Genome-wide identification of chitinase and chitin deacetylase gene families in the oriental fruit fly, Bactrocera dorsalis (Hendel)

Accepted Manuscript Genome-wide identification of chitinase and chitin deacetylase gene families in the oriental fruit fly, Bactrocera dorsalis (Hendel)

Shi-Huo Liu, Hong-Fei Li, Yang Yang, Rui-Lin Yang, Wen-Jia Yang, Hong-Bo Jiang, Wei Dou, Guy Smagghe, Jin-Jun Wang PII: DOI: Reference:

S1744-117X(18)30020-0 doi:10.1016/j.cbd.2018.04.005 CBD 499

To appear in: Received date: Revised date: Accepted date:

14 January 2018 10 April 2018 13 April 2018

Please cite this article as: Shi-Huo Liu, Hong-Fei Li, Yang Yang, Rui-Lin Yang, WenJia Yang, Hong-Bo Jiang, Wei Dou, Guy Smagghe, Jin-Jun Wang , Genome-wide identification of chitinase and chitin deacetylase gene families in the oriental fruit fly, Bactrocera dorsalis (Hendel). The address for the corresponding author was captured as affiliation for all authors. Please check if appropriate. Cbd(2018), doi:10.1016/ j.cbd.2018.04.005

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Genome-wide identification of chitinase and chitin deacetylase gene families in the oriental fruit fly, Bactrocera dorsalis (Hendel)

Shi-Huo Liu1,2, Hong-Fei Li1, Yang Yang1, Rui-Lin Yang1,2, Wen-Jia Yang3,

Key Laboratory of Entomology and Pest Control Engineering, College of Plant

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Hong-Bo Jiang1,2, Wei Dou1,2, Guy Smagghe1,2,4, Jin-Jun Wang*1,2

Academy of Agricultural Sciences, Southwest University, Chongqing 400716,

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Protection, Southwest University, Chongqing 400716, China

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Key & Special Laboratory of Guizhou Education Department for Pest Control and

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Resource Utilization, College of Biology and Environmental Engineering, Guiyang

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University, Guiyang, Guizhou 550005, P. R. China Department of Crop Protection, Faculty of Bioscience Engineering, Ghent

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University, Ghent, Belgium

Running title: Identification of chitinase and chitin deacetylase genes in Bactrocera dorsalis

ms. has 46 pages, 9 figures, 1 table, 5 suppl. files

*Correspondence to: Jin-Jun Wang, College of Plant Protection, Southwest University, Chongqing 400716, P. R. China. E-mail: [email protected],

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[email protected]; Tel: +86-23-68250255; Fax: +86-23-68251269

ACCEPTED MANUSCRIPT Abstract Chitinases (Chts) and chitin deacetylases (CDAs) are important enzymes required for chitin metabolism in insects. In this study, 12 Cht-related genes (including seven Cht genes and five imaginal disc growth factor genes) and 6 CDA genes (encoding

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seven proteins) were identified in Bactrocera dorsalis using genome-wide searching

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and transcript profiling. Based on the conserved sequences and phylogenetic

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relationships, 12 Cht-related proteins were clustered into eight groups (group I–V and VII–IX). Further domain architecture analysis showed that all contained at least

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one chitinase catalytic domain, however, only four (BdCht5, BdCht7, BdCht8 and

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BdCht10) possessed chitin-binding domains. The subsequent phylogenetic analysis revealed that seven CDAs were clustered into five groups (group I–V), and all had

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one chitin deacetylase catalytic domain. However, only six exhibited chitin-binding

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domains. Finally, the development- and tissue-specific expression profiling showed that transcript levels of the 12 Cht-related genes and 6 CDA genes varied

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considerably among eggs, larvae, pupae and adults, as well as among different

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tissues of larvae and adults. Our findings illustrate the structural differences and expression patterns of Cht and CDA genes in B. dorsalis, and provide important information for the development of new pest control strategies based on these vital enzymes. Key words: Bactrocera dorsalis; chitinase; chitin deacetylase; genome-wide; expression profiles

ACCEPTED MANUSCRIPT 1. Introduction Chitin is a high molecular weight polymer of N-acetylglucosamine, and is one of the most common amino polysaccharides in nature (Merzendorfer, 2006). It is widely distributed in fungi, mollusks, and arthropods (Rudall and Kenchington, 1973; Cohen, 2010). In insects, chitin is an important component of the epidermis, trachea,

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peritrophic matrix (PM) and muscle (Kramer and Koga, 1986; Merzendorfer and Zimoch, 2003). Chitin represents up to 60% of dry weight in some insect species,

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which illustrates the importance of this component for insect survival, as well as the

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huge demand for precursors of chitin synthesis (Hackman, 1953; Doucet and

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Retnakaran, 2012).

Chitin forms specific chitinous structures with distinct proteins, catecholamines,

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lipids, and metal ions to organize cuticles and PM in insects (Kramer et al., 1988; Vincent and Wegst, 2004). The rigid insect exoskeleton provides physical support and protection from physical and chemical harm, while the stretchable PM protects

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midgut against pathogen invasion and mechanical injuries. However, chitin restricts

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insect growth and development to some extent, therefore both PM and cuticle must be periodically degraded and restructured to allow for molting and metamorphosis (Zhu

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et al., 2008a; Nakabachi et al., 2010). Therefore, chitin metabolism is an indispensable process for growth, development, and other important physiological

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stages in insects. Based on their functions, the proteins involved in catalyzing insect chitin degradation can be classified into two main categories, chitinases (Chts) and N-acetylglucosaminidases participate in hydrolytic processes, while chitin deacetylases (CDAs) are involved in deacetylation. Chts (EC 3.2.1.14) are one of the largest group of hydrolases, which decompose chitin into N-acetylglucosamines. Chts consist of two families, glycoside hydrolase family 18 (GH18) and family 19 (GH19), based on conserved amino acids, protein folding and other conserved motifs (Henrissat, 1999). GH19 Chts are mostly found in

ACCEPTED MANUSCRIPT plants, while GH18 Chts are widely distributed in animals, plants, and microorganisms. GH18 Cht-like superfamily members not only include Chts with catalytic activity but also some Cht-like proteins, such as imaginal disc growth factors (IDGFs), which lack Cht activity (Funkhouser and Aronson, 2007). All insect Chts belong to the GH18 superfamily, and play roles in food digestion, PM degradation,

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and cuticle formation during molting. In recent years, access to insect genome data has enabled several Cht genes to be identified in many insect species. For instance, 22,

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16, and 17 Cht-related genes were identified in Drosophila melanogaster, Tribolium

