- Research article
- Open Access
Characterization and expression profiling of glutathione S-transferases in the diamondback moth, Plutella xylostella (L.)
- Yanchun You†1, 2, 3,
- Miao Xie†1, 2, 3,
- Nana Ren1, 3,
- Xuemin Cheng1, 3,
- Jianyu Li1, 2, 3,
- Xiaoli Ma1, 3,
- Minming Zou1, 3,
- Liette Vasseur1, 4,
- Geoff M Gurr1, 3, 5 and
- Minsheng You1, 3Email author
© You et al.; licensee BioMed Central. 2015
- Received: 12 September 2014
- Accepted: 12 February 2015
- Published: 5 March 2015
Glutathione S-transferases (GSTs) are multifunctional detoxification enzymes that play important roles in insects. The completion of several insect genome projects has enabled the identification and characterization of GST genes over recent years. This study presents a genome-wide investigation of the diamondback moth (DBM), Plutella xylostella, a species in which the GSTs are of special importance because this pest is highly resistant to many insecticides.
A total of 22 putative cytosolic GSTs were identified from a published P. xylostella genome and grouped into 6 subclasses (with two unclassified). Delta, Epsilon and Omega GSTs were numerically superior with 5 genes for each of the subclasses. The resulting phylogenetic tree showed that the P. xylostella GSTs were all clustered into Lepidoptera-specific branches. Intron sites and phases as well as GSH binding sites were strongly conserved within each of the subclasses in the GSTs of P. xylostella. Transcriptome-, RNA-seq- and qRT-PCR-based analyses showed that the GST genes were developmental stage- and strain-specifically expressed. Most of the highly expressed genes in insecticide resistant strains were also predominantly expressed in the Malpighian tubules, midgut or epidermis.
To date, this is the most comprehensive study on genome-wide identification, characterization and expression profiling of the GST family in P. xylostella. The diversified features and expression patterns of the GSTs are inferred to be associated with the capacity of this species to develop resistance to a wide range of pesticides and biological toxins. Our findings provide a base for functional research on specific GST genes, a better understanding of the evolution of insecticide resistance, and strategies for more sustainable management of the pest.
- Transcriptome analysis
- Phylogenetic analysis
- Insect pest
The diamondback moth (DBM), Plutella xylostella (L.) (Lepidoptera: Plutellidae), is a world-wide destructive pest of wild and cultivated crucifers . The larvae feed on cruciferous plants and may cause significant reductions in yield and quality of economically important crops such as canola and cabbage. Historical reliance on insecticides has led to the rapid development of resistance in P. xylostella populations , making it difficult to control.
Several studies have examined the potential mechanisms underlying the development of insecticide resistance in P. xylostella [3-5]. One of the proposed mechanisms is metabolic resistance through the multifunctional glutathione S-transferases (GSTs, EC18.104.22.168). These enzymes can catalyze electrophilic compounds, making them water soluble and readily excreted . GSTs are known more generally by insects to detoxify various xenobiotics, including insecticides and plant allelochemicals . The recent work has focused on the potential role of GSTs in oxidative stress responses [6,8-11].
Insect GSTs are classified as cytosolic and microsomal. The number of microsomal GSTs is much lower than that of cytosolic GSTs, which have been grouped into six subclasses . Delta and Epsilon subclasses are insect specific, while the other four subclasses, Omega, Sigma, Theta, and Zeta, are found in various animal taxa [10,13,14].
GSTs are involved in the resistance of insects to organophosphate (OPs), chlorine, and pyrethroid insecticides [15,16]. Recombinant GST enzymes from P. xylostella and Drosophila melanogaster have been shown to play a role in the metabolism of organophosphate insecticides [17,18]. It has been suggested that, under elevated GST activity conditions, Anopheles subpictus can detoxify fenitrooxon activation products, leading to organophosphate resistance . The silkworm Zeta GST recombinant protein (rbmGSTz) has been found to initiate the dechlorination of permethrin and to be abundantly distributed in a permethrin-resistant strain . Similarly, an Omega GST is highly expressed in a fenitrothion-resistant strain of silkworm and its recombinant protein (rbmGSTo) shows high affinity with organophosphate insecticides, indicating that it may contribute to insecticide resistance and oxidative stress responses . The antennae-specific GST was found being involved with detoxification of xenobiotics and detection of sex pheromones in Manduca sexta .
The GSTs were found to be one of the major enzyme families in the P. xylostella genome and to be linked to detoxification of plant defense compounds and insecticides . A recent study on the identification and characterization of multiple glutathione S-transferase genes  based on the DBM transcriptome database [24,25] provides a primary base for further investigation of this important gene family. In the present study, the P. xylostella GSTs (PxGSTs) were identified and compared with the equivalent information from published insect genomes to better reveal their phylogenetic relationships and intron-exon organization. We profiled and analyzed expression patterns of the PxGSTs using the published transcriptome  and reverse transcription-quantitative polymerase chain reaction (qRT-PCR) in different life stages and tissues from insecticide susceptible or resistant strains. We then examined the major characteristics of GST subclasses and some particular GST genes in relation to their potential roles in P. xylostella insecticide resistance.
Identification of the PxGSTs
Description of 22 identified cytosolic GSTs in the P. xylostella genome
Gene size (bp)
Gene ID a
Comparison of GST gene numbers of various insect species*
Phylogenetic analysis of the PxGSTs
Delta and Epsilon GST subclasses are unique to insects and have been suggested to be implicated in insecticide resistance [10,36,37]. The earlier diverging insects Hymenoptera and Exopterygota do not have Epsilon subclass GSTs. Hymenoptera has a few genes present in the Delta subclass but none for Exopterogota suggesting that these orders may be from an evolutionary older lineage . Our tree suggests that Delta and Epsilon GSTs have diverged more recently from the other subclasses.
The range of amino acid identities in the insect-specific GSTs of P. xylostella are fairly variable, ranged from 38.39 ~ 84.75% in Delta and 23.05 ~ 60.91% in Epsilon (Additional file 4: Table S1). Except for PxGSTe1, the remaining Epsilon PxGSTs were clustered in a monophyletic clade of Lepidoptera (Figure 1), suggesting a lineage-specific expansion within the Epsilon subclass in lepidopteran order.
Characterization of the PxGST introns
In the PxGSTs, the splice sites of introns were classified into three phases: 0 with 45 introns, 1 with 17 introns, and 2 with 18 introns, according to their positions in the codons. The phase-1 introns were present only in the Omega, Zeta and Sigma subclasses. Most phase-2 introns were found in insect-specific subclasses (Delta and Epsilon) as well as in Zeta subclass. Most of the PxGST introns spliced in a given site tended to be from the same phase, suggesting that they might be relatively conserved (Figure 2).
Intron sites are similar across different PxGST subclasses. There are three highly conserved sites of introns within the Delta and Epsilon GST subclasses, except for the PxGSTd1 and PxGSTe1. These were between the 47th and 51st, the 114th and 118th and the 180th and 186th amino acids. Most of the PxGSTs tended to have a nearby conserved site of the introns located between the 111th and 125th amino acids belonging to phase 0. Both of the intron sites and phases were strongly conserved within Sigma and Zeta subclasses (Figure 2), implying that these genes might have similar functions. There appeared to be a correlation between the intron conservation and the phylogenetic cluster within a given PxGST subclass, indicating that gene structure evolution might be involved in the phylogenetic development of a specific subclass.
Despite the conserved nature of intron sites and phases, the lengths were highly variable in the PxGSTs ranging from 28 to 17,644 bp with a larger proportion ranging from 300 to 399 bp (Additional file 5: Figure S1) and an average of 918 bp. The shortest intron was PxGSTu1 (28 bp), while the longest were PxGSTo3 (17,644 bp) and PxGSTz2 (13,241 bp). A previous study has shown that long introns were considered to involve more functional elements than short introns and could effectively regulate gene expressions, possibly via the formation of pre-mRNA secondary structures . However, the function of the longest introns in PxGSTo3 and PxGSTz2 needs to be further investigated.
GSH and substrate binding sites in the PxGSTs
Stage-specific expression profiling
Strain- and tissue-specific expression profiling
To date, this is the most comprehensive study on genome-wide identification, characterization and expression profile of the GSTs in P. xylostella. Twenty-two GSTs were found in P. xylostella, which is similar in number to another lepidopteran species, B. mori. Variable features and different expression patterns of the genes reveal that the P. xylostella GSTs are evolutionary and functionally diversified, and may be involved in the evolution of adaptive capacity in response to environmental variation. Because GST enzymes are considered to be important in insecticide resistance, many of these newly identified genes are potential candidates for inhibiting the pathway of insecticide resistance and targeting lepidopteran-selective insecticides. Thus, further functional research on the PxGSTs is essential to identify the key genes and their roles in xenobiotic detoxification of insects, and understand the mechanisms underlying the insecticide resistance.
Experimental DBM strains
The experimental population of P. xylostella was derived from a susceptible strain (SS) that was collected from a vegetable field of Fuzhou (26.08°N, 119.28°E) in 2004 and used for genome sequencing . Since then this initial population was reared on potted radish seedlings (Raphanus sativus L.) at 25 ± 1°C, 65 ± 5% RH and L:D = 16:8 h in a separate greenhouse without exposure to insecticides over the past ten years. Two insecticide resistant strains (chlorpyrifos- and fipronil-resistant strains (CRS and FRS)) were selected from this susceptible strain, and detailed in DBM transcriptome .
Identification of P. xylostella GST genes
To identify putative GST genes from the DBM genome database [23,27], the GST protein sequences of D. melanogaster, C. quinquefasciatus, A. aegypti, A. gambiae, T. castaneum, A. mellifera, N. vitripennis, A. pisum, P. humanus, B. mori and other lepidopteran insects were downloaded from their genome databases [53-57] and/or GenBank (http://www.ncbi.nlm.nih.gov/) and Uniprot (http://www.uniprot.org/). These insect GST protein sequences were used as queries to perform local TBLASTN searches against the DBM genome database. The putative genomic sequences were retrieved, and then predicted using Fgenesh + (http://www.softberry.com/). The DBM GST protein sequences were confirmed using online BLASTP in NCBI.
The GSTs of A. aegypti (Aa), A. gambiae (Ag), D. melanogaster (Dm), A. mellifera (Am), N. vitripennis (Nv), Papilio polytes (Pp), Danaus plexippus (Dp), B. mori (Bm) and P. xylostella (Px) were used for the phylogenetic analysis. Putative amino acid sequences of the GSTs were aligned using Clustal X2.0 , and then gaps and missing data were manually trimmed. A phylogenetic tree was constructed with the neighbor-joining method  using MEGA 5.10 . Bootstrap analysis with 1,000 replicates was used to evaluate the significance of the nodes. Poisson correction amino acid model and pairwise deletion of gaps were selected for the tree reconstruction.
RNA extraction and cDNA synthesis
DBM eggs, 1st- to 4th-instar larvae, pupae and adults from susceptible and resistant strains were frozen in liquid nitrogen. Total RNA was extracted using the RNAiso Plus (Takara, Code: D9108A, Japan). The 4th-instar larvae were surface sterilized in 75% ethanol, then dipped in DNAase and RNAse free water and dissected. Tissues (head, midgut, Malpighian tubules, fatbody and epidermis) from resistant strains were briefly immersed in RNAlater™ RNA Stabilization Reagent (QIAGEN, Code: 76104, Germany) then stored at 4°C. Total RNA was extracted with the RNeasy Plus Micro Kit (QIAGEN, Code: 74034, Germany) and RNA concentration was determined using a spectrophotometer (Nanodrop 2000: Thermo, USA).
The cDNA template for PCR was synthesized with 1 μg of total RNA using PrimeScript®RT reagent Kit with gDNA Eraser (Perfect Real Time) (Takara, Code: DRR047A, Japan).
Validation of gene expression by qRT-PCR
The qRT-PCR primers used in the validation of gene expression were identified based on the encoding sequences of the DBM GSTs (Additional file 9: Table S5). DBM ribosomal protein L8 (RPL8) was used as reference gene for different strains and tissues, and DBM ribosomal protein S4 (RPS4) for different stages/instars. The assays were run in triplicate in CFX96 Touch™ Real-Time PCR Detection Systems (Bio-Rad, USA). PCR amplification was performed in a total reaction volume of 20 μL reaction mixture, containing 20 ng cDNA, 10 μL 2 × SYBR® Premix Ex Taq™ (Takara, DRR420A, Japan), 0.2 μM of each primer. PCR was conducted with standard thermal cycle conditions using the two-step qRT-PCR method: an initial denaturation at 95°C for 30s followed by 40 cycles of 3s at 95°C and 30s at 60°C. Specificity of the PCR products was assessed by melting curve analysis for all samples. For each treatment (tissues, strains and developmental stages), there were three biological replicates.
