Introduction

Chemosensory receptors represent an interface between an insect and its chemical environment, mediating pivotal biological processes such as host finding, mate selection and choice of oviposition sites1. One gene family, the gustatory receptors (GRs), plays a central role in co-ordinating insect feeding behaviours1. Taste stimuli from the environment are recognized by GRs located on the dendrites of taste sensilla, which are distributed throughout the insect body1. Despite a growing body of knowledge about the insect taste system, little is known about the molecular and cellular mechanisms that underlie taste signal recognition or how these signals affect feeding behaviours.

Insect Gr genes were first identified from the Drosophila melanogaster genome2. They have been classified into four clades: CO23, GR43a-like4, sugar5,6,7,8,9,10 and bitter11,12. To date, published research on GRs has been focused on Drosophila5,6,8,9,10,13,14,15,16,17, but with the increasing availability of genomic information from other insect species, such as Anopheles gambiae18, Tribolium castaneum19, Apis mellifera20, and Acyrthosiphon pisum21, GR research is being extended to a diverse range of species. The recent genome projects of four lepidopteran species, silkworm (Bombyx mori)11, Monarch butterfly (Danaus plexippus)22,23, Postman butterfly (Heliconius melpomene23 and Diamondback moth (Plutella xylostella)24 provide invaluable resources for looking deeply into the lepidopteran gustatory system. 69 GRs were identified from B. mori11, 58 GRs were identified from D. plexippus22,23, 73 GRs were identified from H. melpomene23 and 69 GRs were identified from P. xylostella24. These four species are all specialist feeders: B. mori is a mulberry leave specialist11; D. plexippus consumes only plants in the milkweed family (Asclepiadacea)22; H. melpomene feeds on either Passiflora oerstedii or Passiflora menispermifolia23; while P. xylostella feeds exclusively on crucifers24. Therefore, the study on gustatory systems of these four species can shed light on the interactions between host plants and specialist herbivores but may be less relevant to generalist species like Helicoverpa armigera, which include significant pests of human agriculture. H. armigera itself is one of the most polyphagous lepidopteran pest species, with larvae that feed on numerous cultivated crops such as cotton, peanuts, soybeans and maize. H. armigera would be an ideal species for studying the gustatory receptor repertoire of a polyphagous lepidopteran species. Our hypothesis is that the gustatory system of H. armigera may have species-specific features that contribute to its robust polyphagy.

In this study, we utilized genome and transcriptome data from H. armigera and manually identified this species repertoire of Gr genes. We applied phylogenetic analysis, comparative gene expression, and topological and calcium imaging analyses to structurally and functionally characterize the GR repertoire, in the process uncovering a markedly expanded family of lepidopteran GRs.

Results

GR Gene Annotation and Identification

A bioinformatics screen of the H. armigera genome and transcriptome data revealed a total of 197 GR and 64 odorant receptor (OR) genes (Fig. 1A). All identified H. armigera GR nucleotide and amino acid sequences, distributions on scaffolds and other information are available online (Table S1). In comparison with all insect species with currently available genomes, H. armigera showed the second highest number of GRs, 197. This is the highest number of GRs identified in a lepidopteran genome with only the red flour beetle T. castaneum having more, 220 (Fig. 1A). Most insect species studied possess comparable total numbers of ORs and GRs (Fig. 1B). H. armigera is unique among insects in having an increase in total GR numbers relative to OR numbers (Fig. 1B). Of the exceptions, the honeybee, Apis mellifera has an expansion of ORs (163) but possess only 10 GRs20 (Fig. 1A) and wasps and ants also have many more ORs than GRs in their genomes (Fig. 1A).

Figure 1: Relationship between numbers of GRs and ORs in different insect species.
figure 1

(A) Phylogeny of various insect species and the number of ORs and GRs identified from each species’ genome: 1 H. armigera, 2 P. xylostella24,28, 3 B. mori4,11, 4 H. melpomene23 and 5 D. plexippus22,23 (green); 6 A. aegypti48, 7 C. quinquefasciatus49, 8 A. gambiae18 and 9 D. melanogaster2, (black); 10 T. castaneum (blue)19; 11 P. barbatus50, 12 L. humile51, 13 A. mellifera20, and 14 N. vitripennis52, (purple); 15 A. pisum (orange)21; 16 P. humanus53 (brown), 17 L. migratoria54 (light green) and 18 Z. nevadensis55 (grey). (B) A map of numbers of ORs (Y-axis) and GRs (X-axis) of insect species mentioned above. Lepidoptra (), Diptera (●), Hymenoptera (), Coleoptera (), Hemiptera (), Phthiraptera (), Orthoptera () and Blattodea ().

