# Expression map of a complete set of gustatory receptor genes in chemosensory organs of Bombyx mori

Huizhen Guo, Tingcai Cheng, Zhiwei Chen, Liang Jiang, Youbing Guo, Jianqiu Liu, Shenglong Li, Kiyoko Taniai, Kiyoshi Asaoka, Keiko Kadono-Okuda, et al.

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Huizhen Guo, Tingcai Cheng, Zhiwei Chen, Liang Jiang, Youbing Guo, et al.. Expression map of a complete set of gustatory receptor genes in chemosensory organs of Bombyx mori. Insect Biochemistry and Molecular Biology, 2017, 82, pp.74-82. 10.1016/j.ibmb.2017.02.001. hal-01602615

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Expression map of a complete set of gustatory receptor genes in chemosensory organs of *Bombyx mori*

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CrossMark

Huizhen Guo a, Tingcai Cheng a, Zhiwei Chen a, Liang Jiang a, Youbing Guo a, Jianqiu Liu a, Shenglong Li a, Kiyoko Taniai b, Kiyoshi Asaoka b, Keiko Kadono-Okuda b, Kallare P. Arunkumar c, Jiaqi Wu d, Hirohisa Kishino d, Huijie Zhang e, Rakesh K. Seth f, Karumathil P. Gopinathan g, Nicolas Montagné h, Emmanuelle Jacquin-Joly i, **, Marian R. Goldsmith j, **, Qingyou Xia a, Kazuei Mita a, *

ARTICLE INFO

Article history:

Received 25 August 2016

Received in revised form

30 January 2017

Accepted 2 February 2017

Available online 7 February 2017

Keywords:

Insect-plant interactions

Annotation

Taste

Gustatory receptor

*Bombyx mori*

RNA-seq

A B S T R A C T

Most lepidopteran species are herbivores, and interaction with host plants affects their gene expression and behavior as well as their genome evolution. Gustatory receptors (Grs) are expected to mediate host plant selection, feeding, oviposition and courtship behavior. However, due to their high diversity, sequence divergence and extremely low level of expression it has been difficult to identify precisely a complete set of Grs in Lepidoptera. By manual annotation and BAC sequencing, we improved annotation of 43 gene sequences compared with previously reported Grs in the most studied lepidopteran model, the silkworm, *Bombyx mori*, and identified 7 new tandem copies of *BmGr30* on chromosome 7, bringing the total number of *BmGrs* to 76. Among these, we mapped 68 genes to chromosomes in a newly constructed chromosome distribution map and 8 genes to scaffolds; we also found new evidence for large clusters of *BmGrs*, especially from the bitter receptor family. RNA-seq analysis of diverse *BmGr* expression patterns in chemosensory organs of larvae and adults enabled us to draw a precise organ specific map of *BmGr* expression. Interestingly, most of the clustered genes were expressed in the same tissues and more than half of the genes were expressed in larval maxillae, larval thoracic legs and adult legs. For example, *BmGr63* showed high expression levels in all organs in both larval and adult stages. By contrast, some genes showed expression limited to specific developmental stages or organs and tissues. *BmGr19* was highly expressed in larval chemosensory organs (especially antennae and thoracic legs), the single exon genes *BmGr53* and *BmGr67* were expressed exclusively in larval tissues, the *BmGr27–BmGr31* gene cluster on chr7 displayed a high expression level limited to adult legs and the candidate CO₂ receptor *BmGr2* was highly expressed in adult antennae, where few other Grs were expressed. Transcriptional analysis of the Grs in *B. mori* provides a valuable new reference for finding genes involved in plant-insect interactions in Lepidoptera and establishing correlations between these genes and vital insect behaviors like host plant selection and courtship for mating.

© 2017 The Author(s). Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

<footnote>a State Key Laboratory of Silkworm Genome Biology, Southwest University, Chongqing 400715, PR China</footnote>

<footnote>b National Institute of Agrobiological Sciences, 1-2 Owashi, Tsukuba 305-8634, Ibaraki, Japan</footnote>

<footnote>c Centre for DNA Fingerprinting and Diagnostics, Hyderabad 500001, India</footnote>

<footnote>d Graduate School of Agricultural and Life Sciences, The University of Tokyo, 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-8657, Japan</footnote>

<footnote>e Ministry of Education Key Laboratory of Diagnostic Medicine, College of Laboratory Medicine, Chongqing Medical University, Chongqing 400016, PR China</footnote>

<footnote>f Department of Zoology, University of Delhi, Delhi 110007, India</footnote>

