2.7. Assay of GST activity and in vitro inhibition The GST activity in fourth-instar larvae of the midge was determined according to the method of Zhu et al. (2000) as modified by Rakotondravelo et al. (2006) using CDNB and DCNB as substrates. The conjugation of GSH to CDNB or DCNB was determined by recording the change in absorbance at 340 nm for CDNB and 344 nm for DCNB for 1 min with 10-s intervals using Ultraspec 3000 UV/visible spectrophotometer (Pharmacia Biotech, Ltd., Cambridge, UK). Nonenzymatic controls were performed in parallel in order to correct for nonenzymatic conjugation of GSH to the substrates. For the assay of in vitro inhibition of GST activities, 10 ml of appropriately diluted alachlor or diethyl maleate (DEM, Sigma– Aldrich, St. Louis, MO) (1 mg/ml in acetone) in 0.1 M phosphate buffer (pH 7.5) were mixed with 10 ml of the enzyme preparation. The GST activities were immediately determined using CDNB and DCNB as substrates as described above. For negative controls, 10 ml of 0.1 M phosphate buffer (pH 7.5) instead of alachlor or DEM were used in the assays. Nonenzymatic controls were also performed in parallel in order to correct for nonenzymatic conjugation of GSH to the substrates. 2.8. Statistical analysis The GST activities and gene expression changes due to the alachlor exposures were subjected to two-way analysis of variance (ANOVA). For percentage data of the relative GST gene expression, data were first transformed using arcsine square root transformation before the ANOVA. Fisher’s least significant difference (LSD) multiple comparisons were then used to separate the means among the treatments. All the statistical analyses were performed using ProStat software (Poly Software International Inc., Pearl River, NY). 3. Results and discussion 3.1. Identification and classification of C. tentans GSTs Insect cytosolic GSTs have been assigned to six classes including delta, epsilon, omega, sigma, theta and zeta (Chelvanayagam et al., 2001; Ranson et al., 2001, 2002; Ding et al., 2003; Yu et al., 2008).