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Phylogenetic analysis

To investigate the types of GRs that were expanded, we selected thirteen GRs from B. mori as representative of the four insect GR subfamilies and compared them in a phylogenetic analysis to all 197 GRs from H. armigera (Fig. 2A). A phylogeny tree with all B. mori GRs and H. armigera GRs was provided (S7 Figure). The analysis reveals HarmGR1–3 are members of the CO2 receptor subfamily25 (Fig. 2A). HarmGR4–8 and 10–12 are members of the insect sugar receptor subfamily and share 46–99% identity with other known moth sugar GRs (Table S1). HarmGR9 and 13 are members of the GR43a-like receptor subfamily26. The remaining 184 HarmGRs form a large “bitter” receptor subfamily with the representative B. mori bitter GRs (BmorGR53, 58 and 66) (Fig. 2A). Four of these GRs, (HarmGR57, 158, 179 and 193) contain a stop codon in the ORF and are likely to be pseudogenes. Thirteen of these GRs (HarmGR44, 45, 63, 78, 85, 88, 103, 106, 118, 123, 151, 185 and 194) are partial sequences, missing their N- or C-terminus.

Figure 2: Phylogenetic analysis and characterisation of H. armigera GRs.
figure 2

(A) Phylogenetic analysis of selected B. mori and all H. armigera GRs. CO2 GRs (Purple), Sugar GRs (Orange), GR43a-like GRs (dark red) are labelled. For bitter GRs, Type 1 (blue), Type 2 (red) and Type 3 (green) are also labelled. Unknown types (partial sequences and pseudogenes) are labelled in black. Type 2 GRs (HarmGR65) and Type 3 GRs (HarmGR35, 50, 170 and 195) which were used in topological and functional characterization experiments are asterixed. B. mori GRs are framed. (B) The gene structures of Type 1, Type 2 and Type 3 bitter GRs in lepidopteran species. Bars represent exon sequence, lines represent intron sequence. (C) Examples of the distribution and organization of some bitter GRs in H. armigera genome scaffolds.

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We then compared the total numbers of CO2, sugar, GR43a-like and bitter GRs found in the H. armigera genome with those of other Lepidoptera; B. mori, D. plexippus, H. melpomene and P. xylostella (Table 1). The largest difference is in the bitter receptor family, comprising 180 GRs in H. armigera, which is three times the number found in B. mori (51 GRs), H. melpomene (57 GRs), or P. xylostella (49 GRs) and four times more than in D. plexippus (40 GRs)4,11,22,23,27,28. There are also small differences in the total numbers of genes for the remaining GR sub-families. For the CO2 GRs, P. xylostella has five while the other four species possess three (Table 1). The number of sugar GRs varies from four to nine across the five species. There are more GR43a-like GRs in the two butterflies (5 or 6 GRs) and P. xylostella (7 GRs) than in B. mori and H. armigera (2 GRs) (Table 1).

Table 1 Numbers of CO2, sugar, GR43a-like and bitter GRs (Type 1, Type 2 and Type 3) in lepdiopteran species H. armigera, B. mori, D. plexippus, Plutella xylostella and H. melpomene.
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Bitter receptors