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castaneum, and Anopheles gambiae, respectively (Zhu et al., 2008a). Based on similarities in amino acids and phylogenetic analysis, insect Chts and Cht-like

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proteins have been classified into eight groups (Arakane and Muthukrishnan, 2010). Functions of Chts differ a lot among groups. For example, RNA interference (RNAi)

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of the characterized Cht genes in T. castaneum showed that enzymes from groups I and II are involved in digesting cuticular chitin, Chts from group III regulate wing

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expansion and abdominal contraction, whereas some group V members are

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indispensable for adult eclosion (Zhu et al., 2008b). CDAs (EC 3.5.1.41) are secreted metalloenzymes that play a catalytic role in

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the N-deacetylation of chitin. CDAs belong to the carbohydrate esterase family 4 (CE4), and have been well studied in pathogenic microorganisms, such as bacteria

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and fungi (Caufrier et al., 2003). Cloning of the first sequence encoding an insect CDA-like protein occurred in the cabbage looper, Trichoplusia ni (Guo et al., 2005). Since then, research into insect CDAs progressed rapidly, with reports of CDA gene characteristics, domain structures, and enzymatic properties in D. melanogaster, An. gambiae and T. castaneum (Dixit et al., 2008; Arakane et al., 2009). However, the number of CDA genes varies among species, for example, there are five CDA genes in An. gambiae, six in D. melanogaster, and nine in T. castaneum and Manduca sexta (Dixit et al., 2008; Tetreau et al., 2015). Many CDA genes are essential for insect development and molting, for instance, knockdown of TcCDA1 and TcCDA2

ACCEPTED MANUSCRIPT affected molting of T. castaneum (Arakane et al., 2009). CDAs may also play roles in trachea development (Luschnig et al., 2006; Wang et al., 2006), and possibly in insect immunity (Zhao et al., 2010). However, little is known about the biochemical and physiological characteristics of insect CDAs, and their underlying molecular mechanisms.

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The oriental fruit fly, Bactrocera dorsalis (Hendel) is an invasive pest, which

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cases huge economic losses in East Asia and the Pacific (Clarke et al., 2005; Wang et al., 2013). Although chemical control is a convenient option, extensive and perennial

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exposure has caused B. dorsalis to develop resistance to a variety of insecticides, including pyrethroids, organophosphates, avermectins and carbamate (Hsu et al.,

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2004; Jin et al., 2011). Because of their crucial role in regulating insect growth and development, chitin degradation enzyme genes could be considered as targets for pest

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control (Kramer and Muthukrishnan, 1997), and environmentally friendly strategies

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could be developed based on these potential targets.

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In this study, we focused on the identification and annotation of Cht and CDA genes in B. dorsalis. We described sequence homology, domain structures, and gene expression patterns of the Chts and CDAs, and propose putative physiological

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functions for these enzymes. Our findings lay the foundation for further study into

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development of novel pest control strategies based on genes of these vital enzymes. 2. Materials and methods 2.1 Insect cultures

The laboratory colony of B. dorsalis was collected from Hainan province, China, in 2008, and maintained in the phytotron at 27.5 ± 0.5 °C, 75 ± 5% relative humidity, and a photoperiod of 14:10 h (L:D). Larvae were fed with a previously described artificial diet mainly consisting of yeast powder, sugar, wheat germ flour, corn flour, while adults were fed with the artificial diet mainly consisting of yeast

ACCEPTED MANUSCRIPT powder, sugar, honey and water (Wang et al., 2013). 2.2 Gene identification To identify genes encoding Cht-related proteins and CDAs in B. dorsalis, sequences of Chts and CDAs from D. melanogaster, An. gambiae, and T. castaneum were downloaded from the databases of FlyBase (http://flybase.org/), VectorBase

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(https://www.vectorbase.org/), and BeetleBase (http://www.beetlebase.org/). The

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conserved domains, including consensus catalytic domains of Chts (GH18, [pfam00704]) and CDAs (CE4-1, [cd10974] and CE4-2, [cd10975]) were used as

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queries to identify homologous genes and related genes in the B. dorsalis genome

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database (i5K, https://i5k.nal.usda.gov/webapp/blast/) and full-length transcriptome data (unpublished data, not released on NCBI) using BLAST search. All related

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sequences of BdChts, BdIDGFs and BdCDAs were validated by reverse transcription PCRs and sequencing. Each 25 μL PCR reaction mixture contained 12.5 μL 2× Taq Master Mix (Novoprotein, Shanghai, China), 1 μL each of forward and reverse

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primers (10 μM), 1 μL template cDNA (about 400 ng/μL) and 9.5 μL ddH2O. All

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primers used for reverse transcription PCR were list in Supplementary Table 1, and PCR parameters were set as described previously (Liu et al., 2017). The purified

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products were sequenced by Invitrogen in Shanghai, China. 2.3 Phylogenetic analysis

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To construct a phylogenetic tree of Chts, 90 proteins containing GH 18 domains ([smart00636]), including 46 from Diptera, 24 from Coleoptera, 4 from Hymenoptera, 6 from Hemiptera and 10 from Lepidoptera were downloaded from the NCBI database (https://www.ncbi.nlm.nih.gov/), FlyBase, BeetleBase, Manduca base

(http://agripestbase.org/manduca/),

(http://silkworm.genomics.org.cn/)

and

(Supplementary

the

Silkworm

Table

2).

To

database perform

phylogenetic analysis of CDAs, 52 proteins containing CE4 domains (CE4-1, [cd10974] and CE4-2, [cd10975]), including 18 from Diptera, 11 from Coleoptera, 4

ACCEPTED MANUSCRIPT from Hymenoptera and 19 from Lepidoptera were obtained from the above-mentioned databases (Supplementary Table 3). ClustalW and MEGA 5.0 software were used for sequence alignment and phylogenetic analysis, respectively (Tamura et al., 2011). Phylogenetic trees were generated using the Maximum Likelihood method with 1,000 bootstrap tests. Chts and CDAs from B. dorsalis were

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named BdCht, BdIDGF, and BdCDA with numbers representing the extent of their homology.

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2.4 Domain Analysis

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Multiple domains including catalytic domain, chitin binding domains, and

CDAs

in

B.

dorsalis

using

the

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transmembrane domains were identified from the protein sequences of all Chts and NCBI

Conserved and

SMART

Search tool

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(https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi)

Domains

(http://smart.embl-heidelberg.de/). The theoretical parameters of the deduced protein and alignments of multiple sequences were performed using online softwares

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ExPASy Proteomics Server (http://cn.expasy.org/tools/pi_tool.html) and MAFFT

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version 7 (http://mafft.cbrc.jp/alignment/server/), respectively.

