The nicotinic acetylcholine receptor gene family of the honey bee, Apis mellifera

doi:10.1101/gr.4549206

Journal List > Genome Res > v.16(11); Nov 2006

Genome Res. 2006 November; 16(11): 1422–1430.

doi: 10.1101/gr.4549206.

PMCID: PMC1626644

The nicotinic acetylcholine receptor gene family of the honey bee, Apis mellifera

Andrew K. Jones,¹ Valerie Raymond-Delpech,² Steeve H. Thany,² Monique Gauthier,² and David B. Sattelle^1,³

¹MRC Functional Genetics Unit, Department of Physiology Anatomy and Genetics, University of Oxford, Oxford OX1 3QX, United Kingdom;

²Centre de Recherches sur la Cognition Animale, Centre National de la Recherche Scientifique, Unité Mixte de Recherche 5169, Université Paul Sabatier, 31062 Toulouse, France

³ Corresponding author.

E-mail david.sattelle/at/anat.ox.ac.uk; fax 44-1865-282-651.

Received August 10, 2005; Accepted November 14, 2005.

Freely available online through the Genome Research Open Access option.

This article has been cited by other articles in PMC.

Abstract

Nicotinic acetylcholine receptors (nAChRs) mediate fast cholinergic synaptic transmission and play roles in many cognitive processes. They are under intense research as potential targets of drugs used to treat neurodegenerative diseases and neurological disorders such as Alzheimer's disease and schizophrenia. