Developmental and tissue-specific expression of CHS1 from Plutella xylostella and its response to chlorfluazuron

www.elsevier.com/locate/ypest

Pesticide Biochemistry and Physiology 89 (2007) 20–30

PESTICIDEBiochemistry & Physiology

Developmental and tissue-specific expression of CHS1 fromPlutella xylostella and its response to chlorfluazuron

Muhammad Ashfaq, Shoji Sonoda *, Hisaaki Tsumuki

Research Institute for Bioresources, Okayama University, Kurashiki, Okayama 710-0046, Japan

Received 21 December 2006; accepted 14 February 2007Available online 24 February 2007

Abstract

The cDNA of chitin synthase-1 gene of Plutella xylostella (PxCHS1) was characterized and expression patterns of the two splice vari-ants PxCHS1A and PxCHS1B were investigated in various developmental stages and in major body tissues by RT-PCR and real-timequantitative PCR (qPCR). The PxCHS1 cDNA was 5461 bp in length with an open reading frame of 4701 bp that encoded a putativeprotein of 1567 amino acids with predicted molecular mass of 179 kDa. The two splice variants were expressed from the mutually exclu-sive exons which were same in size (177 bp) but showed only 66% identity at the nucleotide level. Both splice variants were expressed inall developmental stages. The qPCR data suggested an uneven expression of the two variants in the body where expression of PxCHS1A

was 3.7-fold higher than that of PxCHS1B. The expression of PxCHS1A was 1.5-fold higher in the head than the body whereas in caseof PxCHS1B the difference between head and body was 6.3-fold. Chlorfluazuron did not change the expression of PxCHS1 in larvae.� 2007 Elsevier Inc. All rights reserved.

Keywords: Chitin synthase; Chlorfluazuron; Splice variants; Gene expression; Plutella xylostella

1. Introduction

Insect cuticle is composed of micro fibers of chitin sur-rounded by a matrix of protein [1]. Chitin is a crystallinepolymer of b-1,4-linked N-acetyl-D-glucosamine and inaddition to cuticle it is present in insect gut and trachealtubes. Chitin fibers are synthesized by the enzyme chitinsynthase (CHS), a glycosyltransferase found in fungi, nem-atodes and arthropods [2–4]. The disruption of chitin syn-thase function can impair the formation, sclerotization andpigmentation process of cuticle as evidenced in a study onDrosophila melanogaster [5].

Initially, CHSs were identified from fungi, where at leastsix different genes have been reported from various fungalspecies [3]. Ibrahim et al. [6] had isolated and characterizedthe first insect cDNA of the gene encoding CHS (the genelater classified as CHS2) [7] from peritrophic membrane ofAedes aegypti. From another dipteran, Lucilia cuprina a

0048-3575/$ - see front matter � 2007 Elsevier Inc. All rights reserved.

doi:10.1016/j.pestbp.2007.02.004

* Corresponding author. Fax: +81 86 434 1249.E-mail address: [email protected] (S. Sonoda).

different CHS was characterized and classified as CHS1

[8]. To date two chitin synthase genes, CHS1 and CHS2

have been reported from insects [4,9] and one or both typesof genes have been characterized from at least three insectorders, Diptera, Lepidoptera and Coleoptera [6–12]. Theprevious data have indicated that CHS2 is relatively smal-ler in size than CHS1 although this difference varies in dif-ferent insect species [7,13].

The differential expression of CHSs has been reportedwhere more than one gene are present. There is evidencefrom nematodes that one CHS is expressed and used ineggshell production whereas the other in the cells to formpharynx [2]. Observations about different CHSs expressedin different tissues have been made in fungi too [14]. Ininsects, CHS2 is expressed in the gut where the chitin is uti-lized in the development of lining of the alimentary canal,whereas CHS1 is mainly expressed by the epidermal tissuesand tracheae, where chitin serves as a building block of thecuticle [8,10,12,15,16]. Both genes share a significant num-ber of nucleotides, particularly in the region coding for thecatalytic domain of the putative proteins.

mailto:[email protected]

M. Ashfaq et al. / Pesticide Biochemistry and Physiology 89 (2007) 20–30 21

The CHS1 encodes a large protein comprising a highlyconserved catalytic domain and two relatively less con-served flanking domains. The N-terminal and C-terminaldomains contain 15–18 transmembrane segments and thusreferred to as transmembrane domains [8], making the pro-tein transmembrane in nature. At least seven of the trans-membrane helices are present in C-terminal domain. Thetotal mass of the encoded protein has been reported around170 kDa with slightly acidic isoelectric points [4,17]. Theprotein contains two consensus sequences EDR andQRRRW, also referred to as signature sequences of CHSsand are reported from all the insect CHS studies performedso far [4].

The usage of alternate exon in CHS1 has been reportedfrom several insect species (Tribolium castaneum, D. mela-

nogaster and Anopheles gambiae) [4,11] while there is noevidence of this phenomenon in case of CHS2 [9]. Theseexons have been referred to as ‘A’ or ‘B’ [11] and theirexpression produce two splice variants (isoforms). Thesesplice variants differ in amino acid sequences and are differ-entially expressed in the epidermis and tracheae [4]. Theexpression of these exons is differentially regulated duringdevelopment and between epidermis and tracheae ofManduca sexta, with a relatively higher expression ofexon-A in epidermis and that of exon-B in tracheae [9,12].

Benzoylphenylureas (BPUs) including diflubenzuronand chlorfluazuron are a kind of insect growth regulators(IGR) and disrupt chitin synthesis [18,19]. Ishaaya andCasida [20] noted an increased chitinase activity in housefly fed on diflubenzuron-treated diet, but Deul [21] foundno effects on chitinase when repeated the experiments withPieris brassicae (L.), a lepidopteran. Some researchers havesuggested that BPUs interfere with cuticle synthesis byinhibiting the CHS [22] but others found no effect of BPUson CHS [19,23,24]. In a more recent study, Zhang and Zhu[17] reported an up-regulation of CHS1 expression inAnopheles quadrimaculatus, as a result of diflubenzurontreatment. However, in light of the research published sofar, it can be assumed that the nature of BPU effect onCHS is not conclusive.

Plutella xylostella is an important lepidopteran insectand is a major pest on several agricultural crops. Under-standing the regulation of CHS, a basic enzyme for chitindevelopment can be helpful in development and use ofnew strategies to control this insect. The purpose of thecurrent study was to sequence the full length cDNA codingfor CHS1 (PxCHS1), analyze the structure of putative pro-tein and evaluate the expression of the gene in developmen-tal stages and in body tissues. Further, to investigate thechlorfluazuron effects on PxCHS1 expression.

2. Materials and methods

2.1. Chlorfluazuron treatment

Test insects were obtained from Sumitomo ChemicalCo. Ltd., Osaka, Japan, in 2004 and have been maintained

in a controlled environment on radish seedlings at 25 �Cand 16:8 (L:D) cycle. Chlorfluazuron (Atabron 5% suspen-sion concentrate) was purchased from Ishihara Sangyo,Osaka, Japan. The LD50 dose of the fourth instar larvaeof P. xylostella to chlorfluazuron was estimated to be1.8 ppm [25]. The third instar larvae were used in the pres-ent study to evaluate the effect on PxCHS1 expression.Radish seedlings were dipped for 30 s in the LD50 concen-tration of chlorfluazuron prepared in distilled H2O (dH2O),air-dried for 30 min and provided to the larvae in plasticcups. The control larvae were provided with seedlingsdipped in dH2O only. Total RNA was extracted after24 h for RT-PCR analysis of PxCHS1 mRNA in treatedand control larvae.

2.2. RNA and DNA extractions

Total RNA was extracted from 24–28 h old eggs, larvae(first, second, third and fourth instar), pupae and adults ofP. xylostella. Third instar larval body parts, head, gut andthe remaining body (body) were separated. To do that thelarvae were decapitated, gut was pulled out and the remain-ing body was left as such. Due to small size of the larvae itwas not feasible to remove the attached trachea off the gut.The procedures were repeated three times to replicate theexpression analysis.

For the RNA extraction, insects or the insect body partswere ground using a pot and pestle on ice by mixing inRNA extraction buffer (Tris–HCl, 50 mM; pH 8.5; EDTA,10 mM; NaCl, 100 mM; SDS 2%). RNA was extractedtwice with equal volumes of phenol: chloroform: isoamylal-cohol (25:24:1) and once with chloroform: isoamylalcohol(24:1) and precipitated using 99.5% ethanol in the presenceof 3 M sodium acetate. RNA pellet was washed with 70%ethanol, re-suspended in dH2O treated with DEPC andstored at �80 �C until used.

2.3. cDNA synthesis

Oligo(dT)-primed cDNA was synthesized from totalRNA as described previously [26] or poly-A mRNA waspurified from total RNA of pupae by using an mRNApurification kit (Takara Bio Inc., Otsu, Japan) accordingto the supplier’s instructions. Subsequent synthesis ofdouble-stranded, adaptor-ligated cDNA (library) was per-formed as described in the Marathon cDNA amplificationkit (BD Biosciences, Palo Alto, CA, USA). The adaptor-ligated cDNA was diluted to 1:100 in Tricine–EDTA bufferbefore using in a PCR reaction.

2.4. Polymerase chain reaction (PCR) amplification of

CHS1 sequences

Information on PCR primers used and the cDNA frag-ments amplified with the corresponding PCR conditionshave been stated in Tables 1 and 2, respectively, and overlap-ping fragments amplified are shown in Fig. 1. Nucleotide and

Table 1PCR primers used in this study

Primer Corresponding nucleotides in Fig. 2 Sequence (50–3 0)

CS-1 (F) 2448–2474 GGCTCTGGTCCTATGGTGTGGTATCA (GSGPMVWYQ)CS-2 (R) 2925–2953 GCCACRAAAGCACCCACCAACATAAGGAA

(FLMLVGAFVA)CS-3 (F) 2888–2912 GTTGATTGGCGGTACGATCTTGGGCCS-4 (F) 2977–3002 CTTCATTCGAATACAACCTCTACCCGCS-5 (F) 1839–1855 TGCGCKACTATGTGGCA (CATMWH)CS-6 (R) 2561–2586 CCATGAGAGCCTTCCCTCTGAAGAGCCS-7 (F) 927–947 TCDCTSGGMTGGTGGGARAAY (SLGWWEN)CS-8 (R) 1861–1884 GGAACTCCATCATCTCGTCCTTCGCS-9 (R) 960–983 CTTGATTATACCGATTGGACTCTGCS-10 (F) 3901–3920 AGCAGGCCCGTATCTCGCACCS-11 (R) 4063–4082 CTCTTGCGTCACCTCATCACS-12 (F) 3902–3920 GCAGGCTCGCATTGCAGCTCS-13 (R) 4063–4082 CTCGCCTGTCTCCTCGATGCS-14 (F) 1593–1616 GCCGAGCGTCTGGCGTCCACGGAGCS-15 (R) 1837–1859 CTCGTGCCACATCGTCGCGCATGM4 (adaptor primer) GTTTTCCCAGTCACGACAP1 (adaptor primer, BD biosciences) CCATCCTAATACGACTCACTATAGGGC

(F), forward primer; (R), reverse primer; conserved amino acids in parenthesis indicate the degenerate primers.