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2.5 Temporal and developmental expression profiles Samples at different age (including eggs, 1 to 9-day-old larvae, later larvae,

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pre-pupae, early pupae, 1 to 9-day-old pupae and 0 to 10-day-old adults) and various tissues (including midgut, fat body, trachea, integument, Malpighian tubule and central nervous system of third-instar larvae, and midgut, fat body, Malpighian tubule, ovary and testis of 5-day-old adults) were collected as described previously (Liu et al., 2017). Isolation of total RNA and cDNA synthesis for all samples were carried out using TRIzol® reagent (Invitrogen, Carlsbad, CA, US) and the

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PrimeScript 1st Strand cDNA Synthesis Kit (Takara, Dalian, China), respectively, according to the manufacturers’ instructions. Before cDNA synthesis, all RNA samples were treated with Dnase I (Promega, Madison, WI) according to the manual specification. Specific primers for quantitative real-time PCR (qRT-PCR) were

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designed using online software Primer 3 Web (http://primer3.ut.ee/) (Supplementary

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Table 4) and synthesized by Invitrogen. All qRT-PCRs were carried out with 10 μL

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reaction mixture consisting of 5 μL qPCR Master Mix (Promega, Madison, WI), 0.5

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μL cDNA templates (about 400 ng/μL), and 0.5 mM each of forward and reverse primers. The processes were 95 °C for 2 mins, 40 cycles of 95 °C for 15 s and 60 °C

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for 30 s. The melting curve analysis from 65 to 95 °C was conducted to verify a single PCR product. Three to four independent biological replicates and 2 technical

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replicates were performed for each qRT-PCR experiment. The relative transcript levels were calculated according to ΔCT method using α-tublin as reference gene

3 Results

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(Schmittgen and Livak, 2008; Shen et al., 2012).

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3.1 Chts and Cht-like proteins 3.1.1 Identification and characterization A total of 12 Cht-related genes were identified in the genome of B. dorsalis. Gene names, GenBank accession numbers, and characteristic parameters are listed in Table 1. Besides one previously described Cht gene, BdCht2 (GenBank accession no. KF289944), our extensive search identified 11 genes encoding GH18 Chts. The full-length cDNAs of 10 genes (BdCht -1, -5, -7, -8, -11, and BdIDGF -1, -2, -3, -4, -6) were obtained by reverse transcription PCR (RT-PCR) or searching full-length

ACCEPTED MANUSCRIPT transcriptome data, whereas cDNA of BdCht10 was not obtained in this study. Among 12 Cht-related genes, 7 genes encode Chts, of which the protein length varied from 403 of BdCht1 to more than 2500 amino acids of BdCht10. Other 5 genes encode IDGFs, which belong to GH18 Cht-like superfamily but probably lack chitinase activity. The length of 5 IDGFs ranged from 437 of BdIDGF3 to 452 amino

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acids of BdIDGF6 (Table 1).

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3.1.2 Phylogenetic and structural analysis

Phylogenetic analysis based on amino acid sequences showed that insect Chts

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and Cht-like proteins were clustered into 10 distinct groups (I–X) (Fig. 1). The 12

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Cht-related proteins of B. dorsalis were clustered into eight groups, of which seven only contained one BdCht, with BdCht5 in group I, BdCht10 in group II, BdCht7 in

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group III, BdCht8 in group IV, BdCht2 in group VII, BdCht11 in group VIII, and BdCht1 in group IX. Furthermore, group V contained five Cht-like proteins (BdIDGF1-4 and BdIDGF6) of B. dorsalis, whereas no BdChts were clustered into

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group VI and group X (Fig. 1).

The domain architecture of these Cht-related proteins showed that they all contained the GH-18 catalytic domain. BdCht7 and BdCht10 contained two and four

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catalytic domains, respectively, whereas the others only contained one catalytic domain (Fig. 2). Multiple sequence alignment showed that each of these 16 catalytic had

four

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regions

conserved

motifs

with

the

sequences

KxxxxxGGW,

FDGxDLDWEYP, MxYDxxG, and GxxxWxxDxDD, respectively (Fig. 3). The number of conserved amino acids in the 11-residues critical catalytical domain (motif II) ranged from 5 to 11 (Fig. 4). Among these BdChts and BdIDGFs, only four (BdCht5, BdCht7, BdCht8, and BdCht10) contained 1-4 chitin binding domains (Fig. 2). Eleven proteins were predicted to have a signal peptide, whereas BdCht11 was predicted to have transmembrane domains at the N-terminal (Fig. 2). 3.1.3 Spatio-temporal expression patterns

ACCEPTED MANUSCRIPT Developmental stage- and tissue-specific expressions of BdChts and BdIDGFs were determined using qRT-PCR, and the results were displayed as heatmaps (Fig. 5 and 6). All 12 Cht-related genes were expressed from eggs to adults. Among them, BdIDGF4 and BdIDGF6 were expressed at high levels from eggs to 10-day-old adults, whereas BdCht11 was expressed at a relatively low level during the entire

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development of B. dorsalis (Fig. 5 A and B). Relative mRNA expressions of BdIDGF1 and BdIDGF3 in the adult stage were higher than those of eggs and pupae,

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while the relative expression levels of BdIDGF2, BdCht1 and BdCht2 were stable

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from eggs to adults (Fig. 5 A and B). Of note, BdCht5, BdCht8 and BdCht10 were expressed at high levels during larvae-pupae metamorphosis compared with other

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stages (Fig. 5 A).

Among different tissues, five genes (BdCht1, BdIDGF -2, -3, -4, and -6)

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displayed relatively high expression in all six larval tissues investigated, and all of them were most highly expressed in the larval fat body. In contrast, BdCht -5, -7, -8,

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-10, and -11 were expressed at relatively low levels in larval tissues, with the highest

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expressions found in the midgut and trachea (Fig. 6). In the case of adult tissues, five genes (BdCht1, BdIDGF -2, -3, -4, and -6) were expressed at high levels in all

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tissues investigated, whereas BdCht5 was expressed at low levels in these tissues (Fig. 6). And nine genes (BdCht -1, -2, -7, -10, and all BdIDGFs) were most highly

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expressed in the fat body of female adult, whereas the highest expressions of BdCht5 and BdCht11 were found in the Malpighian tubule of female adults. Noteworthy, BdCht8 was only highly expressed in the midgut of both females and males (Fig. 6). 3.2 Chitin deacetylases 3.2.1 Identification and characterization A total of six CDA genes were identified in the genome of B. dorsalis by BLAST search (Table 1). Two alternatively spliced transcripts were found to arise from BdCDA2, namely BdCDA2A and BdCDA2B. The full-length cDNAs of five

ACCEPTED MANUSCRIPT BdCDAs (BdCDA -1, -2, -3, -4, and -9) were obtained, however, full-length BdCDA5 cDNA could not be obtained either from full-length transcript data or by RT-PCR. The protein lengths of the 7 BdCDAs ranged from 374 of BdCDA3 to 3140 amino acids of BdCDA5 (Table 1). 3.2.2 Phylogenetic and structural analysis

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CDAs from insects were clustered into five groups based on amino acid

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sequence homology, and each group contained one or more BdCDAs (Fig. 7). Among them, BdCDA1, BdCDA2A and BdCDA2B were clustered into group I,

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whereas BdCDA3, BdCDA4, BdCDA5, and BdCDA9 were clustered into groups II,

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III, IV and V, respectively (Fig. 7). Each BdCDA was closely related to the corresponding CDA of D. melanogaster with high bootstrap values (Fig. 7).