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This classification was primarily based on the identity level of the amino acid residues of an unknown GST compared with known insect GSTs. The GSTs with amino acid sequences sharing over 40% identities are assigned to the same class, and given a number that may reflect the order of discovery or the genome organizations. For example, AgGSTd12 is the 12th member of the delta class of GSTs identified in A. gambiae (Chelvanayagam et al., 2001). Based on this approach, 25 out of 28 GSTs have been assigned to six different classes, including 15 in delta, 8 in epsilon, 2 in theta, 1 in each of omega, sigma and zeta, and 3 unclassified in A. gambiae (Ding et al., 2003), whereas 37 and 21 out of 23 GSTs have also been assigned into six GST classes in D. melanogaster (Enayati et al., 2005) and B. mori (Yu et al., 2008), respectively. We identified 11 different GST transcripts from our C. tentans EST database and revealed their identities by BLASTX search against the non-redundant database at NCBI. We then obtained and verified the complete cDNA sequence within the coding region of each gene by sequencing the same EST clone from both 30 and 50 directions. Our phylogenetic analysis of the 11 GSTs deduced from their cDNAs revealed 7 GSTs that belong to three different cytosolic classes, including 2 in delta (CtGSTd1, CtGSTd2), 4 in sigma (CtGSTs1, CtGSTs2, CtGSTs3, CtGSTs4), and 1 in omega (CtGSTo1), based on their sequence similarities to other insect GSTs, particularly those from A. gambiae and D. melanogaster (Chelvanayagam et al., 2001; Ranson et al., 2001, 2002; Ding et al., 2003), both species are in the same order (Diptera) as the aquatic midge. The remaining four GSTs (CtGSTu1, CtGSTu2, CtGSTu3, CtGSTu4) were unclassified due to their low relatedness to the currently known classes of insect GSTs including A. gambiae and D. melanogaster (Chelvanayagam et al., 2001; Ranson et al., 2001, 2002; Ding et al., 2003; Enayati et al., 2005) (Fig. 1). Specifically, three (CtGSTu1, CtGSTu2, CtGSTu3) of the four unclassified midge GSTs were grouped with the two unclassified A. gambiae GSTs (AgGSTu2, AgGSTu3) and the remaining CtGSTu4 appeared to be more related to the epsilon class of GSTs from A. gambiae and D. melanogaster (Fig. 1). Indeed, Drosophila species also have cytosolic GSTs that do not fit into any of the defined classes (Low et al., 2007). The assignment of the seven midge GSTs to the delta, sigma and omega classes is clearly supported by our phylogenetic analysis of all the 11 C. tentans cytosolic GSTs along with GSTs from both A. gambiae and D. melanogaster (Fig. 1). The percentages of deduced amino acid identities are 47.6% between the two delta GSTs, 45.3–55.7% among the four sigma GSTs, but only 12.2 to 35.0% among different classes of the GSTs in midge GSTs (Table 1). Furthermore, the length of the deduced amino acid residues of 201–229 and conserved amino acid residues found in midge GSTs were consistent with those of other insect GSTs. These C. tentans GST cDNA and their deduced amino acid sequences have been deposited in GenBank with the following accession numbers: FJ851365 (CtGSTd1), FJ851366 (CtGSTd2), FJ851367 (CtGSTs1), FJ851368 (CtGSTs2), FJ851369 (CtGSTs3), FJ851370 (CtGSTs4), FJ851371 (CtGSTu1), FJ851372 (CtGSTu2), FJ851373 (CtGSTu3), FJ851374 (CtGSTu4), and FJ851375 (CtGSTo1). 3.2. Comparison of GSTs between C. tentans and other insects Delta and epsilon are the two largest insect specific GST classes and account for over 65% of the total complement of cytosolic GSTs in the genomes of two dipteran species, D. melanogaster and A. gambiae (Ranson et al., 2002). In contrast, sigma is a very small GST class and only one was found in each of the two dipteran species (Enayati et al., 2005). In this study, we found only two delta GSTs but did not find any epsilon GST from our C. tentans EST database. However, we found four sigma GSTs among the 11 C. tentans GSTs. Relatively few delta, no epsilon, but more sigma