The 2−ΔCt method was used to analyze the qRT-PCR-based expression patterns. One-way ANOVA, using PASW Statistics 18, followed by a Duncan’s multiple range test was used to evaluate significant differences among patterns. The results were presented by mean ± standard deviation of the relative mRNA expressions.
Availability of supporting data
The nucleic acid sequences and protein sequences have been deposited in the published DBM genomic database (DBM-DB: http://iae.fafu.edu.cn/DBM/family/PxGSTs.php). Other supporting data are presented in Additional file 7: Tables S3 and Additional file 8: Table S4, and also deposited in the same database.
We are grateful to Liying Yu for her advice on analysis of gene family and phylogenetic analysis, and to Drs. Simon W. Baxter and Mark S. Goettel for their comments on our manuscript at early stage. We thank Yimin Zhuo, Jinyang Cai, Qiuye Qiu, Wenxiu Xie, Xianbin Huang and Xiaohui Huang for their technical assistance in rearing diamondback moth and preparing samples. The work was supported by National Natural Science Foundation of China (No. 31320103922 and No. 31230061) and National Key Project of Fundamental Scientific Research (“973” Programs, No. 2011CB100404) in China. LV is supported by the Minjiang Scholar Program in Fujian Province (PRC), and GMG by the National Thousand Talents Program in China.
- Furlong MJ, Wright DJ, Dosdall LM. Diamondback moth ecology and management: problems, progress, and prospects. Annu Rev Entomol. 2013;58:517–41.View ArticlePubMedGoogle Scholar
- Georghiou GP. Overview of insecticide resistance. In: Managing resistance to agrochemicals. vol. 421. Washington, DC: American Chemical Society; 1990. p. 18–41.View ArticleGoogle Scholar
- Sonoda S. Molecular analysis of pyrethroid resistance conferred by target insensitivity and increased metabolic detoxification in Plutella xylostella. Pest Manage Sci. 2010;66(5):572–5.View ArticleGoogle Scholar
- Kim JI, Joo YR, Kwon M, Kim GH, Lee SH. Mutation in ace1 associated with an insecticide resistant population of Plutella xylostella. J Asia-Pacif Entomol. 2012;15(3):401–7.View ArticleGoogle Scholar
- Dukre AS, Moharil MP, Ghodki BS, Rao NGV. Role of glutathione S-transferase in imparting resistance to pyrethroids in Plutella xylostella(L.). Int J Integr Bio. 2009;6(1):17–21.Google Scholar
- Ranson H, Hemingway J. 5.11 - Glutathione Transferases. In: Gilbert LI, editor. Compr Mol Insect Sci. Amsterdam: Elsevier; 2005. p. 383–402.View ArticleGoogle Scholar
- Enayati AA, Ranson H, Hemingway J. Insect glutathione transferases and insecticide resistance. Insect Mol Biol. 2005;14(1):3–8.View ArticlePubMedGoogle Scholar
- Clark AG. The comparative enzymology of the glutathione S-transferases from non-vertebrate organisms. Comp Biochem Physiol B: Comp Biochem. 1989;92(3):419–46.Google Scholar
- Fournier D, Bride JM, Poirie M, Berge JB, Plapp Jr FW. Insect glutathione S-transferases. Biochemical characteristics of the major forms from houseflies susceptible and resistant to insecticides. J Biol Chem. 1992;267(3):1840–5.PubMedGoogle Scholar
- Ranson H, Rossiter L, Ortelli F, Jensen B, Wang X, Roth CW, et al. Identification of a novel class of insect glutathione S-transferases involved in resistance to DDT in the malaria vector Anopheles gambiae. Biochem J. 2001;359(Pt 2):295–304.View ArticlePubMed CentralPubMedGoogle Scholar
- Enayati AA, Vontas JG, Small GJ, McCarroll L, Hemingway J. Quantification of pyrethroid insecticides from treated bednets using a mosquito recombinant glutathione S-transferase. Med Vet Entomol. 2001;15(1):58–63.View ArticlePubMedGoogle Scholar