<footnote>g Indian Institute of Science, Bangalore, India</footnote>

<footnote>h Sorbonne Universités, UPMC Univ Paris 06, Institute of Ecology and Environmental Sciences IEES-Paris, 4 Place Jussieu, Paris F-75005, France</footnote>

<footnote>i INRA, Institute of Ecology and Environmental Sciences IEES-Paris, Route de Saint-Cyr, Versailles F-78000, France</footnote>

<footnote>j Department of Biological Sciences, University of Rhode Island, Kingston 02881, RI, USA</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>** Corresponding author.</footnote>

<footnote>*** Corresponding author.</footnote>

<footnote>E-mail addresses: emmanuelle.joly@inra.fr (E. Jacquin-Joly), mki101@uri.edu (M.R. Goldsmith), mitakazuei@gmail.com (K. Mita).</footnote>
# 1. Introduction

Insects, especially phytophagous insects, have formed specific relationships with their host plants for a long period of evolution. The interaction and co-evolution between insects and host plants laid the foundation for insects to survive and expand into genetic races (Ehrlich and Raven, 1964; Jiang et al., 2015), and for processes in which chemical senses play critical roles, such as detection of food, oviposition sites, predators, and mates. Chemical reception is mediated by specialized sensory neurons located in appendages such as antennae, mouthparts, legs, and ovipositors. While olfactory neurons recognize volatile cues, gustatory neurons sense soluble chemicals by contact. Both mechanisms involve expression of chemosensory genes whose genomic organization, expression and evolution participate in shaping the process of insect-plant interactions (Engsontia et al., 2014; Gardiner et al., 2008; McBride et al., 2007; Vieira and Rozas, 2011; Xu et al., 2016).

At present, the signal transduction process of insect olfactory sensory cells is relatively clear (Benton et al., 2006; Sato et al., 2008; Smart et al., 2008; Wicher et al., 2008), while less is known about the gustatory process (Sato et al., 2011; Zhang et al., 2011). Both involve transmembrane chemoreceptors encoded by olfactory receptor genes (Ors), gustatory receptor genes (Grs) and ionotropic receptor genes (Irs) (Cande et al., 2013; Hansson and Stensmyr, 2011).

Comparative studies of chemosensory gene families among lepidopteran species can lead to new insights into understanding the mechanisms of host plant specialization and insect adaptation. Whereas much effort has been made to identify Or genes in Lepidoptera (Montagne et al., 2015), only fragmentary data are available about Grs, limited to species with published genome sequences such as Bombyx mori (Wanner and Robertson, 2008; Sato et al., 2011; Zhang et al., 2011), Danaus plexippus (Zhan et al., 2011), Heliconius melpomene (Briscoe et al., 2013), Plutella xylostella (You et al., 2013), Manduca sexta (Koenig et al., 2015) and Hypantria cunea (Zhang et al., 2016).

Depending on their host plant ranges, phytophagous insects can be categorized as monophagous, oligophagous or polyphagous. The domesticated silkworm, *B. mori*, is a typical oligophagous insect that feeds mainly on leaves of mulberry tree and its close taxonomic relatives. *B. mori* genome sequence data published in 2004 (Mita et al., 2004; Xia et al., 2004) led to the identification of 65 candidate Gr genes (further referred as *BmGrs*) (Wanner and Robertson, 2008). Four additional partial *BmGrs* (Sato et al., 2011; Zhang et al., 2011) were identified in the highly improved *B. mori* genome sequence published in 2008 (International Silkworm Genome, 2008). However, due to the very low expression level and high divergence observed in insect chemoreceptor sequences as well as short contigs in their genome assemblies, the precise composition and structure of the *BmGr* family have not been characterized completely, nor has the nomenclature of the chemosensory genes been unified, notably for the *Grs*. Subsequently, RNA-seq technology became available as a highly sensitive and low cost method to detect weakly expressed genes, alternative splice variants and novel transcripts using ultra-high-throughput sequencing technology (Buermans and den Dunnen, 2014). This situation gave us an opportunity to carry out a more comprehensive analysis of the silkworm *Gr* gene family.

Here we identified and characterized a complete set of *BmGrs* from the improved silkworm genome assembly and sequences of additional bacterial artificial chromosome (BAC) clones encompassing *Gr* genes. Subsequently, RNA-seq analysis provided a precise expression map of *BmGrs* in various chemosensory organs. The results of this study yield a valuable new reference for comparative studies of plant-insect interactions in Lepidoptera.