To better understand the bitter receptor expansion in H. armigera we performed an in-depth phylogenetic analysis of the subfamily (Fig. 2A). The three B. mori bitter GRs (BmorGR53, 58 and 66) selected for the phylogenetic comparison with H. armigera, represent three different structures of bitter receptors. BmorGR58 contains four exons and three introns and is representative of the majority of other B. mori bitter GRs (Fig. 2A). BmorGR53 is an intronless bitter GR, the only of its kind found in B. mori, while BmorGR66 is a shorter intronless bitter GR4. We were able to group most lepidopteran full-length bitter GRs into three categories based on their lengths and sequences (Fig. 2B and Table 1). Type 1 describes Gr genes, like BmorGR58, that encode approximately 400 amino acids (Fig. S5) and contain 3~4 exons (Fig. 2B). Type 2 comprises a group of long intronless GRs, also encoding approximately 400 amino acids (Fig. S5), like BmorGR5323,27. Type 3 comprises short intronless genes (Fig. 2B and Table 1) encoding 200~350 amino acids (Fig. S5), like BmorGR664. Compared with other lepidopteran species, H. armigera has a low total number of Type 1 GRs but a larger total number of both Type 2 and especially Type 3 bitter GRs (Table 1). Phylogenetically, the H. armigera Type 2 Gr genes cluster together to form a monophyletic branch, while Type 1 and Type 3 are interspersed in the bitter GR sub-family (Fig. 2A). Analysis of their genomic distribution revealed Type 2 or Type 3 GRs are often clustered together (Fig. 2C). For example, 38 Type 3 Gr genes are localized in a tandem array within 0.2 megabases on scaffold 139 (Fig. 2C) and nine Type 2 genes are clustered within 0.05 megabases on scaffold 152 (unpublished data). Five Type 1 and nine Type 3 Gr genes are interspersed with each other on scaffold 88 (Fig. 2C).

As Type 3 Gr genes are short intronless genes, we wanted to verify if these were complete genes and not artefacts of incorrect genome assembly. We performed RT-PCR on 20 of these genes to verify their sequence and expression and show all 20 are as predicted from the genome (data not shown). Within our transcriptome libraries (established from 31 RNA sequencing libraries based on different tissues and life stages, unpublished data), we identified the complete ORF sequences for seven Type 3 Gr genes (HarmGR24, 35, 50, 168, 169, 171 and 174) confirming the genomic sequences are complete. We also used 3′RACE to sequence the ORFs of two Type 3 genes (HarmGR17 and HarmGR19) and confirm the positions of their stop codons are the same as those identified from the genome (S4 Data). Both HarmGR17 and HarmGR19 are highly expressed in female adult heads (Fig. 3). We did not perform 5′ RACE due to the high level of conservation at the 5′ terminal. We also confirmed that the cDNA sequences of the genes used in our functional studies, HarmGR35, 50, 65, 170 and 195 (Fig. S3), matched the sequences annotated from the genome.

Figure 3: H. armigera GR expression as revealed by read mapping.
figure 3

Red box indicates the presence of reads uniquely mapping to the coding region of each receptor gene model while blue box indicates no expression detected. The expression of GRs in various tissues include: (A) embryos (Em), 3rd-instar larvae whole bodies (3rd), 5th-instar larvae whole bodies (5th) and pupae (Pu); (B) larvae antennae (An), mouthparts (Mo), epidermis (Ep), fat body (FB), foreguts (Fg), midgets (Mg), hindguts (Hg), malpigian tubules (Mp), hemocytes (Hc), hearts (He), trachea (Tr), ventral nerve (NV), silk glands (Si), salivary glands (Sa) and muscle (Mu) of 5th instar; (C) male antennae (♂An), female antennae (♀An), male heads with antennae (♂He), female heads with antennae (♀He), male tarsi (♂Ta), female tarsi (♀Ta), male thorax (♂Th), female thorax (♀Th), male abdomens (♂Ab), female abdomens (♀Ab), male testes (♂Te) and female ovaries (♀Ov) from day 0 to day 5 adults.

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GR gene expression profile

To help characterise the potential function of H. armigera bitter receptor candidates, especially members of the expanded Type 3, we built expression profiles for them from 31 transcriptomic libraries (unpublished data) (Fig. 3). We detected the presence of 84 of the total repertoire of 197 GRs. We used a conservative ‘cutoff’ value to judge if a GR is detected or not in the transcriptome libraries, which is based on relative abundance29. As GRs are lowly expressed, it is difficult to assess if non-detection means there is no expression in a tissue or if the level of expression is very low but would result in expression of functional proteins. As there are no biological repeats of the transcriptomic libraries we cannot meaningfully compare expression levels of Gr genes between the different libraries. We can, however, report the number of GRs expressed within these different libraries (Fig. 3A). Across the different developmental stages, four GRs were detected from 3rd instar larvae, twelve were detected from embryos and 5th instar respectively, and eighteen were detected from pupae (Fig. 3A). Across the libraries of 15 different larval tissues (Fig. 3B) and 13 adult tissues (Fig. 3C) we found twenty Gr genes were expressed in larval tissues (Fig. 3B) and 73 Gr genes was detected in adult tissues (Fig. 3C). Most HarmGr genes detected in adults were found in the heads, abdomens or female ovaries (Fig. 3C). Some individual Gr genes were expressed in multiple tissues. For example, HarmGR185 was expressed in nearly all the tested samples except larval antennae (Fig. 3A). HarmGR180 is detected in 22 different tissues. Conversely some individual Gr genes were only detected in specific tissue(s). For example, HarmGR35 was only detected in the adult heads (Fig. 3C), HarmGR65 was detected in only two tissues, the larva fat body (Fig. 3B) and male adult abdomen (Fig. 3C) and HarmGR195 was only detected in adult tarsi (Fig. 3C). Given the low number of cells expressing GRs and the expected low expression levels, it is probable that their expression is not always detectable by transcriptome sequencing.