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The domain organization of BdCDAs showed that all BdCDAs contained the deacetylase catalytic domain (pfam 01522) (Fig. 8). BdCDA-1, -2A, -2B, and -4

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contained the CE-like 1 domain, whereas BdCDA5 and BdCDA9 had the CE-like 2

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domain (Fig. 8). Among these seven BdCDAs, five (BdCDA -1, -2A, -2B, -4 and -5) contained a chitin binding pertrophin-A domain (CBM-14 domain [pfam 01607]), and only BdCDA -1, -2A, and -2B were predicted to possess a low density

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lipoprotein receptor domain (LDLa, [cd 00112]) (Fig. 8). The amino acid alignment of the catalytic domains of CE-4 revealed that each BdCDA contained five signature

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motifs and one or more predicted N-glycosylation sites (Fig. 9). 3.2.3 Spatio-temporal expression patterns Similar to Cht-related genes, all seven BdCDA genes were expressed from eggs to adults (Fig. 5 C). Among them, BdCDA1 and BdCDA2A were expressed at high levels during the entire developmental period, whereas BdCDA3 and BdCDA9 were expressed at low levels from eggs to 10-day-old adults (Fig. 5 C). Relative BdCDA5 mRNA expression was higher from 5-day-old pupae to 4-day-old adults (both males

ACCEPTED MANUSCRIPT and females) than at other stages of development (Fig. 5 C). BdCDA1 and BdCDA2B were expressed at extremely high levels during the early stage of adult (0- to 2-day-old) (Fig. 5 C). BdCDAs were expressed at highly varied levels among larval and adult tissues. BdCDA1 and BdCDA2A were highly expressed in the trachea, integument and

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central nervous system (CNS) of larvae, as well as the fat body of female adults,

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while they were expressed at low levels in the other larval and adult tissues tested (Fig. 6). However, BdCDA2B was only expressed at relatively high levels in the

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integument and CNS of larvae, as well as the fat body in male and female adults (Fig. 6). The highest mRNA levels of BdCDA -3, -4 and -5 appeared in the midgut,

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CNS and integument of larvae, respectively. Notably, BdCDA9 was expressed at extremely low levels in all larval and adult tissues tested, except for the midgut of

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male adults (Fig. 6).

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4. Discussion

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Open access to genome data of different insect species has made it possible to genome-widely identify Cht-related and CDA genes by bioinformatic approaches. Previous studies have identified 16 Cht-related genes and 6 CDA genes in D.

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melanogaster (Zhu et al., 2004; Dixit et al., 2008), 23 Cht-related genes and 9 CDA genes in T. castaneum (Dixit et al., 2008; Zhu et al., 2008b), 11 Cht-related genes

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and 9 CDA genes in M. sexta (Tetreau et al., 2015), 20 Cht-related genes and 5 CDA genes in An. gambiae (Dixit et al., 2008; Zhang et al., 2011a), and 11 Cht-related genes and 4 CDA genes in Nilaparvata lugens (Xi et al., 2014; Xi et al., 2015), respectively. Based on the sequences of conserved domains, the genome-wide search in this study identified 12 Cht-related genes (including 7 BdChts and 5 BdIDGFs) and 6 chitin deacetylase genes (BdCDAs) in B. dorsalis. In comparison with the model insect D. melanogaster, B. dorsalis lacked 5 Cht-related genes (Cht -4, -6, -9, -12, and IDGF5), while possessed an additional gene namely as BdCht1. This

ACCEPTED MANUSCRIPT indicated the loss of genes in evolution or the fact that they were missed during our identification. Although DmCht1 was initially reported as a Cht gene and later identified as part of the catalytic domain of DmCht10 (Zhu et al., 2004; Zhu et al., 2008a), our identification suggested that BdCht1 was an unique Cht gene that shared a low level of identity (34.8%) with the catalytic domains of BdCht10. Multiple

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sequence alignment also found that BdCht1 shared many conserved regions with Cht1s from T. castaneum, M. sexta, and Harpegnathos saltator (Supplementary

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figure 1).

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Because CDA2 and CDA5 undergo alternative splicing, the total number of CDAs in insect is usually larger than the number of coding genes. In this study, we

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identified two isoforms of the mature mRNA of BdCDA2 (named BdCDA2A and BdCDA2B), which encode 534 and 528 amino acid residues, respectively.

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Unfortunately we failed to obtain the cDNA sequence of BdCDA5 either by RT-PCR or from BLAST searches of the full-length transcript database, therefore we could

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not comment about the existence of BdCDA5 isoforms. Phylogenetically, insect Chts are divided into eight groups (group I-VIII), and the family of Chts are currently classified into 11 groups (group I-X, and group h)

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(Tetreau et al., 2015). Such classification might represent an expansion of the members of Chts. In present study, 90 Cht-related proteins were clustered into 10

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groups (group I-X), and each Cht-related protein of B. dorsalis was shown to be closely related to the homologous Cht or IDGF of D. melanogaster. Eight of ten groups contained only one protein (AgCht5- 1-5 were regarded as duplication genes (Zhang et al., 2011b) and not to be considered) in each insect species, whereas groups IV and V had multiple members. However, as the most divergent group, group IV contained only one Cht (BdCht8) in B. dorsalis, although this may reflect the absence of some group IV Chts from our identification. In contrast, CDAs from insects are usually divided into five groups. Group I includes CDA1 and CDA2, and

ACCEPTED MANUSCRIPT three CDAs from B. dorsalis were clustered into this group. Groups II-V had one representative CDA from each insect species (BmCDA9- 1-3 were regarded as duplication genes), although CDA5 contained multiple isoforms. On the other hand, CDAs differ greatly among themselves in architectural structure. For instance, BdCDAs show differences in their amino acid sequence, type of catalytic domain

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(CE4-1 and CE4-2), and presence or absence of CBD and LDLa domains. These differences may reflect distinct biological functions. As previously reported,

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knockdown of CDAs expression in group I (TcCDA1 and TcCDA2) affected

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larval-larval, larval-pupal and pupal-adult molts (Arakane et al., 2009). The high expression of BdCDA1 and BdCDA2 during larval-pupa-adult molts suggested their

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role involvement in metamorphosis of B. dorsalis.