# 2. Results

## 2.1. Identification and nomenclature of a complete set of BmGr genes

Based on the *BmGr* sequences in the KAIKObase (Shimomura et al., 2009), SilkDB (Duan et al., 2010) and the NCBI reference database, especially the reported BmGr amino acid sequences (Wanner and Robertson, 2008), we performed BLASTp (cutoff e-value: 1e-05) and tBLASTn using the highly improved silkworm genome assembly (International Silkworm Genome, 2008). We identified a total of 76 *BmGrs*, among which 26 gene sequences (*BmGr9*, *11*, *13*, *14*, *15*, *16*, *17*, *18*, *24*, *25*, *26*, *27*, *30*, *33*, *34*, *41*, *50*, *51*, *53*, *56*, *57*, *58*, *60*, *62*, *63* and *64*) were the same as previously reported. For genes initially reported to lack a 5' end (*BmGr1*, 2 and 5), we determined the first exon and completed the full-length sequences. We made minor revisions of the remaining sequences except for *BmGr44* and *BmGr66*, which differed completely from those previously published.

Zhang et al. (2011) reported *BmGr66-69* as new Gr genes which we checked initially with a BLAST search and subsequently reannotated as follows. We chose H. Robertson's *Gr66* (379aa) as *BmGr66* (Wanner and Robertson, 2008) because *BmGr66-Zhang* (341aa) aligned with *BmGr67* (acc. # NM_001246287.1, 351aa) [1–314]. *BmGr67-Zhang*, encoding 275aa, was identical to a portion of *BmGr68* (encoding 418aa), which was derived from a *BmGr66* sequence reported by Sato et al. (2011) (acc. # AB600835.1, 344aa, partial). *BmGr68-Zhang* encoding 177aa was a part of *BmGr44* (306aa) [130–306]. tBLASTn search of *BmGr69-Zhang* (188aa) in KAIKObase showed that it hit sequences between *BmGr17* and *BmGr14* on chr7. A subsequent BLASTp search in NCBI showed homology with *BmGr14* (370aa) with an amino acid identity of 93/202 (46%). Since *BmGr17*, *14*, *15* and *16* formed a gene cluster, we checked whether any new *Grs* were predicted in the domain chr7: 3,547,001–3,552,000 using fgenesh and genescan gene prediction programs and KAIKObase, but could not find any *Gr* genes in this region. Therefore we concluded that *BmGr69-Zhang* was not a genuine *Gr*.

We found one gene, *BmGr31*, located in a gap on chr7: 3,442,653–3,534,450. Since we found several other *Grs* flanking the gap (91 kb), we investigated whether it might harbor other copies of *Grs* in a large gene cluster. Sequencing of BAC clone 092J04 [chr7: 3,376,7764–3,551,821] (http://sgp.dna.affrc.go.jp/KAIKObase/), which covered the gap, revealed 7 additional tandemly aligned copies of *BmGr30*. We also corrected published sequences for *BmGr29* (acc. #: BK006604) and *BmGr30* (acc. #: BK006605), which were located just at the edges of the gap. Comparison of the *BmGr29-BmGr31* sequences showed high sequence identity. Thus, in total we found 10 copies of *BmGr30* in this gap (Fig. 1).

For the nomenclature of these genes, we basically followed the previously published name order (Sato et al., 2011; Wanner and Robertson, 2008; Zhang et al., 2011). For the newly identified genes on chr7, we listed them as *BmGr30-1* to *BmGr30-8* and adjusted the gene numbering based on their chromosomal location. The 154,052 bp sequence of BAC 092J04 is available in DDBJ (http://www.ddbj.nig.ac.jp) under accession number LC056060. We improved 43 genes with suffixes marked "XX" or previously published partial *Gr* gene structures by this work. As noted above, *BmGr66* was based on the *Gr66* sequence kindly provided by H. Robertson, while *BmGr66-AB600835* (Sato et al., 2011) was changed to *BmGr68*, and *BmGr68-NM_001246288* (Sato et al., 2011) was changed to *BmGr69*. This resulted in an increase of the total number of *BmGrs* from 69 to 76 and significant improvement of previous annotations. All gene sequences and detailed gene information for *BmGrs* are shown in Table S1.
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Detailed gene distribution of *BmGrs* in the Chromosome 7 gene cluster. BAC clone 092J04 [chr7:3,376,776 ... 3,551,821] was shown to encompass a gap found in the organization of *BmGr* genes. Sequencing of this BAC revealed 7 additional copies of *BmGr30* tandemly aligned in this gap. The 154,052 bp sequence of BAC092J04 is available in DDBJ under accession number LC056060. Red arrows indicate direction of gene transcription.</figcaption></figure>