Table 2Primer combinations used to amplify the cDNA of PxCHS1

cDNA fragment amplified (size) (refer Fig. 1) Primer combination PCR conditions

I (507 bp) CS-1 + CS-2 94 �C, 2 min; 40 cycles of 94 �C, 30 s/50 �C, 1 min/72 �C, 1 min; 72 �C, 5 minII (1019 bp); 3 0 RACE, partial CS-3 + M4 94 �C, 2 min; 30 cycles of 94 �C, 30 s/55 �C, 1 min/72 �C, 2 min; 72 �C, 5 minIII (2483 bp); 3 0 RACE CS-4 + AP1 94 �C, 30 s; 35 cycles of 94 �C, 5 s/68 �C, 4 minIV (748 bp) CS-5 + CS-6 94 �C, 2 min; 30 cycles of 94 �C, 30 s/55 �C, 1 min/72 �C, 2 min; 72 �C, 5 minV (958 bp) CS-7 + CS-8 94 �C, 2 min; 30 cycles of 94 �C, 30 s/55 �C, 1 min/72 �C, 2 min; 72 �C, 5 minVI (983 bp); 5 0 RACE CS-9 + AP1 95 �C, 1 min; 35 cycles of 95 �C, 30 s/68 �C, 1 min; 68 �C, 1 min

Fig. 1. A schematic diagram of the strategy to amplify the PxCHS1 cDNA. Upper rectangle represents the full cDNA. Lower rectangles stand for each ofthe PCR fragment mentioned in Table 2. A 507 bp cDNA fragment was initially amplified and the obtained sequence was used to develop primers foramplification of rest of the cDNA fragments. The amplified fragment sequences were put together and overlapping sequence were removed to obtain theconsensus cDNA sequence.

22 M. Ashfaq et al. / Pesticide Biochemistry and Physiology 89 (2007) 20–30

deduced amino acid sequences of CHS1 gene from severalinsects available in the DNA Data Bank of Japan (DDBJ)were aligned. The conserved regions from the alignment weredetermined and degenerate primers were developed (Table1). A 507 bp fragment (I) of P. xylostella cDNA was ampli-fied in a PCR reaction using the primer pair, forward primerCS-1 and reverse primer CS-2 following PCR conditionsoutlined in Table 2. The PCR fragments were analyzed onagarose gels, purified (Qiagen Inc., Valencia, CA, USA),and subsequently cloned into pGEM-T Easy vector (Pro-mega Corp., Madison, WI, USA). The nucleotide sequencewas determined using BigDye� Terminator Cycle Sequenc-ing Kit (Applied Biosystems,Warrington, UK) with M13

forward and reverse primers and subjecting the samples toa DNA sequencer (Applied Biosystems, 3100 Avant GeneticAnalyzer).

2.5. Rapid amplification of cDNA ends (RACE)

Gene specific primers (GSPs) were developed usingsequence of the first amplified cDNA fragment or the over-lapping sequences of the subsequent fragments obtained.The homology of each amplified fragment with other insectCHS1 sequences was confirmed before developing the newGSPs. The 3 0 RACE was performed with the primer pairCS-3 and M4 using oligo(dT)-primed cDNA following


PCR conditions described (Table 2) which produced a1019 bp PCR product (II). The fragment was cloned andsequenced (which turned out to be a partial 3 0 end frag-ment) and GSPs were developed. A second 3 0 RACEPCR was performed with the primer pair CS-4 and AP1using cDNA synthesized from mRNA (the cDNA was usedfor all the subsequent amplifications too) following PCRconditions described (Table 2). The PCR reaction con-tained 2.5 ll of 100-fold diluted cDNA, 0.2 lM of each pri-mer in a primer pair, 0.2 lM dNTP mix, 1· cDNA PCRreaction buffer and 1· Advantage 2 Polymerase Mix (BDBiosciences) in a final volume of 50 ll. The 2483 bp PCRproduct obtained (III) was cloned and sequenced andhomology with other insect CHS1 genes was confirmedby DDBJ Blast Search.

To obtain the full sequence towards 5 0 end, fragment IV(748 bp) and fragment V (958 bp) (Fig. 1) were amplifiedwith the primer pairs CS-5/CS-6 and CS-7/CS-8, respec-tively (Table 2). The 5 0 RACE was performed with the pri-mer pair CS-9/AP1 which produced a 983 bp PCR product(Table 2). Cloning and sequencing of the product con-firmed the complete sequence of 5 0 end.