- Chelvanayagama G, Parker MW, Board PG. Fly fishing for GSTs: A unified nomenclature for mammalian and insect glutathione transferases. Chem-Biol Interact. 2001;133:256–60.Google Scholar
- Low WY, Ng HL, Morton CJ, Parker MW, Batterham P, Robin C. Molecular evolution of glutathione S-transferases in the genus Drosophila. Genetics. 2007;177(3):1363–75.View ArticlePubMed CentralPubMedGoogle Scholar
- Sawicki R, Singh SP, Mondal AK, Benes H, Zimniak P. 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. 2003;370(Pt 2):661–9.View ArticlePubMed CentralPubMedGoogle Scholar
- Ketterman AJ, Saisawang C, Wongsantichon J. Insect glutathione transferases. Drug Metab Rev. 2011;43(2):253–65.View ArticlePubMedGoogle Scholar
- Abel EL, Bammler TK, Eaton DL. Biotransformation of methyl parathion by glutathione S-transferases. Toxicol Sci. 2004;79(2):224–32.View ArticlePubMedGoogle Scholar
- Huang HS, Hu NT, Yao YE, Wu CY, Chiang SW, Sun CN. Molecular cloning and heterologous expression of a glutathione S-transferase involved in insecticide resistance from the diamondback moth. Plutella xylostella Insect Biochem Mol Bio. 1998;28(9):651–8.View ArticleGoogle Scholar
- Wei SH, Clark AG, Syvanen M. 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 Bio. 2001;31(12):1145–53.View ArticleGoogle Scholar
- Hemingway J, Miyamoto J, Herath PRJ. A possible novel link between organophosphorus and DDT insecticide resistance genes in Anopheles: Supporting evidence from fenitrothion metabolism studies. Pestic Biochem Physiol. 1991;39(1):49–56.View ArticleGoogle Scholar
- Yamamoto K, Shigeoka Y, Aso Y, Banno Y, Kimura M, Nakashima T. Molecular and biochemical characterization of a Zeta-class glutathione S-transferase of the silkmoth. Pestic Biochem Physiol. 2009;94(1):30–5.View ArticleGoogle Scholar
- Yamamoto K, Nagaoka S, Banno Y, Aso Y. Biochemical properties of an omega-class glutathione S-transferase of the silkmoth, Bombyx mori. Comp Biochem Physiol C: Toxicol Pharmacol. 2009;149(4):461–7.Google Scholar
- Rogers ME, Jani MK, Vogt RG. An olfactory-specific glutathione-S-transferase in the sphinx moth Manduca sexta. J Exp Biol. 1999;202(Pt 12):1625–37.PubMedGoogle Scholar
- You M, Yue Z, He W, Yang X, Yang G, Xie M, et al. A heterozygous moth genome provides insights into herbivory and detoxification. Nat Genet. 2013;45(2):220–5.View ArticlePubMedGoogle Scholar
- Chen X, Zhang YL. Identification and characterisation of multiple glutathione S-transferase genes from the diamondback moth, Plutella xylostella. Pest Manag Sci. 2014Google Scholar
- Jouraku A, Yamamoto K, Kuwazaki S, Urio M, Suetsugu Y, Narukawa J, et al. KONAGAbase: a genomic and transcriptomic database for the diamondback moth, Plutella xylostella. BMC Genomics. 2013;14:464.View ArticlePubMed CentralPubMedGoogle Scholar
- He W, You M, Vasseur L, Yang G, Xie M, Cui K, et al. Developmental and insecticide-resistant insights from the de novo assembled transcriptome of the diamondback moth. Plutella xylostella Genomics. 2012;99(3):169–77.View ArticleGoogle Scholar
- Tang W, Yu L, He W, Yang G, Ke F, Baxter SW, et al. DBM-DB: the diamondback moth genome database. Database (Oxford). 2014;2014:bat087.View ArticleGoogle Scholar
- Yu Q, Lu C, Li B, Fang S, Zuo W, Dai F, et al. Identification, genomic organization and expression pattern of glutathione S-transferase in the silkworm, Bombyx mori. Insect Biochem Mol Bio. 2008;38(12):1158–64.View ArticleGoogle Scholar
- Ding Y, Ortelli F, Rossiter LC, Hemingway J, Ranson H. The Anopheles gambiae glutathione transferase supergene family: annotation, phylogeny and expression profiles. BMC Genomics. 2003;4(1):35.View ArticlePubMed CentralPubMedGoogle Scholar