Similarly, Chts considerably with respect to domain structure and sequence

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content. Group I Chts (Cht5s) usually contains an N-terminal signal peptide, a GH-18 catalytic domain, and a C-terminal CBM-14 chitin binding domain (CBD). catalytically

critical

motif

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The

sequence

of

GH18

Chts

consists

of

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FDG(L/F/I)D(L/I)DW(L/E/Q)(F/Y)P (Merzendorfer and Zimoch, 2003). These highly conserved residues, especially the proton donor glutamate (E) (Lu et al.,

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2002), were detected in many Chts in groups I-IV and groups VI-VIII of the current study (Fig. 4). In T. castaneum, TcCHT5 was found to be required for pupal-adult

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molting (Zhu et al., 2008b), however, BdCht5 was highly expressed at larval-pupal and pupal-adult stages, indicating that BdCht5 regulated chitin degradation during larval-pupal and pupal-adult metamorphosis of B. dorsalis. Chts from group II (Cht10s) are long-length Chts that usually contain 4-7 CBDs and 4-5 catalytic domains. The locations and numbers of CBDs and catalytic domains in Cht10s show conserved arrangements depending on the insect species. For example, Lepidopterans such as B. mori (Pan et al., 2012) and M. sexta (Tetreau et al., 2015) have five catalytic domains and seven CBDs with the arrangement

ACCEPTED MANUSCRIPT represented by▲-■-▲-■-■-■-■-■-▲-▲-■-▲, where ▲ represents a catalytic domain and ■ represents a CBD. Coleopterans have five catalytic domains and five CBDs (Royer et al., 2002; Arakane and Muthukrishnan, 2010) with the arrangement ▲-■-▲-■-■-■-▲-▲-■-▲, whereas dipterans only have four catalytic domains and four CBDs (Zhu et al., 2008a; Zhang et al., 2011a) with the arrangement

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▲-■-■-■-▲-▲-■-▲. The domain architecture of BdCht10 was found to be identical

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to that of dipterans.

BdCht7 was classified into group III in the present study, and Chts from this

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group usually contained two catalytic domains and one CBD with the arrangement ▲-▲-■. Many Chts in this group possess transmembrane spans at the N-terminal,

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such as DmCht7, AmCht7, and TcCht7 (Zhu et al., 2008a), suggesting that they act as membrane-anchored enzymes. However, recombinant HlCht7 of Haemophysalis

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longicornis (with a signal peptide at the N-terminal region) could be secreted into the culture medium (You et al., 2003). Moreover, BdCht7 was predicted to possess a

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signal peptide but no transmembrane segment at the N-terminus, suggesting that it

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might also be a secreted enzyme.

Group IV Chts constitute the most divergent and the largest group of Chts.

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Members in this group usually contain a signal peptide, one catalytic domain, and most lack a CBD, whereas BdCht8 possessed a CBD and lied in this group. In the

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current study, BdIDGFs were placed in group V, which mainly consisted of Cht-like proteins sharing high levels of identity with the catalytic domain of Chts. Members in this group share high similarity in domain structure with group IV Chts. Among all five BdIDGFs, the aspartic acid (D) and glutamic acid (E) closest to the tryptophan (W) residue in conserved motif II sequence FDGLDLDWEFP were changed to alanine (A) and glutamine (Q), respectively. The replacement of these two critical residues might result in the total loss of enzymatic activity (Lu et al., 2002). Group VI Chts (Cht6s) are large proteins that are similar to group I Chts with

ACCEPTED MANUSCRIPT respect to their domain structure, however, we failed to identify any BdChts that are homologous with them. Group VII Chts (Cht2s) usually contain a signal peptide and a catalytic domain, but no CBD. They exhibit highly similar domain structures to group IV Chts, and they were placed as an outlier group near group IV Chts in the present study (Fig. 1).

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BdCht11 and BdCht1 were clustered into group VIII and IX, respectively.

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Members of group VIII usually had no signal peptides or CBDs. The predicted transmembrane domain at the N-terminal of BdCht11 indicated that it might be a

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membrane-associated protein. Chts in group IX (Cht1s) usually lack CBDs and contain different residues in the four conserved motifs (Arakane and Muthukrishnan,

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2010) (Supplementary figure 1). Most notably, Cht1s only retain the first four and the sixth amino acids in the sequence FDGLDLDWEFP. The lack of other acidic

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groups and the critical glutamic acid (E) might influence catalytic activity (Lu et al.,

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2002; Zhang et al., 2002).

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Regarding spatio-temporal expression, only a limited number of Chts, IDGFs and CDAs were highly expressed at specific stages or tissues. BdIDGFs might be the exception to this, because they were expressed at relatively high levels at all

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developmental stages and in all tissues tested, which is similar to BmIDGFs in B. mori (Pan et al., 2010). This suggests that IDGFs are essential enzymes required for

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insect growth and development. Previous studies reported that group I Chts (Cht5s) had multiple functions involvement in metamorphosis of insects, knockdown of TcCht5 in T. castaneum influenced pupal-adult molting (Zhu et al., 2008b), while RNAi of group I Cht gene in Spodoptera exigua caused abnormal and lethal effects during larval-pupal and pupal-adult molting (Zhang et al., 2012). However, the current results suggested that BdCht5 might play important roles in larval-pupal and pupal-adult molting because the transcripts were highly detected during these developmental stages. In the larval stage, BdCht1, BdCht2, all BdIDGFs, BdCDA1,

ACCEPTED MANUSCRIPT and BdCDA2A demonstrated high expressions, suggesting their functional involvement in larval molting. BdCDAs in group I were highly expressed in the integument and tracheal tube of larvae, which is consistent with previous findings in D. melanogaster (Luschnig et al., 2006; Wang et al., 2006), T. castaneum (Arakane et al., 2009) and N. lugens

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(Xi et al., 2014). These findings suggest that group I CDAs function in chitin

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degradation in the integument and trachea of insects. Unexpectedly, BdCht1 was found to be highly expressed in the ovary and testis. However, MsCht1 was

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expressed exclusively in these tissues (Tetreau et al., 2015), and a chitin-like component was detected in the ovaries of Aedes aegypti (Moreira et al., 2007). The

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exact function of these is still unknown. Further work should be carried out to determine if Cht1s are active enzymes that participate in chitin degradation in

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reproductive organs. Additionally, BdCht8 was uniquely expressed in the midgut of adult. Similar results were also reported in previous studies. For instance, MsCht8

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was highly expressed in midgut of adult M. sexta (Tetreau et al., 2015), both of

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PxCht8-1 and PxCht8-2 were expressed exclusively in gut tissue of Plutella xylostella (Liao et al., 2016), and TcCht8 was expressed predominantly in the

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posterior midgut of T. castaneum (Arakane and Muthukrishnan, 2010). All these findings suggested that Cht8 may modulate the effect of PM-associated chitin in the

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midgut.