## 2.2. *BmGrs* are distributed on 16 chromosomes and most are arranged in clusters

We improved a previously constructed map of *BmGrs* (Engsontia et al., 2014) to localize 68 genes to chromosomes (Fig. 2). The remaining 8 *Grs* were mapped to scaffolds. *BmGr65*, composed of four exons, had been problematic, because the first two exons were on chr12 and the other two exons were on chr23 in the previous annotation. Here we corrected the location of *BmGr65* and determined that it was close to *BmGr64* on chr12 (Document S1). *BmGr41*, *BmGr42* and *BmGr43* were located on unmapped *Bm_scaf444* with a size of 21,432bp. KAIKObase showed a gap of 22 kb between *Bm_scaf1_contig631* and contig630 [chr13: 11,532,372–11,554,690]. Here, we determined that *BmGr41–43* were localized in this gap by PCR using genomic DNA (Document S2). We also confirmed the gene structure of *BmGr46* and the order of the genes in the corresponding cluster which should be *BmGr46–BmGr41–BmGr42–BmGr43* (Document S3).

From the chromosome distribution map, we found that most *BmGrs* were in clusters. Putative CO₂ receptors (*BmGr1*, 2 and 3) (Wanner and Robertson, 2008) were located separately on chr7, chr8 and chr23, whereas putative sugar receptors, *BmGr4*, *BmGr5* and *BmGr6* (Mang et al., 2016; Wanner and Robertson, 2008), formed a cluster on chr15; *BmGr7* and *BmGr8* remained unmapped. Most of the remaining *Grs*, provisionally considered as putative bitter receptors (Wanner and Robertson, 2008), formed clusters, especially on chr7 (16 genes) and chr13 (16 genes). *BmGr14–BmGr17*, located in the 3' region of the chr7 gene cluster, and 12 *Grs*, *BmGr27–BmGr31*, in the 5' portion, presented a high level of sequence identity. *BmGr49–BmGr52* formed a cluster on chr6, expansion of which is proposed to have occurred by recent gene duplication events (Figs. 2 and 3).

A phylogenetic analysis (Fig. 3) was carried out using the updated *BmGr* repertoire together with *Grs* identified in *M. sexta* and *H. melpomene*. As observed previously (Engsontia et al., 2014), candidate CO₂ receptors (*BmGr1*–3) grouped within a single clade and exhibited a high conservation level among the three lepidopteran species. The same applies for candidate sugar receptors (*BmGr4*–8) and for the *Drosophila DmGr43a* orthologs, which include D-fructose receptor *BmGr9* (Sato et al., 2011), and the inositol receptor *BmGr10* (Kikuta et al., 2016). The remaining 66 *BmGrs*, including all the newly identified genes, belong to highly

divergent clades of candidate bitter receptors, with very few one-to-one orthology relationships evidenced among the three species (e.g., *BmGr54* and 63). Genes clustered on the same chromosome also belonged to the same clade in the phylogenetic tree. Most bitter *Grs* shared a similar gene structure composed of 3–5 exons, except for *BmGr53* and *BmGr67* which had a single exon each, whereas CO₂ and sugar receptor genes had more complex gene structures composed of 5–9 exons and 8–12 exons, respectively. The CDS sequence of bitter receptor genes within each gene cluster shared more than 70% identity, whereas intron identity decreased to around 45%. Notably, we observed 95%–99% homology for CDS sequences and 62%–94% for introns among the newly identified copies of the chr7 gene cluster, suggesting they expanded through very recent gene duplication events. Thus, not only the exons but also the introns were congruent with the phylogeny (Fig. 3).

## 2.3. Transcriptional analysis of *BmGrs* in silkworm larval and moth chemosensory organs

Determining whether identified *Gr* genes are indeed expressed and in which tissue they are expressed is essential for understanding their role in insect interactions with the environment. Here we used RNA-seq analysis based on the more precise gene annotation to determine the spatio-temporal expression profiles of *BmGrs* in various chemosensory organs. RNA-seq detected most transcripts in one or more of the following tissues: larval antennae, larval maxillary galea + palps, larval thoracic legs, adult antennae and adult legs, all collected from both males and females (Figs. S1 and S2). Only two genes (*BmGr25* and *61*) had no evidence of expression in any of the tissues investigated. By contrast, *BmGr63* was highly expressed in all chemosensory tissues examined in larvae and adults, and *BmGr18* and 19 were expressed in every tissue except adult antennae. It is notable that 46 *Gr* genes (Fig. S1) were expressed in larval maxillae, 44 *Gr* genes expressed in larval thoracic legs and 52 *Gr* genes (Fig. S2) expressed in adult legs, indicating that these pairs of appendages are important gustatory tissues in *B. mori*. Relatively fewer *Gr* genes were expressed in larval or adult antennae. Among these, *BmGr19* showed an extremely high expression level in larval antennae, whereas the candidate CO₂ receptor *BmGr2* was highly expressed in adult antennae. *BmGr53* and *BmGr67*, which shared the same single exon gene structure,
<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Chromosomal distribution map of *BmGr* genes. The current status of gene cluster distribution is indicated. Newly identified *Gr* genes are marked with dark dots.</figcaption></figure>

were expressed highly in the larval stage, but showed no expression in adult organs.