2.6. Identification of alternate exons

The amplification and sequencing of the 3 0 end of thePxCHS1 had revealed that a 177 bp sequence segment(nucleotide position 3906–4082) is variable and fabricatestwo sequence fragments. Nucleotide and deduced aminoacid alignments showed that these sequences correspondto the exon-8A or exon-8B sequences of CHS1 reportedpreviously [11]. We designed PCR primers from the 3 0

and 5 0 ends of this variable segment and specifically ampli-fied and sequenced each exon. At the PCR conditions,2 min at 94 �C; 30 s at 94 �C, 30 s at 60 �C, 1 min at72 �C (30 cycles); 5 min at 72 �C, the primer pairs CS-10/CS-11 and CS-12/CS-13 (Table 1) specifically amplified177 bp PCR products of exon-A and exon-B, respectively.As the amplified PCR products were of the same size, weconfirmed the specificity of amplification of the respectivePCR products by cloning and sequencing each exon.

2.7. Computer and phylogenetic analyses

Analyses of the nucleotide and deduced amino acidsequences were performed using Genetyx Ver. 8.0 (SoftwareDevelopment, Tokyo, Japan). Blast Search tool of the DDBJwas used to search homology for the cloned sequences.Transmembrane helices and N-glycosylation sites in theputative protein sequence were predicted using tools pro-vided by the ExPASy Service (http://www.expasy.org/).The deduced amino acid sequences of insect CHS1 availablein the GenBank were aligned using ClustalW (ver 1.83) [27].For phylogenetic analysis a dendrogram was developed fromClustalW alignment of available CHS sequences using njplotsoftware. Sequences from the following organism specieswere used in the alignment or phylogenetic analysis

(accession numbers follow in parenthesis). A. gambiae,AgCHS1 (XM_321337); L. cuprina, LcCHS1 (AF221067);A. quadrimaculatus, AqCHS1 (DQ415985); D. melanogaster,DmCHS1 (NM_079509); Spodoptera exigua, SeCHS1

(DQ062115); M. sexta, MsCHS1 (AY062175); T. castane-

um, TcCHS1-A (AY291475); T. castaneum, TcCHS1-B(AY291476); A. aegypti, AeCHS2 (AF223577.2); A. gam-

biae, AgCHS2 (AF223577); Spodoptera frugiperda, SfCHS2

(AY525599); Ostrinia furnacalis, OfCHS2 (DQ294306); M.

sexta, MsCHS2 (AY821560); T. castaneum, TcCHS2

(AY291477); D. melanogaster, DmCHS2 (NM_079485);Exophiala dermatitidis, EdCHS1 (AF054503); Caenorhabdi-

tis elegans, CeCHS1 (AY874871); C. elegans, CeCHS2

(AY874872).The cDNA and deduced amino acid sequences of

PxCHS1 were subjected to blast searches of the DDBJ todetermine the sequence homology with other homologousgenes.

2.8. PxCHS1 expression analysis by reverse transcription

PCR (RT-PCR)

Oligo(dT)-primed cDNA was prepared from 1 lg oftotal RNA treated with DNase I (Toyobo, Osaka,Japan) which had been derived from various P. xylostella

life stages or chlorfluazuron-treated larvae. Equal quan-tity of the synthesized cDNA was subjected to PCRfor amplification of the cDNA using primers developedfrom the respective sequences of each exon (Fig. 4) forexon-specific gene expression. The primer pairs CS-10/CS-11 and CS-12/CS-13 (Table 1) specifically amplifiedexon-A or exon-B, respectively, producing a 177 bpPCR product at the PCR conditions mentioned above.RT-PCR for PxCHS1 expression in response to chlorflu-azuron was performed with primer pair CS-14/CS-15developed from sequence region common for both theisoforms. Comparison of the relative quantity of totalcDNA in each of the samples was made using a PCRapproach specific for cDNA coding for b-actin proteinwhich produced a 586 bp PCR product with primer com-bination, forward 5 0-GCTCCCGAGGAGCACCCCGTGCTGC-3 0 and reverse 5 0-CCTTACGGATGTCCACGTCGCACTTC-3 0 at the same PCR conditions mentionedabove. The PCR products were electrophoresed into1.2% agarose gels, ethidium bromide stained andvisualized using Dolphin-Chemi (Wealtec Corp., Sparks,NV, USA).

2.9. Real-time quantitative PCR (qPCR) analysis of exon-A

and exon-B mRNA in major tissues

The mRNA levels of PxCHS1A and PxCHS1B in thehead, gut and body obtained from the third instar larvaewere evaluated by real-time qPCR using relative quantifi-cation function. Three replications of qPCR were per-formed using equal amounts of oligo(dT)-primed cDNAas template which had been synthesized from each body

http://www.expasy.org/


part as mentioned above. Primer combinations mentionedabove in the RT-PCR analysis were used to target therespective exons of each isoform. The 50 ll PCR reactions,cDNA template were used in contained 25 ll of PowerSYBR Green PCR Master Mix (Applied Biosystems) and0.2 lM of each primer. Relative quantification was con-ducted in Applied Biosystems 7500 Real-Time PCR system(Applied Biosystems). The P. xylostella b-actin (control)was amplified to normalize the threshold cycle (Ct) valuefor the target PxCHS1-exon amplifications between the tis-sues. The optimized real-time qPCR was used for bothb-actin and isoform-A/B of PxCHS1, which consisted ofinitial step at 95 �C for 2 min followed by 40 cycles of95 �C for 30 s, 60 �C for 30 s and 72 �C for 1 min. After-wards, amplification specificity was confirmed by obtainingthe dissociation curve, and also running the PCR productson agarose gels. The Ct values of the exons were normali-zed for specific tissues by adding or subtracting the devia-tion proportion of the Ct values of the b-actin in thesamples.