In summary, we identified multiple genes encoding Cht-related proteins and CDAs in B. dorsalis, which differ a lot in their domain structures and expression patterns. They were clustered into eight of the 10 groups of the GH18 Cht family and into five groups of the CDA family, respectively. Results from spatiotemporal expression suggested that they exhibited distinct biological functions at specific developmental stages and tissues. These findings lay the foundation for further investigation of the properties and physiological functions of individual enzymes.

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ACCEPTED MANUSCRIPT Acknowledgements This work was financially supported by the National Key R & D Program of China (2017YFD0202002), Chongqing Research Program of Basic Research and Frontier Technology (CSTC, 2015jcyjBX0061), the Fundamental Research Funds for the

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Central Universities (XDJK2017A014) of China and the earmarked fund for the

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Modern Agro-industry (Citrus) Technology Research System of China (CARS-26).

ACCEPTED MANUSCRIPT References

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Arakane, Y., Dixit, R., Begum, K., Park, Y., Specht, C.A., Merzendorfer, H., Kramer, K.J., Muthukrishnan, S., Beeman, R.W., 2009. Analysis of functions of the chitin deacetylase gene family in Tribolium castaneum. Insect Biochem. Mol. Biol. 39, 355-365. Arakane, Y., Muthukrishnan, S., 2010. Insect chitinase and chitinase-like proteins. Cell. Mol. Life Sci. 67, 201-216. Caufrier, F., Martinou, A., Dupont, C., Bouriotis, V., 2003. Carbohydrate esterase family 4 enzymes: substrate specificity. Carbohyd. Res. 338, 687-692. Clarke, A.R., Armstrong, K.F., Carmichael, A.E., Milne, J.R., Raghu, S., Roderick, G.K., Yeates, D.K., 2005. Invasive phytophagous pests arising through a recent tropical evolutionary radiation: the Bactrocera dorsalis complex of fruit flies. Annu. Rev. Entomol. 50, 293-319. Cohen, E., 2010. Chitin biochemistry: synthesis, hydrolysis and inhibition, in: J. Casas, S.J. Simpson (Eds.), Adv. Insect Physiol., 5-74. Dixit, R., Arakane, Y., Specht, C.A., Richard, C., Kramer, K.J., Beeman, R.W., Muthukrishnan, S., 2008. Domain organization and phylogenetic analysis of proteins from the chitin deacetylase gene family of Tribolium castaneum and three other species of insects. Insect Biochem. Mol. Biol. 38, 440-451. Doucet, D., Retnakaran, A., 2012. Insect chitin: metabolism, genomics and pest management, in: T.S. Dhadialla (Ed.), Adv. Insect Physiol., Academic Press, Burlington, 437-511. Funkhouser, J.D., Aronson, N.N., Jr., 2007. Chitinase family GH18: evolutionary insights from the genomic history of a diverse protein family. BMC Evol. Biol. 7, 96. Guo, W., Li, G.X., Pang, Y., Wang, P., 2005. A novel chitin-binding protein identified from the peritrophic membrane of the cabbage looper, Trichoplusia ni. Insect Biochem. Mol. Biol. 35, 1224-1234. Hackman, R.H., 1953. Chemistry of Insect Cuticle. Biochem. J. 54, 371-377. Henrissat, B., 1999. Classification of chitinases modules, in: P. Jolles, R.A.A. Muzzarelli (Eds.), EXS, Birkhauser Verlag, Basel/Switzerland, 137-156. Hsu, J.C., Feng, H.T., Wu, W.J., 2004. Resistance and synergistic effects of insecticides in Bactrocera dorsalis (Diptera: Tephritidae) in Taiwan. J. Econ. Entomol. 97, 1682-1688. Jin, T., Zeng, L., Lin, Y., Lu, Y., Liang, G., 2011. Insecticide resistance of the oriental fruit fly, Bactrocera dorsalis (Hendel) (Diptera: Tephritidae), in mainland China. Pest Manage. Sci. 67, 370-376. Kramer, K.J., Hopkins, T.L., Schaefer, J., 1988. Insect cuticle structure and metabolism. ACS Sym. Ser. 379, 160-185. Kramer, K.J., Koga, D., 1986. Insect chitin: physical state, synthesis, degradation and metabolic regulation. Insect Biochem. 16, 851-877. Kramer, K.J., Muthukrishnan, S., 1997. Insect chitinases: molecular biology and