Considering the putative bitter receptor gene cluster found on chr7, the 12 highly homologous *Grs* constituting the 5' part of the gene cluster (*BmGr27–31*) were expressed exclusively in adult legs. In contrast, *BmGr14–17*, found on the same chromosome, were weakly expressed in larval maxillae, but were not expressed in other tissues. Four genes that formed a cluster on chr6 (*BmGr49–52*) were highly expressed in larval organs, but presented almost no expression in adult organs. The large *Gr* gene cluster on chr13 (Figs. 2 and 4) was split into three parts [*BmGr41/42/43/45/46/48*], [*BmGr32–37/40/47*] and [*BmGr39*]. *Grs* in the first and third parts were highly expressed in larval organs, with almost no expression in adult tissues, whereas the seven *Grs* in the central part were mainly expressed in adult legs (Fig. 4 and Fig. S1). These expression patterns were consistent with the phylogenetic tree (Fig. 3).

Several of these *Grs* showed sex-biased expression. *BmGr10*, 16, 28, 29, 37, 38, 45, 49, 58 and 66 were male-biased, whereas *BmGr6*, 15, 18, 20, 24, 30-1–30-8, 33–36, 40, 55, 62, 67 and 69 were female-biased. Comparison of *Gr* gene expression between larval legs and moth legs called attention to two sets of genes with complementary patterns (Fig. S2). Notably, *BmGr19* (chromosome site unknown) was expressed at an extremely high level in larval thoracic legs, whereas expression of the *BmGr27–31* gene cluster (chr7) was strictly limited to moth legs.

Larval mouthparts contain many sensilla located on maxillary galea, maxillary palps and epipharynx, which are the most important chemosensory organs for larval food selection (Ishikawa and Hirao, 1961). In order to understand which genes in

mouthparts could be responsible for silkworm larval food selection, we dissected out larval maxillary palps, maxillary galea, and epipharynx separately to perform RNA-seq (Fig. S3). More *BmGrs* were expressed in the maxillary palps than maxillary galea or epipharynx. The inositol receptor *BmGr8* (Zhang et al., 2011) was mainly expressed in the maxillary galea, whereas the D-fructose receptor *BmGr9* was expressed highly in maxillary palps. The three candidate CO₂ receptors (*BmGr1*, 2 and 3) were highly expressed in the maxillary palps of both sexes, and sugar receptors were mainly expressed in the maxillary galea and palps; however, bitter receptors could be detected not only in maxilla, but also in epipharynx. Genes highly expressed at the larval stage (*BmGr18* and *BmGr19*) had much higher expression in epipharynx and maxillary palps than maxillary galea in both sexes. The recently expanded *BmGrs* identified on chr7 (*BmGr27* to 31) showed almost no expression in these organs.

### 3. Discussion

Thanks to genome sequencing and comparative genomics, in recent years data have accumulated on the size of *Gr* gene families in different species and on their evolution (Engsontia et al., 2014; Lavagnino et al., 2012; Zhang et al., 2016). Information on the expression patterns of *Grs* in different developmental stages, different sexes and different chemosensory organs is much sparser, and functional data have been mostly limited to *Drosophila Grs* (Isono and Morita, 2010). Such data are needed for understanding how the *Gr* repertoires of various species have evolved for discrimination of suitable host plants. In Lepidoptera, only two large scale *Gr* expression analyses have been reported: one focused
<figure><img src="image_7.png" /><figcaption>**Fig. 3.** Evolutionary relationships of the Grs identified in three lepidopteran insect genomes. Amino acid sequences were automatically aligned by the Mafft program version 7, using the E-INS-i strategy. The evolutionary history was inferred using a maximum likelihood tree with RaxML version 8 using the LG + Gamma + I model. Model selection was conducted by Mega version 6 and LG + Gamma + I mode. Bootstrap support was 1000 replicates. Putative CO₂ and sugar gustatory receptors show conserved relationships among the three Lepidoptera species, while the remaining bitter receptors are more divergent. Bm, *Bombyx mori*; Hm, *Heliconius melpomene*; Ms, *Manduca sexta*.</figcaption></figure>

only on adult tissues (Briscoe et al., 2013) and the other on adult and larval tissues (Xu et al., 2016). At the functional level, only three Grs have been functionally characterized in *B. mori*, all responding to sugars (Mang et al., 2016; Sato et al., 2011; Zhang et al., 2011).