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DmGSTd2

65 83

DmGSTd4 DmGSTd5

25

DmGSTd6

34 75

DmGSTd7

37

DmGSTd3 DmGSTd1

49 72

DmGSTd10

52

DmGSTd8 DmGSTd9

65

AgGSTd2 24

CtGSTd1

37

AgGSTd1

43 50

AgGSTd3

Delta

CtGSTd2 37

DmGSTd-CG17639

99

20

AgGSTd7 AgGSTd8 AgGSTd11

57

AgGSTd6

23 77

AgGSTd9 AgGSTd12

33

AgGSTd4

49

AgGSTd5

96 99

10

AgGSTd10 AgGSTu1 CtGSTu1 AgGSTu3

89 16

CtGSTu2

23 87

Unclassified

CtGSTu3

29

AgGSTu2 DmGSTe-CG4688

70 20

AgGSTe8 CtGSTu4 DmGSTe-CG11784 AgGSTe1

91 82

25

AgGSTe2 AgGSTe7

43

AgGSTe4 51

98

AgGSTe5

36

DmGSTe-CG16936

42 26

Epsilon*

DmGSTe-CG5224

24

AgGSTe3 AgGSTe6

82

DmGSTe3

39

DmGSTe1

82

DmGSTe2

64

DmGSTe10 DmGSTe9

19

DmGSTe4

43

DmGSTe7

97

DmGSTe8

95

DmGSTe5

57 54

DmGSTe6 99

99

DmGSTz-CG9363 AgGSTz1

Zeta

DmGSTz-CG9362 DmGSTo-CG6781

88 99 99

31

DmGSTo-CG6673PB DmGSTo-CG6673PA DmGSTo-CG6776

41

Omega

DmGSTo-CG6662 AgGSTo1

85 61

CtGSTo1 DmGSTt-CG30000

99 69

DmGSTt-CG30005 DmGSTt-CG1702

99

DmGSTt-CG1681

Theta

AgGSTt1

64 89

GSTs identified in this study is likely biased by the limited number of ESTs examined, although we can not rule out possible variations regarding the composition of different GSTs among the insect species. According to the numbers of GSTs identified in the two dipteran species: 28 in A. gambiae (Ding et al., 2003) and 37 in D. melanogaster (Enayati et al., 2005), we probably only obtained 1/ 3 of GSTs that might exist in C. tentans. Nevertheless, our estimation is rather rough since we did not take the insect developmental stage or tissue type that was used to construct cDNA libraries into account. Alternative splicing of delta GST genes has been reported as a unique phenomenon in mosquitoes, including A. gambiae (Ranson et al., 1998), Anopheles dirus (Pongjaroenkit et al., 2001), Aedes aegypti (Lumjuan et al., 2007), and Culex quinquefasciatus (Kasai et al., 2009). A total of four different GSTd1 transcripts have been identified in A. gambiae, A. dirus, and C. quinquefasciatus. However, only three GSTd1 variants were identified in A. aegypti, suggesting that one variant could be lost in A. aegypti, or gained in A. gambiae, A. dirus, and C. quinquefasciatus during the evolution of the mosquitoes (Lumjuan et al., 2007; Kasai et al., 2009). Because such a phenomenon of alternative splicing of delta GST genes has not been found in other insect species such as Apis mellifera or D. melanogaster, it is probably mosquito-specific (Kasai et al., 2009). Although C. tentans is closely related to mosquito species, we do not know if alternative splicing of delta GST genes also occur in this species. All the C. tentans delta and sigma GSTs show the characteristics of other insect delta GSTs (Chen et al., 2003; Mittapalli et al., 2007) and sigma GSTs (Agianian et al., 2003), respectively. Specifically, multiple alignments of the four C. tentans sigma GSTs along with a D. melanogaster sigma GST revealed several key residues that are conserved across different insect species (Fig. 2). These residues constituted the putative GSH binding site (Y54-W85-Q96-S110 numbered based on D. melanogaster GST: DmGSTs1) and the electrophilic-binding site (V57-A59-R145-V150-Y153-Y208-Y211-V249) in the deduced amino acid sequence for the GSTs in the sigma class. In addition, L60 represents the putative H-site residue, interacting with GSH, whereas G204 may be the bulge-inducing residue (Agianian et al., 2003). Similarly, multiple alignments of the amino acid sequences of the two delta GSTs along with those of the delta GSTs from other insect species also revealed conserved amino acid residues across different species (Fig. 3). These delta GSTs contain the amino acid residues S10 and N48 (numbered based on DmGSTd1) which represent the catalytic pocket, and P54-L142-G150-D157 which may be involved in protein folding (Mittapalli et al., 2007). As expected, the amino acid sequences among the four unclassified GSTs (CtGSTu1, CtGSTu2, CtGSTu3, CtGSTu4) and the omega GST (CtGSTo1) are less conserved (Fig. 4). However, three (P59, I74 and D162) out of six conserved amino acid residues (Y10, P59, D63, I74, G155 and D162 numbered based on CtGSTu2) of GSTs, as reported by Wilce and Parker (1994), and Ono et al. (2005), are complete conserved, whereas the remaining three residues are identical in at least three GSTs (Y10 in CtGSTu1, CtGSTu2 and CtGSTu4; D63 in CtGSTu1, CtGSTu3 and CtGSTu4; and G155 in CtGSTu1, CtGSTu2, CtGSTu3 and CtGSTo1) among these five GSTs. Because these GSTs show only <40% identities of amino acid