Topology

Insect GRs have been shown to have a similar topologies to insect ORs, with seven transmembrane domains, an intracellular N-term and extracellular C-term27. As Type 3 receptors are shorter in length and likely to have fewer transmembrane domains than the longer GRs, we analysed their topology. Using the algorithms TMPred, HMMTOP and TMHMM (Fig. S6) we predict that the H. armigera Type 3 bitter receptors have between three and seven transmembrane domains (TMDs. Type 1 and Type 2 GRs are predicted to have five to nine TMDs (Fig. S6). We selected two Type 3 receptors (GR35 and 50) expressed in the male adult head and one Type 2 receptor (GR65) expressed in the fat body, and expressed them as N- and C-terminally MYC-tagged fusions in S2 cells. Untagged receptors were used as controls. In all three cases, strong green immunofluorescence could be visualized from permeabilized cells transfected with either MYC: HarmGR or HarmGR: MYC (Fig. 4). In contrast, when cells were not permeabilised, we observed three different results. No fluorescence was observed on either N- or C-terminally tagged HarmGR35, indicating the tags are intracellular (Fig. 4A). Green fluorescence was seen in cells transfected with both N- and C- terminally tagged HarmGR50, indicating both tags are extracellular (Fig. 4B). Green fluorescence was seen for only C- terminally tagged HarmGR65, HarmGR65: MYC, but not from the N-terminally tagged HarmGR65 (Fig. 4C). These results indicate that HarmGR35, HarmGR50 and HarmGR65 exhibit three different topological structures. The N and C-termini of HarmGR35 are both intra-cellular. The N and C-termini of HarmGR50 are both extra-cellular. HarmGR65, a member of the Type 2 bitter receptor family, predicted to have seven TMDs, has the same topology as BmorGR5327 with an intracellular N-terminus and an extracellular C-terminus.

Figure 4: Topological studies of H. armigera “bitter” receptors.
figure 4

Immunohistochemical localisation of the N-termini and the C-termini of (A) HarmGR35, (B) HarmGR50 and (C) HarmGR65. HarmGRs were expressed in native form or fused with two myc epitopes at its N or C-termini. Cells were labelled with mouse anti-myc and Alexa-labelled anti-mouse antibodies to determine the accessibility of a myc antigen under permeabilised or unpermeabilised conditions. Schematic of expression constructs for untagged Grs (controls), N-terminally MYC-tagged GRs (MYC:GRs) and C-terminally MYC-tagged Grs (GRs:MYC). Note: The actual number of transmembrane domains present is not known. Green indicates MYC-directed Alexa immunofluorescence. Red indicates DAPI nuclear counter stain. Scale Bar = 5 μm.

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Functional characterization

Characterising the function of the entire Type 3 GR category would be extremely challenging due to the labour intensive methods needed for functional studies and the lack of information on possible ligands. We therefore focused on a small number of Type 3 Gr genes to establish whether Type 3 GRs exhibited responses to physiologically relevant ligands. We chose genes expressed in adult tissues relevant to the insect taste system. HarmGR35 and HarmGR50 are expressed in adult male heads (Fig. S3); HarmGR195 is expressed in adult tarsi (Fig. 3) and HarmGR170 is expressed in the adult female abdomen (Fig. S3). We also chose a Type 2 GR (HarmGR65), which is expressed in only 2 libraries, the larval fat body and the male abdomen. Using quantitative calcium imaging, we tested whether, when expressed in Sf9 cells, the receptors would respond to compounds present in crude extracts of host plants. Sf9 cells transfected with an empty expression vector (negative control) showed a low but detectable response to the extract, possibly due to membrane receptors that are expressed natively in the cells. A crude extract of cotton leaves initiated significantly higher responses from cells transfected with HarmGR35, HarmGR50 or HarmGR195, when compared to the negative control, HarmGR65 and HarmGR170 (Fig. 5A). These responses were dose-dependent (Fig. 5E,F) with EC50 values of 0.062 ± 0.006 (SEM, N = 3) extract μg/μL for HarmGR35 and 0.185 ± 0.017 (SEM, N = 3) extract μg/μL for HarmGR50.