In the present work, we re-annotated the previously reported Grs of *B. mori* with precise gene structures and a unified nomenclature based on the improved silkworm genome assembly. Furthermore, we identified 7 additional copies of *BmGr30* by sequencing. The *BmGrs* annotated here are located on 16 chromosomes, mostly in clusters in the same chromosomal regions. Based on their high level of sequence identity and similar exon structure, the gene clusters of chr7 and chr13 were formed by recent gene duplication and expansions events. The phylogenetic analysis shows that candidate bitter receptor gene duplications generally occurred after *M. sexta*, *H. melpomene* and *B. mori* separated. In other words, bitter receptors may have evolved faster than other chemosensory genes. This strongly suggests that bitter receptors made a significant contribution to adaptation of Lepidoptera to novel ecological niches. Although the total number of Gr genes we identified is in the range of what has been described in most other lepidopteran species, a recent study revealed significant expansion

of the Gr gene family in the armyworm, *H. armigera* (Lepidoptera: Noctuidae), with a description of up to 197 genes (Xu et al., 2016). Our study confirms the hypothesis that Gr expansion may be linked to polyphagy, since a much lower number of Grs could be identified in the oligophagous *B. mori*.

We also performed the first comprehensive and detailed transcriptional analysis of *BmGr* genes by RNA-seq of a wide range of chemosensory organs including larval antennae, epipharynx, maxillary galea, maxillary palps, and thoracic legs, and moth antennae and legs, each separated by sex. Interestingly, this analysis revealed that most genes located in the same cluster (e.g., genes in the different sections of chr7 and chr13, and *BmGr49–52* on chr6 (Fig. 5) shared the same expression pattern, suggesting their expression is controlled by the same upstream regulatory element(s). It will be interesting to clarify if the clustered Grs are expressed in the same GRN (gustatory receptor neurons) or in a subset of GRNs, as it is known that GRNs can express multiple Gr genes (Scott et al., 2001), contrary to olfactory receptor neurons which usually express only one *Or* gene in addition to the co-receptor *Orco*. This also calls attention to the need for understanding how the stage- and tissue-limited expression of gustatory genes is
<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Distribution map of *BmGr* genes in the Chromosome 13 gene cluster.</figcaption></figure>

coordinated with the regulatory neuronal network required for food choice and other sensory perception.

Several genes showed sex-biased, developmental stage or tissue specific expression and thus may be involved in stage or sex specific behaviors, such as food choice (larvae), oviposition site selection (females) or contact behaviors (males and females). These distinctive expression patterns suggest prioritization of future *Gr* functional studies: *BmGr67* was not only female-biased, but also highly expressed in larval antennae, with no expression in adult tissues. Two candidate CO₂ receptors contrasted in relatively specific expression patterns: *BmGr1* in larval maxilla and *BmGr2*, which was highly expressed in moth antennae. *BmGr63*, *18* and *19* exhibited high expression levels in nearly all organs examined in both larval and adult stages, suggesting an important role in gustation.

Specific expression of *Gr(s)* ensures the functional diversity of GRNs that together shape the way the insect senses its external environment. In caterpillars, these *Grs* are expressed in a pair of styloconic sensilla located on the maxillary galea, each housing 4 GRNs, and in basiconic sensilla located at the top of the maxillary palps, as well as in sensilla on the epipharynx (Dethier, 1937; Ishikawa and Hirao, 1961; Schoonhoven, 1969; Shields, 2009). In *B. mori* larvae, one GRN in the lateral styloconic sensillum is sensitive to sucrose, and the other three respond to myo-inositol, glucose, or salts (Ishikawa, 1963, 1966; Ishikawa and Hirao, 1963). One GRN in the medial sensillum is sensitive to bitter compounds

and the other GRNs respond to water, salts, acids, ecdysone, or 20-hydroxyecdysone (Descoins and Marion-Poll, 1999; Tanaka et al., 1994). Our results are consistent with these physiological data since we found evidence for the expression of sugar receptors, especially the inositol sensitive BmGr8 (Zhang et al., 2011), as well as candidate bitter receptors, in the galea. The gustatory sensilla found in the maxillary palps are involved in food detection and selection (Ishikawa et al., 1969) and we found many *Grs* expressed in this tissue.