AgGSTt2 CtGSTs1 CtGSTs3

99

CtGSTs2

45

CtGSTs4

43

DmGSTs1

93 59

0.1

AgGSTs1

Sigma

Fig. 1. Phylogenetic relationships of 76 GST proteins from three dipteran species including C. tentans (Ct, 11), Drosophila melanogaster (Dm, 37) and Anopheles gambiae (Ag, 28). All the 76 GST protein sequences are provided in Supplementary material. Amino acid sequences were aligned using ClustalW (www.ebi.ac.uk/clustalW) and a distance neighbour-joining tree was generated using MEGA. Nodes with distance bootstrap values (1000 replicates) are shown. The 11 C. tentans GSTs are marked with filled circles. The letter ‘‘u’’ in each of the four CtGSTs (CtGSTu1, CtGSTu2, CtGSTu3 and CtGSTu4) simply refers to ‘‘unclassified’’ due to their low relatedness to the currently known classes of insect GSTs (see text in Section 3.1). These four CtGSTs share <40% amino acid identities from each other.

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Table 1 Percent identities of amino acid residues among the 11 C. tentans GSTs.

CtGSTd1 CtGSTd2 CtGSTo1 CtGSTs1 CtGSTs2 CtGSTs3 CtGSTs4 CtGSTu1b CtGSTu2 CtGSTu3 CtGSTu4

CtGSTd1

CtGSTd2

CtGSTo1

CtGSTs1

CtGSTs2

CtGSTs3

CtGSTs4

CtGSTu1

CtGSTu2

CtGSTu3

CtGSTu4

–

47.6a –

19.5 18.1 –

17.1 13.6 13.5 –

17.1 16.8 12.5 45.3 –

17.1 14.7 13.1 45.3 49.3 –

14.8 12.4 14.4 46.7 55.7 55.7 –

31.6 30.0 19.4 15.0 15.6 19.5 16.8 –

29.2 27.2 13.9 14.6 16.8 17.2 16.9 27.2 –

32.6 33.7 21.7 14.6 15.2 13.6 12.2 28.6 35.0 –

32.9 34.6 19.2 15.5 14.0 16.5 16.2 22.5 29.4 28.0 –

a The percentages of the amino acid identity of different GSTs were determined using the DNASTAR software. The italic numbers indicate the amino acid identities of the GSTs within the same class. b The letter ‘‘u’’ in each of the four GSTs (CtGSTu1, CtGSTu2, CtGSTu3 and CtGSTu4) simply refers to ‘‘unclassified’’ due to their low relatedness to the currently known classes of insect GSTs (see text in Section 3.1). These unclassified GSTs show only <40% identities of amino acid residues as compared with each other and, therefore, are unlikely to belong to the same class.

residues as compared with each other, they are likely to belong to different classes. 3.3. Tissue- and stage-specific expression patterns of GST genes in C. tentans Tissue-specific expression patterns of the 11 C. tentans GST genes were analyzed in each of six different tissues, including salivary glands, hemolymph, midgut, Malpighian tubules, fatbodies, and carcass, by using RT-PCR. Our results indicated that eight of the 11 GST genes, including CtGSTd1, CtGSTu2, CtGSTu4, CtGSTs1, CtGSTs2, CtGSTs3, CtGSTs4 and CtGSTo1, were expressed in all tissues examined, although there were some noticeable variations in expression levels among different tissues (Fig. 5). Specifically, CtGSTd2 was expressed in a limited number of tissues including midgut and fatbodies, whereas CtGSTu1 was virtually undetectable in salivary glands and hemolymph. The widespread expressions of GST genes in midgut and fatbodies have also been demonstrated in other insect species including D. melanogaster (Nakamura et al., 1999; Chintapalli et al., 2007) and lepidopterans (Snyder et al., 1995; Krishnan and Kodrı´k, 2006). Indeed, 74% of the GST genes found in the genome of