3. Results

3.1. cDNA and deduced amino acid sequences of PxCHS1

We amplified overlapping cDNA fragments by PCR toobtain the full length PxCHS1 cDNA sequence. The strat-egy to obtain the full length cDNA sequence has been out-lined in Fig. 1. The complete cDNA sequence of PxCHS1

is revealed in Fig. 2. Total length of the cloned cDNA was5461 bp with 4701 bp of open reading frame (ORF) begin-ning with the methionine start codon ATG and ending withtranslation stop codon TAA at nucleotide positions 135and 4836, respectively (Fig. 2) (GenBank/EMBL/DDBJAccession No. AB271784). The ORF translates to 1567amino acids producing a putative protein of 179 kDa witha 6.5 isoelectric point. The polyadenylation signal(AATAAA) was detected 25 nucleotides upstream fromthe poly-A tail. The 5 0- and 3 0-untranslated regions (UTRs)comprised of 134 and 623 bp, respectively. The 5 0-UTR ofthe start codon of the cDNA partially coincided with theKozak consensus sequence (GCC (G/A) CCAUGG) [28].The sequence analysis of the cDNA revealed the presenceof two splice variants of the mRNA of PxCHS1 corre-sponding to nucleotide positions 3906–4082. The splicevariants are hereafter referred to as PxCHS1A andPxCHS1B. Nucleotide sequence shown in Fig. 2 representsPxCHS1A. PxCHS1A and PxCHS1B were, respectively,supposed to be expressed by exon-A and exon-B, the alter-nate exons which have previously been reported from sev-eral insect species (Fig. 3). Nucleotide sequence of exon-Bfrom P. xylostella was deposited in GenBank/EMBL/DDBJ with Accession No. AB281490.

Alignment of the deduced amino acid sequence of 1567amino acid residues with those of other CHSs suggestedthat the N-terminus sequence was complete (Fig. 4). Theuse of TMHMM Server v. 2.0 [29] predicted 17 transmem-

brane helices in the amino acid sequence of the putativeprotein spanning on both sides of the catalytic domain.The CHS signature sequences, EDR and QRRRW werepresent in the catalytic domain (Fig. 2). Amino acidsequences for possible glycosylation sites (Asn-X-Ser andAsn-X-Thr) were found in five locations (Fig. 2). Analysisof deduced amino acid sequence using Signal 3.0 Server[30] did not predict a signal peptide, suggesting that theputative protein is non-secretory in nature.

3.2. Sequence alignments and phylogenetic analysis

The full length alignment of deduced amino acidsequence of PxCHS1A against other homologoussequences from insect species revealed a significant homol-ogy throughout the sequence in general and the catalyticdomain in particular, where the overall identity was 82%(Fig. 4). Amino acid and nucleotide sequence alignmentsagainst two other lepidopterans showed an identity of91% and 89%, and 76% and 75% with those of S. exigua

and M. sexta, respectively. The two PxCHS1 alternativelyexpressed exons, exon-A and exon-B, showed a sequenceidentity of 66% and 73% at nucleotide and amino acid lev-els, respectively (Fig. 3). On the other hand amino acidsequence identity of PxCHS1 exon-A with exon-8A ofM. sexta and T. castaneum was 90% and 83%, respectively.The identity of PxCHS1 exon-B with exon-8B of M. sexta

and T. castaneum was 93% and 83%, respectively. A den-drogram was developed from the alignment of insect aminoacid sequences of CHS1 and CHS2, and three CHSs repre-senting non-insect species. The insect CHS1 and CHS2were clearly separated into two groups, with PxCHS1

along with other insect CHS1 and much closer to the twolepidopterans (Fig. 5).

3.3. Developmental and tissue-specific expression pattern of

PxCHS1A and PxCHS1B

The levels of PxCHS1A and PxCHS1B in various devel-opmental stages of the insect were examined by RT-PCR.The results indicated that mRNA for both isoforms wasdetectable from egg through adult stage; however expres-sion was significantly lower in the fourth instar larvae(Fig. 6).

To measure the mRNA levels of the two isoforms in thehead, gut and the body we analyzed cDNA samples pre-pared from the total RNA of the respective tissues in areal-time qPCR (Fig. 7). The Ct values revealed a 1.5-foldhigher mRNA level of PxCHS1A in the head than in thebody (0.82 ± 0.1 vs. 0.56 ± 0.22). The mRNA level ofPxCHS1B was 6.3-fold higher in the head than in the body(0.94 ± 0.15 vs. 0.15 ± 0.02). Comparison between the twoisoforms indicated that the expression was almost equiva-lent in the head (PxCHS1A, 0.82 ± 0.1 vs. PxCHS1B,0.94 ± 0.15) while in the body mRNA level of PxCHS1Awas 3.7-fold higher than that of PxCHS1B (0.56 ± 0.22vs. 0.15 ± 0.02). There was little but equal level of


transcription of both isoforms in the gut (PxCHS1A,0.09 ± 0.02 and PxCHS1B, 0.1 ± 0.04) (Fig. 7).