ACCEPTED MANUSCRIPT

AC

CE

PT E

D

MA

NU

SC

RI

PT

potential use as biopesticides. Insect Biochem. Mol. Biol. 27, 887-900. Liao, Z.H., Kuo, T.C., Kao, C.H., Chou, T.M., Kao, Y.H., Huang, R.N., 2016. Identification of the chitinase genes from the diamondback moth, Plutella xylostella. Bull. Entomol. Res. 106, 769-780. Liu, S.H., Wei, D., Yuan, G.R., Jiang, H.B., Dou, W., Wang, J.J., 2017. Antimicrobial peptide gene cecropin-2 and defensin respond to peptidoglycan infection in the female adult of oriental fruit fly, Bactrocera dorsalis (Hendel). Comp. Biochem. Physiol. B 206, 1-7. Lu, Y.M., Zen, K.C., Muthukrishnan, S., Kramer, K.J., 2002. Site-directed mutagenesis and functional analysis of active site acidic amino acid residues D142, D144 and E146 in Manduca sexta (tobacco hornworm) chitinase. Insect Biochem. Mol. Biol. 32, 1369-1382. Luschnig, S., Batz, T., Armbruster, K., Krasnow, M.A., 2006. Serpentine and vermiform encode matrix proteins with chitin binding and deacetylation domains that limit tracheal tube length in Drosophila. Curr. Biol. 16, 186-194. Merzendorfer, H., 2006. Insect chitin synthases: a review. J. Comp. Physiol. B 176, 1-15. Merzendorfer, H., Zimoch, L., 2003. Chitin metabolism in insects: structure, function and regulation of chitin synthases and chitinases. J. Exp. Biol. 206, 4393-4412. Moreira, M.F., dos Santos, A.S., Marotta, H.R., Mansur, J.F., Ramos, I.B., Machado, E.A., Souza, G.H.M.F., Eberlin, M.N., Kaiser, C.R., Kramer, K.J., Muthukrishnan, S., Vasconcellos, A.M.H., 2007. A chitin-like component in Aedes aegypti eggshells, eggs and ovaries. Insect Biochem. Mol. Biol. 37, 1249-1261. Nakabachi, A., Shigenobu, S., Miyagishima, S., 2010. Chitinase-like proteins encoded in the genome of the pea aphid, Acyrthosiphon pisum. Insect Mol. Biol. 19, 175-185. Pan, Y., Chen, K., Xia, H., Yao, Q., Gao, L., Lue, P., Huojuan, He, Y., Wang, L., 2010. Molecular cloning, expression and characterization of BmIDGF gene from Bombyx mori. Z. Naturforsch. C 65, 277-283. Pan, Y., Lu, P., Wang, Y., Yin, L., Ma, H., Ma, G., Chen, K., He, Y., 2012. In silico identification of novel chitinase-like proteins in the silkworm, Bombyx mori, genome. J. Insect Sci. 12, 150. Royer, V., Fraichard, S., Bouhin, H., 2002. A novel putative insect chitinase with multiple catalytic domains: hormonal regulation during metamorphosis. Biochem. J. 366, 921-928. Rudall, K.M., Kenchington, W., 1973. The chitin system. Biol. Rev. Camb. Philos. Soc. 48, 597-636. Schmittgen, T.D., Livak, K.J., 2008. Analyzing real-time PCR data by the comparative CT method. Nat. Protoc. 3, 1101-1108. Shen, G.-M., Wang, X.-N., Dou, W., Wang, J.-J., 2012. Biochemical and molecular

ACCEPTED MANUSCRIPT

AC

CE

PT E

D

MA

NU

SC

RI

PT

characterisation of acetylcholinesterase in four field populations of Bactrocera dorsalis (Hendel) (Diptera: Tephritidae). Pest Manage. Sci. 68, 1553-1563. Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S., 2011. MEGA5: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol. 28, 2731-2739. Tetreau, G., Cao, X., Chen, Y.-R., Muthukrishnan, S., Jiang, H., Blissard, G.W., Kanost, M.R., Wang, P., 2015. Overview of chitin metabolism enzymes in Manduca sexta: Identification, domain organization, phylogenetic analysis and gene expression. Insect Biochemistry and Molecular Biology 62, 114-126. Vincent, J.F.V., Wegst, U.G.K., 2004. Design and mechanical properties of insect cuticle. Arthropod Struct. Dev. 33, 187-199. Wang, J.J., Wei, D., Dou, W., Hu, F., Liu, W.F., Wang, J.J., 2013. Toxicities and synergistic effects of several insecticides against the oriental fruit fly (Diptera: Tephritidae). J. Econ. Entomol. 106, 970-978. Wang, S.Q., Jayaram, S.A., Hemphala, J., Senti, K.A., Tsarouhas, V., Jin, H.N., Samakovlis, C., 2006. Septate-junction-dependent luminal deposition of chitin deacetylases restricts tube elongation in the Drosophila trachea. Curr. Biol. 16, 180-185. Xi, Y., Pan, P.L., Ye, Y.X., Yu, B., Xu, H.J., Zhang, C.X., 2015. Chitinase-like gene family in the brown planthopper, Nilaparvata lugens. Insect Mol. Biol. 24, 29-40. Xi, Y., Pan, P.L., Ye, Y.X., Yu, B., Zhang, C.X., 2014. Chitin deacetylase family genes in the brown planthopper, Nilaparvata lugens (Hemiptera: Delphacidae). Insect Mol. Biol. 23, 695-705. You, M., Xuan, X., Tsuji, N., Kamio, T., Taylor, D., Suzuki, N., Fujisaki, K., 2003. Identification and molecular characterization of a chitinase from the hard tick Haemaphysalis longicornis. J. Biol. Chem. 278, 8556-8563. Zhang, D., Chen, J., Yao, Q., Pan, Z., Chen, J., Zhang, W., 2012. Functional analysis of two chitinase genes during the pupation and eclosion stages of the beet armyworm Spodoptera exigua by RNA interference. Arch. Insect Biochem. Physiol. 79, 220-234. Zhang, H., Huang, X., Fukamizo, T., Muthukrishnan, S., Kramer, K.J., 2002. Site-directed mutagenesis and functional analysis of an active site tryptophan of insect chitinase. Insect Biochem. Mol. Biol. 32, 1477-1488. Zhang, J., Zhang, X., Arakane, Y., Muthukrishnan, S., Kramer, K.J., Ma, E., Zhu, K.Y., 2011a. Comparative genomic analysis of chitinase and chitinase-like genes in the African malaria mosquito (Anopheles gambiae). Plos One 6, e19899. Zhang, J., Zhang, X., Arakane, Y., Muthukrishnan, S., Kramer, K.J., Ma, E., Zhu, K.Y., 2011b. Identification and characterization of a novel chitinase-like gene cluster (AgCht5) possibly derived from tandem duplications in the African

ACCEPTED MANUSCRIPT

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PT E

D

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malaria mosquito, Anopheles gambiae. Insect Biochem. Mol. Biol. 41, 521-528. Zhao, Y., Park, R.D., Muzzarelli, R.A.A., 2010. Chitin deacetylases: properties and applications. Mar. Drugs 8, 24-46. Zhu, Q.S., Arakane, Y., Banerjee, D., Beeman, R.W., Kramer, K.J., Muthukrishnan, S., 2008a. Domain organization and phylogenetic analysis of the chitinase-like family of proteins in three species of insects. Insect Biochem. Mol. Biol. 38, 452-466. Zhu, Q.S., Arakane, Y., Beeman, R.W., Kramer, K.J., Muthukrishnan, S., 2008b. Functional specialization among insect chitinase family genes revealed by RNA interference. P. Natl. Acad. Sci. USA 105, 6650-6655. Zhu, Q.S., Deng, Y.P., Vanka, P., Brown, S.J., Muthukrishnan, S., Kramer, K.J., 2004. Computational identification of novel chitinase-like proteins in the Drosophila melanogaster genome. Bioinformatics 20, 161-169.