D. melanogaster were observed to be expressed in the midgut by oligoarray analysis (Li et al., 2008). The GST expression has also been observed in Malphigian tubules in other insect species (Konno and Shishido, 1992). For example, one of the GST genes (GstE10) was expressed at 40-fold higher in Malpighian tubules than in the whole fly of D. melanogaster (Wang et al., 2004). Stage-specific expression patterns of C. tentans GST genes were determined in eggs, four different larval instars (1st, 2nd, 3rd and 4th), pupae and adults by using RT-PCR. Among the 11 genes, only two genes (CtGSTs1 and CtGSTs4) showed significant expressions in eggs, whereas the remaining nine genes did not show any detectable expressions in eggs (Fig. 6). In contrast, all the 11 GST genes showed various levels of expressions in all the four larval instars, whereas the expressions of CtGSTd2, CtGSTu1 and CtGSTu2 were not detectable in pupal and adult stages. 3.4. In vivo and in vitro inhibition of GST activities by alachlor in C. tentans The in vivo inhibitions of GST activities by the herbicide alachlor were compared among the fourth-instar midges exposed to water containing the solvent acetone (control) and different

Fig. 2. Similarity comparisons of the amino acid sequences of four C. tentans sigma GSTs (CtGSTs1, CtGSTs2, CtGSTs3, CtGSTs4) with a Drosophila melanogaster sigma GST (DmGSTs1, AAM48357). The amino acid residues of the sigma GSTs, which are shaded in green and yellow, represent residues that constitute the putative glutathione (GSH) and electrophilicsubstrate binding sites, respectively. The putative H-site residue of C. tentans sigma GSTs (L14 in CtGSTs4), which also contacts GSH, is shaded in red. The putative bulge-inducing residues (A156 in CtGSTs4, A158 in CtGSTs2, G159 in CtGSTs3, and A156 in CtGSTs1) of C. tentans sigma GSTs are shaded in blue. Dashes are used to denote gaps introduced for a maximum alignment. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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Fig. 3. Similarity comparisons of the amino acid sequences of two C. tentans dalta GSTs (CtGSTd1, CtGSTd2) with a D. melanogaster (DmGSTd1, AAM52032) and an Anopheles gambiae delta (AgGSTd1, 40889324) GSTs. S9 and N47 of CtGSTd1 and CtGSTd2, which represent the catalytic pocket, are shaded in red. Amino acid residues determining folding are shaded in green. Dashes are used to denote gaps introduced for a maximum alignment. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

concentrations of the herbicide alachlor for 72 h. Alachlor at 10, 100, 1000 mg/L decreased the GST activities by 1.3-, 1.7-, and 2.2-fold, respectively, compared with those of the control when CDNB were used as a substrate (Table 2). Similarly, alachlor at 10, 100, 1000 mg/L decreased the GST activities by 1.3-, 1.7-, and 1.8-fold, respectively, when DCNB was used as a substrate. These results were consistent not only between the two substrates used in this study, but also with the results of a recent report by Jin-Clark et al. (2008), which showed significant decreases of GST activities in C. tentans larvae exposed to alachlor. Significant inhibitions of GST activities by alachlor were also observed in in vitro assays (Fig. 7). Alachlor at 1000 mg/L decreased GST activity by 28.3% as compared with the control when CDNB was used as a substrate, whereas alachlor at 100 and 1000 mg/L decreased the enzyme activities by 65.5 and 73.5%, respectively, as compared with the control when DCNB was used as a substrate. In order to examine whether or not DEM, a known GST inhibitor (Susanto et al., 1998), can also inhibit GST activities in vitro, we performed parallel assays using DEM. Indeed, DEM at 100 and

1000 mg/L significantly decreased the GST activities when CDNB was used as a substrate. Because DEM is an effective GSH depletor at relatively high concentrations, its inhibition to GST activities is likely due to rapid depletion of GSH in our assays. Both the chloroacetanilide herbicides alachlor and metolachlor have been known to inhibit protein synthesis in plants thereby conferring herbicide toxicity (Chesters et al., 1989; HSDB, 2002). Previous study also showed that metolachlor reduced a total protein production by 3.2-fold in the midges exposed to the chemical at 1000 mg/L for 72 h (Jin-Clark et al., 2008). However, alachlor did not cause a significant reduction of total protein of the midge in a parallel study (Jin-Clark et al., 2008). It is possible that the concentration necessary for alachlor to affect the protein synthesis in midges is higher than that of metolachlor. Nevertheless, it has been reported that alachlor is an electrophilic substrate that can be conjugated by GSH (Zablotowicz et al., 1995). Indeed, many bacteria (Stamper and Tuovinen, 1998) and plants (Rossini et al., 1998; Labrou et al., 2005) possess GSTs capable of conjugating alachlor to GSH. Thus, significant inhibitions of GST