In this report, we found that not only maxillae but also antennae and legs expressed gustatory receptors in silkworm, as observed in other Lepidoptera (Briscoe et al., 2013; Legeai et al., 2011; Xu et al., 2016; Zhang et al., 2016), which is in agreement with the observation that these organs carry taste sensilla. In particular, *Gr* expression in female moth legs has been proposed to be a determinant of oviposition site choice in *H. melpomene* (Briscoe et al., 2013). In accordance with this hypothesis, we found over-expression of many *Grs* in female moth legs compared to males. In addition, data in Figs. S1 and S2, clearly show that a set of *BmGrs* was over-expressed in adult legs compared to larval legs and another subset was enriched in larval legs compared to adults, suggesting they play different roles in these differing life stages. It has been reported that during mating, a male fruit fly taps the female with its tarsal leg sensilla to make contact with the female abdomen or to detect the cuticular hydrocarbons (Bray and Amrein, 2003; Ling et al., 2014). In several lepidopteran species, it has been
<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Expression map of *BmGr* genes in larval and adult chemosensory organs. Colored dots on larval and adult chemosensory organs correspond to positions of high expression of *BmGr* genes clustered in chr 7 and chr 13. Other genes mentioned in the text are listed near tissues where they are also highly expressed. Bar graphs are based on the FPKM values of the genes which are shown on the Y axis. The X axis stands for tissues as follows: white, LA; black, LM; horizontal stripes, LTL; vertical stripes, MA; and grey, ML. LA, larval antenna; LM, larval maxilla; LTL, larval thoracic legs; MA, moth antenna; ML, moth legs.</figcaption></figure>

reported that scales act as a releaser of the copulation attempt, or play a key role in the recognition of the correct object for copulation (Ono, 1977). Although not yet described in *B. mori*, such a behavior may exist in this species. *Grs* with high expression in silkworm legs, for example, *BmGr27–31*, are good candidates to mediate such interactions, possibly via recognition of cuticular hydrocarbons during courtship behavior. Those expressed in larval legs would be responsible for food recognition, such as *BmGr18*, *BmGr19* *BmGr53* and so on. Thus, the precise *Gr* expression map in chemosensory organs provides clues for identifying candidate *Grs* involved in critical insect behaviors.

# 4. Materials and methods

## 4.1. Bioinformatics, re-annotation, and nomenclature

We referenced sequence data from published papers on the identification of silkworm *BmGrs* (Wanner and Robertson, 2008; Zhang et al., 2011) and combined it with the silkworm genome data (http://sgp.dna.affrc.go.jp/KAIKobase/ (Shimomura et al., 2009) and http://www.silkdb.org/silkdb/ (Duan et al., 2010)) and NCBI reference data (http://www.ncbi.nlm.nih.gov/nucleotide/), followed by checking manually all members of the gene family individually. In addition, H. Robertson kindly provided us with all BmGr amino acid sequences that he identified (Wanner and Robertson, 2008). We performed tBLASTn search (cutoff e-value: 1e-05) using amino acid sequences of reported *Grs* to identify all possible candidate genes. For each identified gene, each exon/intron boundary was checked manually. Each identified gene was also evaluated by BLASTp search in public protein databases, and examined by HMMER3 search (cutoff e-value: 1e-03) using the Pfam database as well as ExPASy Prosite Release 20.120<prosite.expasy.org>. We determined a unified nomenclature by following that of the *Grs* in the papers cited (Sato et al., 2011; Wanner and Robertson, 2008; Zhang et al., 2011).

## 4.2. BAC sequencing

In order to complete the *BmGr* gene cluster on chr7, we sequenced BAC 092J04 by the BAC shotgun method as follows. We constructed two shotgun libraries of 2 kb and 5 kb from which we picked 590 clones randomly for each library, followed by pair-end sequencing with an ABI3730 DNA Analyzer (Applied Biosystems). After vector-trimming and removal of low quality reads (QV < 20), we assembled all pair-end reads with the programs Phrap 1.08081222 (de la Bastide and McCombie, 2007) and Consed 16.0 (Gordon et al., 1998). The sequence of BAC092J04 (154,052 bp) is available in DDBJ under accession number LC056060.

## 4.3. Chromosomal distribution of chemosensory gene families in the silkworm genome

Using the physical map of *B. mori* provided in SilkDB we imported the *BmGr* nucleotide sequence data into the online SilkMap tool (http://www.silkdb.org/silksoft/silkmap.html) to output a gene-distribution map automatically.