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Figure legends:

Figure 1. Phylogenetic analysis of 90 insect chitinases (Chts) and chitinase-like proteins (IDGFs) from 10 different species. Acromyrmex echinatior (Ae),

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Anopheles gambiae (Ag), Bactrocera dorsalis (Bd), Camponotus floridanus (Cf),

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Danaus plexippus (Dpl), Drosophila melanogaster (Dm), Harpegnathos saltator

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(Hs), Manduca sexta (Ms), Nilaparvata lugens (Nl) and Tribolium castaneum (Tc). The accession numbers of all the proteins used are listed in Supplementary Table 2.

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A bootstrap analysis of 1000 replications was carried out on the trees inferred from

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the Maximum Likelihood method and bootstrap values are shown at each branch of

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the tree. Chts and IDGFs from B. dorsalis are indicated in bold.

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Figure 2. Domain architectures of 12 chitinase-related proteins in B. dorsalis. The deduced amino acid sequences were used to predict the domain architectures by

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SMART software (http://smart.embl-heidelberg.de/) and NCBI Conserved Domains

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Search (https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi).

Figure 3. Conserved regions in the glycoside hydrolase family 18 (GH18) domain of 12 chitinase-related proteins from B. dorsalis. Amino acid sequences of the catalytic domains of GH18 family enzymes were aligned using MAFFT software (https://mafft.cbrc.jp/alignment/server/). Two and four catalytic domains of BdCht7 and BdCht10 were named as BdCht7-1/2 and BdCht10-1/2/3/4, respectively. Identical and highly conserved amino acids are indicated by (*) and (:), respectively.

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Boxed regions are the four conserved motifs represented by the sequences KxxxxxGGW, FDGxDLDWEYP, MxYDxxG and GxxxWxxDxDD.

Figure 4. Alignment of the 11 amino acids long motif present in GH18 catalytic

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domains of the 12 chitinase-related proteins from B. dorsalis. Two and four motifs of BdCht7 and BdCht10 were named as BdCht7-1/2 and BdCht10-1/2/3/4,

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respectively. For each sequence, the conserved amino acids are indicated in bold and

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the number of conserved amino acids regarding to the reference conserved motif is

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indicated in parentheses. The tree on the left was generated in MEGA5.0 using

was performed 1000 replications.

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Maximum Likelihood method based on 11 amino acids, and bootstrapping analysis

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Figure 5. Developmental mRNA expression profiles of 7 chitinase genes (A), 5

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chitinase-like genes (B), and 7 chitin deacetylase genes (C) of B. dorsalis. The

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ΔCT (ΔCT = CT gene of interest  CT α-tubulin) was used to generate the heatmap. E, eggs; 1-9 d-L, 1 to 9-day-old larvae; later-L, wandering stage larvae; pre-P,

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pre-pupae; early-P, early pupae; 1-9 d-P, 1 to 9-day-old pupae; 0-10 d-FA, 0 to 10-day-old female adults; 0-10 d-MA, 0 to 10-day-old male adults. Warm colors (i.e., red) mean high expression levels and cold colors (i.e., blue) mean low expression levels.

Figure 6. Tissue-specific expression of the chitinase-related genes and chitin deacetylase genes from B. dorsalis at third instar larva and adult. The ΔCT

ACCEPTED MANUSCRIPT (ΔCT = CT gene of interest  CT α-tubulin) was used to generate the heatmap. L-MG, midgut of larvae; L-FB, fat body of larvae; L-Tr, trachea of larvae; L-In, integument of larvae; L-MT, Malpighian tubule of larvae; L-CNS, central nervous system of larvae; F/M-MG, midgut of female/male adults; F/M-FB, fat body of female/male

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adults; F/M-MT, Malpighian tubule of female/male adults; F-Ov, ovary of female

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adults; M-Te, testis of male adults. Warm colors (i.e., red) mean high expression

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levels and cold colors (i.e., blue) mean low expression levels.

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Figure 7. Phylogenetic analysis of 52 insect chitin deacetylases (CDAs) from 7

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different species. Anopheles gambiae (Ag), Apis mellifera (Am), B. dorsalis (Bd), Bombyx mori (Bm), Drosophila melanogaster (Dm), Manduca sexta (Ms) and

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Tribolium castaneum (Tc). The accession numbers of all the proteins used are listed

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in Supplementary Table 3. A bootstrap analysis of 1000 replications was carried out on the trees inferred from the Maximum Likelihood method and bootstrap values are

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shown at each branch of the tree. CDAs from B. dorsalis are indicated in bold.

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Figure 8. Domain architectures of 7 chitin deacetylases in B. dorsalis. The deduced amino acid sequences were used to predict the domain architectures by SMART software (http://smart.embl-heidelberg.de/) and NCBI Conserved Domains Search (https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi).

Figure 9. Alignment of the catalytic domains of the chitin deacetylases containing CE4-like1 (BdCDA 1 to 4) and CE4-like2 domains (BdCDA5 and

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BdCDA9).

Alignment

was

performed

using

MAFFT

software

(https://mafft.cbrc.jp/alignment/server/). Identical and highly conserved amino acids are indicated by (*) and (:), respectively. Boxed regions are the four conserved

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motifs responsible for the catalytic activity.

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Table 1. List of the chitinase-related proteins and chitin deacetylases identified in the B. dorsalis genome and full-length transcriptome.

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Chitin deacetylases

M.W. a /kDa 46.8 54.3 67.5 114.6 56.2 279.2 55.7 49.3 49.0 48.7 48.4 50.3 61.0 60.6 60.0 44.0 56.0 348.3 45.0

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Protein length/aa 403 483 595 1022 505 2504 493 439 438 437 439 452 536 534 528 374 495 3140 394

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Chitinase-like proteins

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Chitinases

Genbank accession no. MF926351 KF289944 KY681041 KY681042 KY426795 XM_019992490 KY426794 KY681043 KY681044 KY681045 KY681046 KY426796 KY681047 KY681049 KY681048 KY681050 KY681051 XM_011211717 KY681052

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Gene name BdCht1 BdCht2 BdCht5 BdCht7 BdCht8 BdCht10 b BdCht11 BdIDGF1 BdIDGF2 BdIDGF3 BdIDGF4 BdIDGF6 BdCDA1 BdCDA2A BdCDA2B BdCDA3 BdCDA4 BdCDA5 b BdCDA9

pI a 9.1 6.0 5.6 5.9 5.3 6.1 6.3 6.3 6.2 6.1 7.2 7.0 5.0 5.0 5.0 5.9 4.9 8.61 6.5

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a, molecular weight (M.W.) and isoelectric point (pI) were predicted using the ExPASy ProtParam tool available at http://web.expasy.org/protparam/; b, full-length cDNA sequences were not obtained in this study, characteristic parameters were predicted based on the NCBI Reference Sequences.

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