Fig. 4. Similarity comparisons of the amino acid sequences among the four unclassified GSTs (CtGSTu1, CtGSTu2, CtGSTu3, CtGSTu4) and the omega GST (CtGSTo1) from C. tentans. Amino acid residues shaded in red represent six conserved amino acid residues (Y10, P59, D63, I74, G155 and D162 numbered based on CtGSTu2) typically found in GST proteins. Dashes are used to denote gaps introduced for a maximum alignment. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

X. Li et al. / Insect Biochemistry and Molecular Biology 39 (2009) 745–754

Fig. 5. Tissue-specific expression patterns of the 11 GST genes in C. tentans as evaluated using RT-PCR in salivary glands (SG), hemolymph (HL), midgut (MG), Malpighian tubules (MT), carcasses (CA) and fatbody (FB). The ribosomal S3 (CtRPS3) gene was used as a reference gene.

activities by alachlor as observed in our in vitro inhibition studies suggested that alachlor’s inhibition to GSTs was probably due to a rapid conjugation between alachlor and GSH, therefore, leading to the depletion of GSH for the enzyme activities. Such depletions due to the presence of alachlor can occur both in vivo and in vitro assays of the GST activities as observed in our studies. The inhibition mechanism between DEM and alachlor are likely the same because both can be conjugated by GSH, therefore leading to the depletion of GSH. However, the effect of alachlor on GST activities was more pronounced when DCNB was used as a substrate as compared with CDNB (Fig. 7). This can probably be explained by DCNB being a less reactive substrate than CDNB for GSH conjugation (Patskovsky et al., 2000). Thus, the effect of GSH depletion on GST activities would be expected to be greater for DCNB than for CDNB as substrates.

Fig. 6. Developmental stage-specific expression patterns of the 11 GST genes in C. tentans as evaluated using RT-PCR in eggs (EG); first- (L1), second- (L2), third- (L3) and fourth-instar (L4) larvae; pupae (PU) and adults (AD). The ribosomal S3 (CtRPS3) gene was used as a reference gene.

alachlor significantly increased the mRNA levels of the CtGSTd1, CtGSTs2 and CtGSTs3 genes in the midges exposed to alachlor (Fig. 8). Specifically, the exposure of forth-instar larvae to alachlor at 1000 mg/L for 72 h increased the CtGSTd1, CtGSTs2 and CtGSTs3 mRNA levels by 2.1-, 2.8- and 4.3-fold, respectively. In addition, the increased expressions of CtGSTs2 and CtGSTs3 appeared to positively correlate with the increases of alachlor concentration. These results suggested that alachlor can actually induce the expression of some genes in at least two classes of the GST gene family, including delta (CtGSTd1) and sigma (CtGSTs2, CtGSTs3). Furthermore, these results clearly indicated that decreased GST activities in the midges exposed to alachlor were not due to reduced expression of GSTs at transcriptional levels.

Table 2 Comparisons of the GST activities among fourth-instar larvae of C. tentans exposed to water containing the solvent acetone (control) or the herbicide alachlor for 72 h. GST activity

CDNB (nmol/min/mg protein)

DCNB (nmol/min/mg protein)

Acetone (100 ml/L) Alachlor (1 mg/L) Alachlor (10 mg/L) Alachlor (100 mg/L) Alachlor (1000 mg/L)

502.37 492.28 388.22 304.03 227.32

48.40 50.47 38.63 29.67 26.65

3.5. Effect of alachlor on GST gene expression in C. tentans To clarify whether or not the decreased GST activities in the midges exposed to alachlor was due to the reduced expression of the GST genes at the transcriptional level, we first used the RT-PCR to examine the relative transcriptional levels of 11 GST genes and then used qPCR to quantify the expression changes for the genes that were revealed by RT-PCR. Our results clearly showed that

751

(21.89) a (65.95) a (15.63) b (7.44) c (13.77) d

(1.96) (6.45) (6.69) (3.48) (3.79)

a a b bc c

Data are presented as the mean  standard error (n ¼ 3). Means followed by the same letter within the same column are not significantly different (p  0.05, Fisher’s LSD multiple comparison test).