3.4. Chlorfluazuron effect on PxCHS1 expression

To evaluate the effects of a BPU on CHS1 expression inP. xylostella, we extracted total RNA from the larvae fedon chlorfluazuron-treated radish seedlings and used forcDNA synthesis for RT-PCR analysis. Chlorfluazurondid not cause any changes in gene expression level ofPxCHS1 in the treated P. xylostella larvae as detected byRT-PCR (data not shown).

Fig. 2. Nucleotide and deduced amino-acid sequences of PxCHS1. Stop codtransmembrane segments predicted by the computer program TMHMM Serv‘signature’ sequence is fully shaded. Potential N-glycosylation sites are boxed.

4. Discussion

CHS is a key enzyme involved in the process of chitinsynthesis which has been studied more rigorously in fun-gal and nematode species than insects [3]. So far, twotypes of CHS genes have been reported from insects,CHS1 and CHS2 [4] which share a significant nucleotidesequence homology, particularly in the catalytic domains[11]. The CHS1 is relatively larger in size and its expres-sion is the cuticle- and tracheae-specific while CHS2 isreportedly expressed exclusively in the gut [9,16]. In thepresent study we cloned and sequenced the complete

on is asterisked and poly(A) signal sequence is underlined. The putativeer v. 2.0 are shaded gray and the EDR and QRRRW the chitin synthase

Fig. 2 (continued)

Fig. 3. Nucleotide (a) and deduced amino acid (b) sequence alignment ofPxCHS1 alternate exon-A and exon-B. Symbols indicate identicalnucleotides or amino acids (*), highly conserved amino acids (:), conservedamino acids (.). Nucleotide positions used to develop exon-specific PCRprimers are indicated by arrow.


cDNA of a CHS gene of P. xylostella, PxCHS1, by PCRamplification. The nucleotide sequence was 5.5 kb inlength producing a putative protein of 1567 amino acidresidues. The size of the cDNA is very similar to the epi-dermal CHS cDNA from other insect species [7,11,12].The putative protein of PxCHS1 showed the presenceof CHS signature sequences (EDR and QRRRW) in thecatalytic domain [4,7]. With no signal peptide predicted,and 17 transmembrane helices on both sides of the cata-lytic domain, the protein is indicated as non-secretedmembrane-associated protein. We predicted seven trans-membrane regions in the C-terminal domain, which is inagreement with CHS1 studies from other insects [4].The nucleotide and the deduced amino acid sequence ofPxCHS1 cDNA showed a significant homology with


other CHS1 from insects, particularly lepidopterans [7,16]where amino acid residues matched to those of M. sexta

and S. exigua up to 90%. When phylogenetic analysis ofPxCHS1 was performed included with deduced amino

Fig. 4. Deduced amino acid sequence alignment of PxCHS1 with other insect(*), highly conserved amino acids (:), conserved amino acids (.). Amino acidsequences of the putative catalytic domain (Arakane et al., 2004) are shaded g

acid sequences of other CHS1 and CHS2, PxCHS1

sequence was grouped with CHS1 sequences of otherinsects. These results suggest that the gene belongs toCHS1 class.

CHS1 sequences using ClustalW. Symbols indicate identical amino acidssequences to develop degenerate primers are underlined. The amino acidrey.

Fig. 4 (continued)


Investigation from several insects has revealed thatCHS1 contains two alternate exons, thus producing twosplice variants, which is not the case in CHS2 [9,11]. Inthe present study, we found the presence of two alternateexons A and B of 177 bp in PxCHS1. Nucleotide sequenc-ing of the mRNA of PxCHS1 showed that exons A and B

are transcribed in a mutually exclusive manner. The loca-tions of these exons in the cDNA sequence of PxCHS1

matched with those of exon 8A and 8B of other insects[11], and in fact, these exons showed higher homology withthe respective exons than they showed between each other.Studies from M. sexta, T. castaneum, and D. melanogaster

Fig. 5. A dendrogram showing the phylogenetic relationships of aminoacid sequences of the known insect chitin synthase genes. The tree wasobtained by a clustalW alignment and developed by njplot. Insect speciesand the respective accession numbers of the sequences in the tree areprovided in the text.

Fig. 6. RT-PCR expression analysis of PxCHS1 isoforms, PxCHS1A andPxCHS1B, during development using isoform-specific primers. cDNAamplified from the respective samples by primers specific for a constitu-tively expressed gene, b-actin, was used as an internal control. Lane 1,eggs; lane 2, first instar larvae; lane 3, second instar larvae; lane 4, thirdinstar larvae; lane 5, fourth instar larvae; lane 6, pupae; lane 7, adults.

Fig. 7. Real-time qPCR expression analysis of PxCHS1 isoforms,PxCHS1A and PxCHS1B in major body tissues. cDNA synthesized fromeach body tissue, head, body and gut was used in a quantitative PCR andamplified with primers specific for each isoform. The constitutivelyexpressed b-actin gene was used as internal control. The Ct values of theisoforms were normalized for specific tissues by adding or subtracting thedeviation proportion of the Ct values of the b-actin in the samples.


have suggested that only two isoforms of CHS1 gene areexpressed in those insects [11].