Figure 5: Functional characterization of GR35, GR50 and GR195 with leaf extracts and amino acids.
figure 5

(A) Calcium imaging analysis of HarmGR35, HarmGR50, HarmGR65, HarmGR170 and HarmGR195 exposed to a crude extract of cotton leaves. (B) Calcium imaging analysis of HarmGR35 and HarmGR50 in response to crude extract from tobacco leaves. (C) HarmGR195 in response to 50 mM amino acids; proline, glysine, serine, arginine and lysine. (D) Concentration-dependent response curve for HarmGR195 to proline. (E) Concentration-dependent response curves for HarmGR35 and (F) HarmGR50 to crude extract from cotton leave. The average responses of control cells have been subtracted. Error bars indicate the calculated standard divisions of the difference. Analysis of the statistical significance between each response and control was conducted by Dunnett tests as part of one-way ANOVA using arcsine transformation. *p < 0.05.

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For HarmGR35 and 50, we also tested a crude extract of tobacco leaves, since tobacco is also a host for H. armigera. However, the crude extract of tobacco leaves did not trigger significant responses from either HarmGR35 or HarmGR50 (Fig. 5B).

We further tested the HarmGR195 response to the amino acids, proline, glycine, serine, arginine and lysine, as this GR is expressed in adult tarsi (Fig. 3 and Fig. S3), which have been shown by electrophysiological studies to detect amino acids30. We found HarmGR195 is selectively activated by proline (ΔF = 0.106, p < 0.05) but not glycine, serine, arginine or lysine at 50 mM (Fig. 5C). The response to proline is dose-dependent with an EC50 = 43 ± 7 mM (SEM, N = 3) (Fig. 5D).

Discussion

After manual curation of the genome and analysis of H. armigera taste receptor genes, we found an expanded subfamily of gustatory receptors in H. armigera compared with other Lepidoptera. The total number of GRs and ORs may be linked to an insect species’ behaviour and ecology. For example, the honeybee, A. mellifera has an expansion of the olfactory receptor gene family (163 ORs)20 compared with other insects, presumably enhancing its olfactory ability and therefore facilitating the typical foraging and social behaviour of bees20. On the other hand, the honey bee genome contains only 10 Grs20. It has been hypothesized that bees have limited need for Grs for plant secondary metabolite discrimination since flowering plants have evolved visual and olfactory cues to attract bees20. We postulate that the expansion of H. armigera GRs may be linked to this species’ capacity for being a successful generalist as the expansion presumably broadens the range of plant secondary metabolites detected by this species.

We focused our comparison of the GRs on lepidopteran species because of the high degree of amino acid dissimilarity in GRs across even modest evolutionary distances. In comparison with B. mori’s 69 GRs, D. plexippus’s 58 GRs, H. melpomene’s 73 GRs and P. xylostella’s 69 GRs, the nearly three fold expansion of the GR repertoire in H. armigera is mainly in the bitter receptor family. Lepidopteran gustatory receptors have been classified into ‘CO2’, ‘GR43a-like’, ‘sugar’ and ‘bitter’ clades28. While CO2 and ‘sugar’ receptors have been confirmed to respond to CO225, sugars27 and ‘GR43a-like’ have been confirmed to also respond to sugars26, there are no studies confirming that ‘bitter’ receptors respond to bitter tastants. Here we named this large clade of GRs as putative “bitter receptors” because they are grouped in the same branch with B. mori “bitter receptors”11 in the phylogenetic analysis, whose ligands are not yet identified (Fig. S7). From the expression profile on 31 libraries, only half of the total GRs were detected above the conservative cut-off value. Gr gene expression itself is very low and may only be expressed in very few cells within a given tissue hence our conservative cut off may exclude some GRs that are actually functional in the tissue. For example, GR50 is not detected in male heads in our expression profile but can be detected using RT-PCR. When we went back to the raw data we did detect GR50 reads but at a very low level. A further reason for low expression levels in this study may be because lab colonised H. armigera were used in library construction. This colony has been fed on artificial diet for over 10 years which may have resulted in lower GR expression compared with wild insects.