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CDNB

a 250

ab

ab b

ab

b

50 b

CtGSTd1

200 40

a 30

ab

100

ab ab

50

20

0

10 a

14

a

DCNB

ab

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8 bc

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cd 4 2 0

l

A A A M M M AL AL AL DE DE DE L L L L L L / / / / / / C µg µg µg µg µg µg 0 10 00 00 00 10 1 0 0 10 1 1

t on

ro

1

10

100

1000

50

ab

10

Fig. 7. In vitro inhibition of GST extracted from fourth-instar C. tentans by three different concentrations of diethyl maleate (DEM) and alachlor (ALA) using CDNB and DCNB as substrates. Vertical bars indicate standard errors of the mean (n ¼ 4). Different letters on the bars indicate that the means are significant different among the treatments in Fisher’s LSD multiple comparison tests (p < 0.05).

Indeed, it has been reported that alachlor may regulate different GST genes using a switchable regulation mechanism in maize (Rossini et al., 1998). Such a mechanism appears to exist also in C. tentans because expressions of only 3 (CtGSTd1, CtGSTs2 and CtGSTs3) out of 11 GST genes were up-regulated by alachlor. The delta GSTs have been considered to have a key role in detoxifying xenobiotics including insecticides (Poupardin et al., 2008) and plant allelochemicals (Ahmad and Pardini, 1990), and in conferring insecticide resistance (Papadopoulos et al., 2000; Ranson et al., 2001; Ortelli et al., 2003). In contrast, the functions of the sigma GSTs are unclear among the insect GST family. Sigma GSTs in D. melanogaster had been initially reported to have structural functions in indirect flight muscle tissues (Bullard et al., 1988; Clayton et al., 1998). However, the catalytic specificity of the sigma GST (GST-2) was established by Singh et al. (2001) and confirmed by Agianian et al. (2003). The enzyme shows a clear preference for elongated unsaturated compounds with carbonyl oxygen atoms. These results suggested a distinctly different function for a sigma GST as compared with the delta GSTs. However, further studies would be necessary to understand the mechanism conferring the increased gene expressions of CtGSTd1, CtGSTs2 and CtGSTs3 in the midge larvae exposed to alachlor and to examine the effect of such

Relative gene expression (%)

GST specific activity (nmol/min/mg protein)

150

CtGSTs2 40

a ab 30

bc bc

20

c 10

0 1

0

10

100

1000

50

CtGSTs3

a

40

30

b bc

20

bc c

10

0 0

1

10

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1000

Concentration of alachlor (µg/L) Fig. 8. Quantification of relative expression levels of three GST genes (CtGSTd1, CtGSTs2 and CtGSTs3) in the fourth-instar C. tentans exposed to water containing acetone (100 ml/L) as control (0) and the herbicide alachlor at four different concentrations (1, 10, 100, and 1000 mg/L) for 72 h. The mRNA level in the control and each treatment was normalized using CtRPS3 as a reference gene. Vertical bars indicate standard errors of the mean (n ¼ 4). Different letters on the bars indicate that the means are significantly different among the control and treatments in Fisher’s LSD multiple comparison test (p < 0.05).