We performed expression analysis of the two isoformsof PxCHS1, PxCHS1A and PxCHS1B, in P. xylostelladevelopmental stages using the exon-specific primers inan RT-PCR. Both the isoforms were detected in all the lifestages used in the analysis. The expression levels in fourthinstar were, however, significantly lower and in pupae rel-atively higher. This might suggest that PxCHS1 is notevenly regulated throughout the insect life and changeswith the developmental stage. From a study on D. melano-

gaster, transcripts of DmeCHSA were barely detectablebetween third instar larval stages and the first 2 h afterpupariation [13]. Hogenkamp et al. [9] determined the low-

est expression of MsCHS1 in the fifth instar larvae of M.

sexta at the wandering stage. The fourth instar larvae fromour studies were 24–28 h old and were getting closer towandering stage. A relatively higher expression at pupalstage is in agreement with the results reported for CHS1

expression of M. sexta and S. frugiperda [9,16].Real-time qPCR was performed to measure the mRNA

levels of the two isoforms in three major body tissues, head,body and gut. The outcome indicated uneven levels ofmRNA of the two isoforms in the head and body.Although mRNA levels of PxCHS1A were relativelyhigher in the head (1.5-fold) than the body, the differencewas more pronounced in PxCHS1B (6.3-fold). Significantdifference in expression was also observed betweenPxCHS1A and PxCHS1B in the body. There is evidencefrom M. sexta that expression levels of exon-A and exon-B of MsCHS1 were uneven in epidermis and tracheae withexon-B predominantly expressed in tracheae [9], but theinformation about the specific function of these exons arenot available. Analyzing the functional roles of the differ-ences in expression on chitin development are intriguingfuture investigations. A weaker expression of both iso-forms was also detected in the gut. Expression of CHS1

is known to be cuticle- and tracheae-specific and we haddetected some expression in the gut. Presence of CHS1

mRNA in the gut may be due to the tracheae which wereimpossible to separate from the gut and the attached tissuesdue to smaller size of the larvae.

We treated the third instar larvae with a BPU, chlorflu-azuron, to evaluate the effects on PxCHS1 expression.However, no suppressed or increased mRNA level ofPxCHS1 was observed in the treated larvae. These resultswere in agreement with those of Cohin and Casida [23]and Mayer [19] who had reported that diflubenzuron hadno effects on CHS in T. castaneum and Stomoxys calcitrans,


respectively. Recently, Zhang and Zhu [17] detectedincreased levels of AqCHS1 mRNA in a dipteran, A.

quadrimaculatus after larval treatment with diflubenzuron.At present we have no data to explain the contradictoryconclusions about the BPU effects. More extensive analysisof the tissue-specific expression of PxCHS1 in response toBPUs is necessary to conclude the effects of BPUs on theexpression.

Acknowledgments

This study was supported in part by a Grant-in-Aid forEncouragement of Young Scientists from the Japan Societyfor the Promotion of Science (JSPS) (No. 16780036) andthe Ohara Foundation. M. Ashfaq thanks JSPS for theaward of Postdoctoral Fellowship for Foreign Researchers.

References

[1] A.C. Neville, Biology of the Arthropod Cuticle, Springer-Verlag,New York, 1975.

[2] P. Veronico, L.J. Gray, J.T. Jones, P. Bazzicalupo, S. Arbucci, M.R.Cortese, M. Di Vito, C. De Giorgi, Nematode chitin synthase: genestructure, expression and function in Caenorhabditis elegans and theplant parasitic nematode Meloidogyne artiellia, Mol. Genet. Genom-ics 266 (2001) 28–34.

[3] C. Roncero, The genetic complexity of chitin synthesis in fungi, Curr.Genet. 41 (2002) 367–378.

[4] H. Merzendorfer, Insect chitin synthases: a review, J. Comp. Physiol.B 176 (2006) 1–15.

[5] B. Moussian, H. Schwarz, S. Bartoszewski, C. Nuslein-volhard,Involvement of chitin in exoskeleton morphogenesis in Drosophila

melanogaster, J. Morphol. 264 (2005) 117–130.[6] G.H. Ibrahim, C.T. Smartt, L.M. Kiley, B.M. Christensen, Cloning

and characterization of a chitin synthase cDNA from the mosquitoAedes aegypti, Insect Biochem. Mol. Biol. 30 (2000) 1213–1222.

[7] Y.-C. Zhu, C.A. Specht, N.T. Dittmer, S. Muthukrishnan, M.R.Kanost, K.J. Kramer, Sequence of a cDNA and expression of thegene encoding a putative epidermal chitin synthase of Manduca sexta,Insect Biochem. Mol. Biol. 32 (2002) 1497–1506.

[8] R.L. Tellam, T. Vuocolo, S.E. Johnson, J. Jarmey, R.D. Pearson,Insect chitin synthase cDNA sequence, gene organization andexpression, Eur. J. Biochem. 267 (2000) 6025–6042.

[9] D.G. Hogenkamp, Y. Arakane, L. Zimoch, H. Merzendorfer, K.J.Kramer, R.W. Beeman, M.R. Kanost, C.A. Specht, S. Muthukrish-nan, Chitin synthase genes in Manduca sexta: characterization of agut-specific transcript and differential tissue expression of alternatelyspliced mRNAs during development, Insect Biochem. Mol. Biol. 35(2005) 529–540.

[10] L. Zimoch, H. Merzendorfer, Immunolocalization of chitin synthasein the tobacco hornworm, Cell Tissue Res. 308 (2002) 287–297.