- Friedman R. Genomic organization of the glutathione S-transferase family in insects. Mol Phylogen Evol. 2011;61(3):924–32.View ArticleGoogle Scholar
- Oakeshott JG, Johnson RM, Berenbaum MR, Ranson H, Cristino AS, Claudianos C. Metabolic enzymes associated with xenobiotic and chemosensory responses in Nasonia vitripennis. Insect Mol Biol. 2010;19:147–63.View ArticlePubMedGoogle Scholar
- Nair PMG, Choi J. Identification, characterization and expression profiles of Chironomus riparius glutathione S-transferase (GST) genes in response to cadmium and silver nanoparticles exposure. Aquat Toxicol. 2011;101(3–4):550–60.View ArticlePubMedGoogle Scholar
- Deng H, Huang Y, Feng Q, Zheng S. Two epsilon glutathione S-transferase cDNAs from the common cutworm, Spodoptera litura: Characterization and developmental and induced expression by insecticides. J Insect Physiol. 2009;55(12):1174–83.View ArticlePubMedGoogle Scholar
- Lumjuan N, Rajatileka S, Changsom D, Wicheer J, Leelapat P, Prapanthadara L-a, et al. The role of the Aedes aegypti Epsilon glutathione transferases in conferring resistance to DDT and pyrethroid insecticides. Insect Biochem Mol Bio. 2011;41(3):203–9.View ArticleGoogle Scholar
- Tamura K, Peterson D, Peterson N, Stecher G, Nei M, Kumar S. MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol Biol Evol. 2011;28(10):2731–9.View ArticlePubMed CentralPubMedGoogle Scholar
- Vontas JG, Small GJ, Nikou DC, Ranson H, Hemingway J. Purification, molecular cloning and heterologous expression of a glutathione S-transferase involved in insecticide resistance from the rice brown planthopper, Nilaparvata lugens. Biochem J. 2002;362(Pt 2):329–37.View ArticlePubMed CentralPubMedGoogle Scholar
- Wang JY, McCommas S, Syvanen M. Molecular cloning of a glutathione S-transferase overproduced in an insecticide-resistant strain of the housefly (Musca domestica). Mol Gen Genet. 1991;227(2):260–6.View ArticlePubMedGoogle Scholar
- Shi H, Pei L, Gu S, Zhu S, Wang Y, Zhang Y, et al. Glutathione S-transferase (GST) genes in the red flour beetle, Tribolium castaneum, and comparative analysis with five additional insects. Genomics. 2012;100(5):327–35.View ArticlePubMedGoogle Scholar
- Savard J, Tautz D, Richards S, Weinstock GM, Gibbs RA, Werren JH, et al. Phylogenomic analysis reveals bees and wasps (Hymenoptera) at the base of the radiation of Holometabolous insects. Genome Res. 2006;16(11):1334–8.View ArticlePubMed CentralPubMedGoogle Scholar
- Haddrill PR, Charlesworth B, Halligan DL, Andolfatto P. Patterns of intron sequence evolution in Drosophila are dependent upon length and GC content. Genome Biol. 2005;6(8):R67.View ArticlePubMed CentralPubMedGoogle Scholar
- Atkinson HJ, Babbitt PC. Glutathione transferases are structural and functional outliers in the thioredoxin fold. Biochemistry. 2009;48(46):11108–16.View ArticlePubMed CentralPubMedGoogle Scholar
- Armstrong RN. Structure, catalytic mechanism, and evolution of the glutathione transferases. Chem Res Toxicol. 1997;10(1):2–18.View ArticlePubMedGoogle Scholar
- Zhou WW, Liang QM, Xu Y, Gurr GM, Bao YY, Zhou XP, et al. Genomic insights into the gutathione S-transferase gene family of two rice planthoppers, Nilaparvata lugens (Stal) and Sogatella furcifera (Horvath) (Hemiptera: Delphacidae). PLoS One. 2013;8(2):e56604.View ArticlePubMed CentralPubMedGoogle Scholar