Some GRs are detected in internal tissues like gut, heart and fat body suggesting they may be functioning in internal nutrient detection, as reported in Drosophila31. H. armigera GRs were also detected in embryo stages, suggesting they play a role in insect development as previously reported in Drosophila32.

We further categorised the bitter GRs into three ‘types’ based on gene structure and length. Type 1 exhibit structural features which are conserved across lepidopteran species GRs while Type 2 and Type 3 are intronless GRs that are less commonly found in the available genomes of other Lepidoptera. Type 2 refers to those genes that are relatively long (>400 AA) and Type 3 as those that are relatively short (<360 AA) (Fig. 2). Interestingly, in mammals, intronless genes are shown to be more lowly expressed, present in a narrower range of tissues and evolving faster than intron containing genes33. The majority of the bitter receptors in the H. armigera expansion fall into the Type 3 category.

Type 3 Gr genes are often arranged in the H. armigera genome in gene clusters (Fig. 2), suggesting they arise from a few ancestral genes that have undergone successive duplications. Previous to this study only a limited number of lineage specific expansions in ORs or GRs have been observed in Lepidoptera28. It is also suggested that the common ancestor of the Lepidoptera had a very small number of Gr genes, particularly within the bitter receptor clade, so further analysis of the genomic structure of these clusters in H. armigera may shed light on the evolution of this unique GR expansion.

It would be extremely challenging to comprehensively de-orphan all of these Type 3 GRs, instead we focused on testing whether at least some Type 3 GRs exhibited responses to physiologically relevant ligands. Since there is little behavioural or neurophysiological information on ligands that could activate these receptors, we used crude extractions of leaves from cotton and tobacco, two common host plants for H. armigera that are known to deploy secondary metabolites as defence against herbivores34. Of the five genes we tested, HarmGR35, HarmGR50 (expressed in adult head) and HarmGR195 (expressed in adult tarsi) showed responses to a crude extract of cotton leaves. It is interesting that GR35 and 50, expressed specifically in male head, detect plant compounds. Males also use plant compounds to find food sources and might use host-plant chemical cues to identify the habitat of calling females35.

HarmGR195, which is specifically expressed in the adult tarsi, responded to proline in a dose-dependent manner. Plant nectars may contain up to 2 mM proline36, and this high level of proline is thought to be an attractant, as several species of insects prefer high-proline nectars36. Given that it is highly expressed on the tarsi, HarmGR195 may have a role in regulating the insects’ feeding or oviposition.

Insect GRs are predicted to have seven TMDs, as is typical for insect odorant receptors2,20,37,38 and the topologies of insect GRs are predicted or confirmed to have an intracellular N-terminus and an extracellular C-terminus11,27,37. We found two different topologies for the two Type 3 GRs; HarmGR35, has two intracellular termini, while HarmGR50, has two extracellular termini, indicating both have an even number of TMDs. Type 3 GRs are also predicted to have fewer TMDs, making them the first report of an insect GR family with multiple topologies (Fig. S6).

Previous studies on Drosophila GRs have shown that insect GRs form functional heteromultimers in vivo3,9,12 with genetic studies indicating that co-expression of multiple GRs is essential for the detection of compounds like CO2, sucrose, D-glucose and trehalose3,6,8,9,39. In vitro studies with GRs from B. mori, BmorGR8 and BmorGR9 have shown that responses to myo-inositol or D-fructose do not require the coexpression of other GRs4,27, however these studies do not conclude that GRs can function as homo-oligomers because the lepidopteran-derived Sf9 cells used may express native co-receptors. For example, Orco, the canonical OR co-receptor is expressed in Sf9 cells and probably enables the correct functioning of odorant receptors in Sf9 heterologous assays40. In this study Type 3 receptors were able to function alone in Sf9 cells, however it is possible that multiple GRs are required for function in vivo.

In summary, we have discovered H. armigera has evolved a high number of Gr genes, most of which are intronless and belong to the bitter gustatory receptor clade. We have shown these genes have topologies not previously seen in insect GRs and at least three of them show functional responses that could be related to their feeding behaviour. We propose this expansion in bitter receptors allows H. armigera to detect a broad range of plant secondary metabolites and contributes to the highly successful polyphagous behaviour of this species.