[11] Y. Arakane, D.G. Hogenkamp, Y.C. Zhu, K.J. Kramer, C.A. Specht,R.W. Beeman, M.R. Kanost, S. Muthukrishnan, Characterization oftwo chitin synthase genes of the red flour beetle, Tribolium castaneum,and alternate exon usage in one of the genes during development,Insect Biochem. Mol. Biol. 34 (2004) 291–304.

[12] L. Zimoch, D.G. Hogenkamp, K.J. Kramer, S. Muthukrishnan,H. Merzendorfer, Regulation of chitin synthesis in the larval

midgut of Manduca sexta, Insect Biochem. Mol. Biol. 35 (2005)515–527.

[13] M.E. Gagou, M. Kapsetaki, A. Turberg, D. Kafetzopoulos, Stage-specific expression of the chitin synthase DmeChSA and DmeChSBgenes during the onset of Drosophila metamorphosis, Insect Biochem.Mol. Biol. 32 (2002) 141–146.

[14] M. Ichinomiya, E. Yamada, S. Yamashita, A. Ohta, H. Horiuchi,Class I and class II chitin synthases are involved in septum formationin the filamentous fungus Aspergillus nidulans, Eukaryotic Cell 4(2005) 1125–1136.

[15] Y. Arakane, S. Muthukrishnan, K.J. Kramer, C.A. Specht, Y.Tomoyasu, M.D. Lorenzen, M. Kanost, R.W. Beeman, The Tribo-

lium chitin synthase genes TcCHS1 and TcCHS2 are specialized forsynthesis of epidermal cuticle and midgut peritrophic matrix, InsectMol. Biol. 14 (2005) 453–463.

[16] R. Bolognesi, Y. Arakane, S. Muthukrishnan, K.J. Kramer, W.R.Terra, C. Ferreira, Sequences of cDNAs and expression of genesencoding chitin synthase and chitinase in the midgut of Spodoptera

frugiperda, Insect Biochem. Mol. Biol. 35 (2005) 1249–1259.[17] J. Zhang, K.Y. Zhu, Characterization of a chitin synthase cDNA and

its increased mRNA level associated with decreased chitin synthesis inAnopheles quadrimaculatus exposed to diflubenzuron, Insect Biochem.Mol. Biol. 36 (2006) 712–725.

[18] S.M. Meola, R.T. Mayer, Inhibition of cellular proliferation ofimaginal epidermal cells by diflubenzuron in pupae of the stable fly,Science 207 (1980) 985–987.

[19] R.T. Mayer, A.C. Chen, J.R. DeLoach, Chitin synthesis inhibitinginsect growth regulators do not inhibit chitin synthase, Experientia 37(1981) 337–338.

[20] I. Ishaaya, J.E. Casida, Dietary TH 6040 alters composition andenzyme activity of housefly larval cuticle, Pest. Biochem. Physiol. 4(1974) 484–490.

[21] D.H. Deul, B.J. de Jong, J.A.M. Kortenbach, Inhibition of chitinsynthesis by two 1-(2,6-disubstituted benzoyl)-3-phenylurea insecti-cides. II, Pest. Biochem. Physiol. 8 (1978) 98–105.

[22] W.H. van Eck, Mode of action of two benzoylphenyl ureas asinhibitors of chitin synthesis in insects, Insect Biochem. 9 (1979) 295–300.

[23] E. Cohen, J.E. Casida, Inhibition of Tribolium gut chitin synthase,Pest. Biochem. Physiol. 13 (1980) 129–136.

[24] R.T. Mayer, A.C. Chen, J.R. DeLoach, Characterization of a chitinsynthase from the stable fly, Stomoxys calcitrans (L.), Insect Biochem.10 (1980) 549–556.

[25] S. Sonoda, H. Tsumuki, Studies on glutathione S-transferase geneinvolved in chlorfluazuron resistance of the diamondback moth,Plutella xylostella L. (Lepidoptera: Yponomeutidae), Pest. Biochem.Physiol. 82 (2005) 94–101.

[26] Y. Tsukahara, S. Sonoda, Y. Fujiwara, F. Nakasuji, H. Tsumuki,Molecular analysis of the para-sodium channel gene in the pyre-throid-resistant diamondback moth, Plutella xylostella (Lepisoptera:Yponomeutidae), Appl. Entomol. Zool. 38 (2003) 23–29.

[27] J.D. Thompson, D.G. Higgins, T.J. Gibson, ClustalW: improving thesensitivity of progressive multiple sequence alignment throughsequence weighting, position-specific gap penalties and weight matrixchoice, Nucleic Acids Res. 22 (1994) 4673–4680.

[28] M. Kozak, Structural features in eukaryotic mRNAs that modulatethe initiation of translation, J. Biol. Chem. 266 (1991) 19867–19870.

[29] A. Krogh, B. Larsson, G. von Heijne, E.L.L. Sonnhammer, Predict-ing transmembrane protein topology with a hidden markov model:application to complete genomes, J. Mol. Biol. 305 (2001) 567–580.

[30] J.D. Bendtsen, H. Nielsen, G. Heijne, S. Brunak, Improved predictionof signal peptides: SignalP 3.0, J. Mol. Biol. 340 (2004) 783–795.

Documents

Developmental and tissue-specific expression of CHS1 from Plutella xylostella and its response to chlorfluazuron