## 4.4. Phylogenetic analysis of BmGrs

We conducted phylogenetic analysis with silkworm (Table S1) and other lepidopteran Grs from *H. melpomene* (Briscoe et al., 2013) and *M. sexta* (Koenig et al., 2015). Amino acid sequences were automatically aligned by the Mafft program version 7 (http://mafft.cbrc.jp/alignment/software/algorithms/algorithms.html), using E-INS-i strategy (Katoh and Standley, 2013). As the alignment showed highly conservative and non-conservative regions, only the conservative regions were retained for further analysis, and sequences with lengths of 340 aa were used for tree inference. Model selection was conducted by Mega version 6 (Tamura et al., 2013) and LG + Gamma + I mode (Hasegawa et al., 1985; Le and Gascuel, 2008; Yang, 1994) was found best for our dataset. The maximum
likelihood tree was inferred by RaxML version 8 (Stamatakis, 2014) using the LG + Gamma + I model. To evaluate the confidence of the tree topology, the bootstrap method (Sanderson and Wojciechowski, 2000) was applied with 1000 replications using the rapid bootstrap algorithm (Stamatakis et al., 2008).

### 4.5. Silkworm strains and sample preparation

The silkworm strain, *Dazao*, was maintained in the Silkworm Gene Resource Library, Southwest University, China, by rearing on fresh mulberry leaves in standard conditions of 12 h light and 12 h dark cycle at 25 °C. We collected the maxillae, antennae and thoracic legs from about 500 2 to 3-day-old fifth instar larvae of each sex, and antennae, forelegs, midlegs and hindlegs from moths, also separated by sex. We washed all tissues with Phosphate-buffered saline (PBS) buffer (NaCl 137 mmol/L, KCl 2.7 mmol/L, Na₂HPO₄ 4.3 mmol/L, KH₂PO₄ 1.4 mmol/L, pH 7.4), and then put them directly into Trizol reagent (Invitrogen, USA) to avoid RNA degradation followed by storage at –80 °C until use.

### 4.6. RNA extraction and Illumina sequencing

We extracted total RNA from the tissues prepared above using Trizol reagent according to the manufacturer's instructions (Invitrogen, USA), and digested contaminating genomic DNA with RNase-free DNase I (Takara, China). We suspended purified RNA in 20 mM sodium acetate buffer (pH 5.2) and quantified samples using a Qubit 2.0 Fluorometer (Life Technologies, Grand Island, NY). We assessed the integrity and quality of the mRNA samples using an Agilent Bioanalyzer 2100 (Agilent Technologies, Santa Clara, CA). We used 1 µg total RNA to make cDNA libraries using a TruSeq RNA sample preparation kit (Illumina, San Diego, CA). In total we prepared 24 individual cDNA libraries by ligating sequencing adaptors to cDNA fragments by PCR amplification and synthesized cDNA products using random hexamer primers, yielding an average length of 260bp. We generated raw sequencing data using an Illumina HiSeq2000 system (Illumina, USA).

### 4.7. Statistical analysis

We manually removed the polyA using *fqtrim* (v0.93) (http://ccb.jhu.edu/software/fqtrim/index.shtml), rRNA and tRNA with Bowtie2 (v2.2.3) (Langmead and Salzberg, 2012) and low quality reads (QV < 20) with Trimmomatic (v0.32) (Bolger et al., 2014), then evaluated and calculated the gene expression level using RSEM software (Li and Dewey, 2011) with the fragments per kb per million reads (FPKM) method (Mortazavi et al., 2008). We directly used the gene sequences as an alignment reference. We integrated each sample into an expression matrix and illustrated the data with heatmaps using R (Logiciel) (https://www.r-project.org/).

## Competing financial interests

The authors declare no competing financial interests.

## Author contributions

HG and KM designed research, MRG, HK, KA, KT, KK, KPA, HZ and QX provided suggestions for research. HG and KM performed most of experiments with the assistance of LJ, ZC, JL and SL. HG, KM, TC, YG and JW analyzed data. HG and KM wrote the primary manuscript. MRG, KK, HK, KPG, RKS, KT, KPA, NM, EJJ and KA revised the manuscript.

## Acknowledgements

This work was supported by the Chinese government fund "One thousand plan for the foreign experts recruitment program (No. WQ20125500074)" and the National Basic Research Program of China (No. 2012CB114600).

## Appendix A. Supplementary data

Supplementary data related to this article can be found at http://dx.doi.org/10.1016/j.ibmb.2017.02.001.

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