Materials and Methods

Insects and cell culture

H. armigera (CSIRO general laboratory (GR) rearing strain) were fed an artificial diet as previously described41. Spodoptera frugiperda Sf9 and D. melanogaster Schneider S2 cells were cultured at 28 °C as previously described26.

Gene annotation and analysis

Genes encoding GRs were identified from the contigs of the H. armigera genome and transcriptome assemblies, using tBLASTn searches with known D. melanogaster and B. mori GR sequences (Table S1). Amino acid sequences were used for phylogenetic analysis in MEGA 5.142. A maximum-likelihood tree was calculated using default settings and the Jones-Taylor-Thornton (JTT) model with partial deletions and 1000 bootstrap replications. TMpred (http://www.ch.embnet.org/software/TMPRED_form.html)43, HMMTOP (http://www.enzim.hu/hmmtop/)44 and TMHMM (http://www.cbs.dtu.dk/services/TMHMM/)45 were used to predict transmembrane domains.

Gene expression profile

RNAseq libraries were prepared by the Helicoverpa genome consortium in the following tissues: antennae, mouthparts, epidermis, fat body, foregut, midgut, hindgut, malpighian tubules, haemocytes, hearts, trachea, ventral nerve, silk glands, salivary glands and muscle of 5th instars; male antennae, female antennae, male head, female head, male tarsi, female tarsi, male thorax, female thorax, male abdomens, female abdomens, male testes and female ovaries from day 0 to day 5 adults; embryos; and whole bodies of 3rd-instars, 5th-instars and pupae. RNAseq data was pre-processed using the default settings of Just_Preprocess_My_Reads (http://justpreprocessmyreads.sourceforge.net), which conducts mild quality control and trimming, pooled and assembled using Trinity-RNASeq, using the default settings46. Open reading frames (ORFs) were predicted using the TranscriptDecoder software in Trinity-RNASeq (http://sourceforge.net/projects/transdecoder) with the PFAM option. In silico expression profiles were generated using DEW (http://dew.sourceforge.net/), as described29.

Molecular biology

cDNA samples were synthesized and RT-PCR was performed as previously described29. 3′ RACE PCR was performed using a SMART RACE cDNA amplification kit with universal and gene-specific primers (Table S2) according to the manufactures’ manuals. PCR products were purified using QIAquick gel extraction reagents (Qiagen, USA), cloned into the pGEM-T Easy vector (Promega, USA) and subsequently sequenced. Successfully cloned full-length ORF sequences were further amplified using specific primers (Table S2) and cloned into the pIB/V5-His vector for expression in insect Sf9 or S2 cells, followed by immunocytochemical studies and calcium imaging analysis.

Immunocytochemistry

S2 cells were subcultured on poly-L-Lysine coated coverslips in 6-well plates and transfected with 1 μg plasmid [PIB/V5-His vector plus a H. armigera GR (HarmGR) as control, or MYC-epitope tagged HarmGR] with 6 μL of Fugene HD transfection reagent (Promega, USA) in 200 μL of medium per well. Forty-eight hours after transfection, immunofluorescence analysis was performed under permeabilised and non-permeabilised conditions as previously described26.

Calcium Imaging

Sf9 cells were plated into 12-well plates and left to settle for 20 min before being transfected by 500 ng of plasmid (pIB/V5-His vector as control or pIB/V5-His-HarmGR vector) and 3 μL of Fugene HD transfection reagent (Promega, USA) in 100 μL per well of Sf-900 medium (Invitrogen, USA). Forty-eight hours after transfection, cells were prepared for calcium imaging and data analysis as described previously26,40,47. Graphpad Prism5 and Microsoft Excel 2012 were utilized for data analysis.

Tastants

One gram of fresh cotton (Gossypium hirsutum) or tobacco (Nicotiana tabacum) leaves were ground in 4 mL 1× Hank’s balanced salt solution (HBSS) buffer and centrifuged to extract soluble compounds (0.25 μg/μL), which were used directly (or diluted) to characterized selected HarmGRs using calcium imaging analysis. Proline (purity ≥99%) was purchased from Sigma-Aldrich.

Additional Information

How to cite this article: Xu, W. et al. Expansion of a bitter taste receptor family in a polyphagous insect herbivore. Sci. Rep. 6, 23666; doi: 10.1038/srep23666 (2016).