X. Li et al. / Insect Biochemistry and Molecular Biology 39 (2009) 745–754

alterations on the responses of the midge to various environmental toxic chemicals in aquatic ecosystems. Acknowledgements The authors thank Jianxiu Yao, Chitvan Khajuria, the editor, and three anonymous reviewers for their helpful comments; Ludex Zurek for letting us use his iCycler iQ real-time PCR detection system; and Ming-Shun Chen for reviewing an earlier version of this manuscript. This research was partially supported by K-State Ecological Genomics Institute funded by K-State Targeted Excellence and Kansas Agricultural Experiment Station to KYZ, and China Scholarship Council to XL. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by Kansas State University. This manuscript is contribution No. 09-288-J from Kansas Agricultural Experiment Station. The Chironomus tentans voucher specimens (110) are located in the Museum of Entomological and Prairie Arthropod Research, Kansas State University, Manhattan, Kansas, USA. Appendix. Supplementary material Supplementary data associated with this article can be found in the online version at doi:10.1016/j.ibmb.2009.08.010. References Agianian, B., Tucker, P.A., Schouten, A., Leonard, K., Bullard, B., 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, 151–165. Ahmad, S., Pardini, R.S., 1990. Antioxidant defense of the cabbage looper, Trichoplusia ni: enzymatic responses to the superoxide-generating flavonoid, quercetin, and photodynamic furanocoumarin, xanthotoxin. Photochem. Photobiol. 51, 305–311. Anderson, T.D., 2006. Toxicological, Biochemical, and Molecular Effects of Atrazine to the Aquatic Midge Chironomus tentans (Diptera: Chironomidae). Ph.D. Dissertation, Kansas State University, Manhattan, KS. Anderson, T.D., Jin-Clark, Y., Begum, K., Starkey, S.R., Zhu, K.Y., 2008. Gene expression profiling reveals decreased expression of two hemoglobin genes associated with increased consumption of oxygen in Chironomus tentans exposed to atrazine: a possible mechanism for adapting to oxygen deficiency. Aquat. Toxicol. 86, 148–156. Anderson, T.D., Lydy, M.J., 2001. Increased toxicity to invertebrates associated with a mixture of atrazine and organophosphate insecticides. Environ. Toxicol. Chem. 21, 1507–1514. Anderson, T.D., Zhu, K.Y., 2004. Synergistic and antagonistic effects of atrazine on the toxicity of organophosphorodithioate and organophosphorothioate insecticides to Chironomus tentans (Diptera: Chironomidae). Pestic. Biochem. Physiol. 80, 54–64. Belden, J.B., Lydy, M.J., 2000. Impact of atrazine on organophosphate insecticide toxicity. Environ. Toxicol. Chem. 19, 2266–2274. 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., 2000. Identification, characterisation, and crystal structure of the Omega class glutathione transferases. J. Biol. Chem. 275, 24798–24806. Bullard, B., Leonard, K., Larkins, A., Butcher, G., Karlik, C., Fyrberg, E., 1988. Troponin of asynchronous flight muscle. J. Mol. Biol. 204, 621–637. Che-Mendoza, A., Penilla, R.P., Rodrı´guez, D.A., 2009. Insecticide resistance and glutathione S-transferases in mosquitoes: a review. Afr. J. Biotechnol. 8, 1386–1397. Chelvanayagam, G., Parker, M.W., Board, P.G., 2001. Fly fishing for GSTs: a unified nomenclature for mammalian and insect glutathione transferases. Chem.-Biol. Inter. 133, 256–260. Chen, L., Hall, P.R., Zhou, X.E., Ranson, H., Hemingway, J., Meehan, E.J., 2003. Structure of an insect delta-class glutathione S-transferase from a DDT-resistant strain of the malaria vector, Anopheles gambiae. Acta Cryst. D59, 2211–2217. Chesters, G.C., Simsiman, G.V., Levy, J., Alhajja, B.J., Fathulla, R.N., Harkin, J.M., 1989. Environmental fate of alachlor and metolachlor. Rev. Environ. Contam. Toxicol. 110, 1–74. Chintapalli, V.R., Wang, J., Dow, J.A.T., 2007. Using FlyAtlas to identify better Drosophila melanogaster models of human disease. Nat. Genet. 39, 715–720. Clayton, J.D., Cripps, R.M., Sparrow, J.C., Bullard, B., 1998. Interaction of troponin-H and glutathione S-transferase-2 in the indirect flight muscles of Drosophila melanogaster. J. Muscle Res. Cell Motil. 19, 117–127.

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