- Wongtrakul J, Pongjaroenkit S, Leelapat P, Nachaiwieng W, Prapanthadara LA, Ketterman AJ. Expression and characterization of three new glutathione transferases, an epsilon (AcGSTE2-2), omega (AcGSTO1-1), and theta (AcGSTT1-1) from Anopheles cracens (Diptera: Culicidae), a major Thai malaria vector. J Med Entomol. 2010;47(2):162–71.View ArticlePubMedGoogle Scholar
- Li X, Schuler MA, Berenbaum MR. Molecular mechanisms of metabolic resistance to synthetic and natural xenobiotics. Annu Rev Entomol. 2007;52:231–53.View ArticlePubMedGoogle Scholar
- Sun XQ, Zhang MX, Yu JY, Jin Y, Ling B, Du JP, et al. Glutathione S-transferase of brown planthoppers (Nilaparvata lugens) is essential for their adaptation to gramine-containing host plants. PLoS One. 2013;8(5):e64026.View ArticlePubMed CentralPubMedGoogle Scholar
- Zhang Y, Yan H, Lu W, Li Y, Guo X, Xu B. A novel Omega-class glutathione S-transferase gene in Apis cerana cerana: molecular characterisation of GSTO2 and its protective effects in oxidative stress. Cell Stress Chaperones. 2013;18(4):503–16.View ArticlePubMed CentralPubMedGoogle Scholar
- Lumjuan N, McCarroll L, Prapanthadara LA, Hemingway J, Ranson H. Elevated activity of an Epsilon class glutathione transferase confers DDT resistance in the dengue vector, Aedes aegypti. Insect Biochem Mol Bio. 2005;35(8):861–71.View ArticleGoogle Scholar
- Yamamoto K, Ichinose H, Aso Y, Banno Y, Kimura M, Nakashima T. Molecular characterization of an insecticide-induced novel glutathione transferase in silkworm. Biochim Biophys Acta. 2011;1810(4):420–6.View ArticlePubMedGoogle Scholar
- Samra AI, Kamita SG, Yao H-W, Cornel AJ, Hammock BD. Cloning and characterization of two glutathione S-transferases from pyrethroid-resistant Culex pipiens. Pest Manage Sci. 2012;68(5):764–72.View ArticleGoogle Scholar
- Dow JA. Insights into the Malpighian tubule from functional genomics. J Exp Biol. 2009;212(Pt 3):435–45.View ArticlePubMedGoogle Scholar
- Yu QY, Lu C, Li WL, Xiang ZH, Zhang Z. Annotation and expression of carboxylesterases in the silkworm, Bombyx mori. BMC Genomics. 2009;10:553.View ArticlePubMed CentralPubMedGoogle Scholar
- Megy K, Emrich SJ, Lawson D, Campbell D, Dialynas E, Hughes DS, et al. VectorBase: improvements to a bioinformatics resource for invertebrate vector genomics. Nucleic Acids Res. 2012;40(Database issue):D729–34.View ArticlePubMed CentralPubMedGoogle Scholar
- Marygold SJ, Leyland PC, Seal RL, Goodman JL, Thurmond J, Strelets VB, et al. FlyBase: improvements to the bibliography. Nucleic Acids Res. 2013;41(D1):D751–7.View ArticlePubMed CentralPubMedGoogle Scholar
- Legeai F, Shigenobu S, Gauthier JP, Colbourne J, Rispe C, Collin O, et al. AphidBase: a centralized bioinformatic resource for annotation of the pea aphid genome. Insect Mol Biol. 2010;19 Suppl 2:5–12.View ArticlePubMedGoogle Scholar
- Kim HS, Murphy T, Xia J, Caragea D, Park Y, Beeman RW, et al. BeetleBase in 2010: revisions to provide comprehensive genomic information for Tribolium castaneum. Nucleic Acids Res. 2010;38(Database issue):D437–42.View ArticlePubMed CentralPubMedGoogle Scholar
- Munoz-Torres MC, Reese JT, Childers CP, Bennett AK, Sundaram JP, Childs KL, et al. Hymenoptera Genome Database: integrated community resources for insect species of the order Hymenoptera. Nucleic Acids Res. 2011;39(Database issue):D658–62.View ArticlePubMed CentralPubMedGoogle Scholar
- Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, et al. Clustal W and Clustal X version 2.0. Bioinformatics. 2007;23(21):2947–8.View ArticlePubMedGoogle Scholar
- Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4(4):406–25.PubMedGoogle Scholar
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly credited. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.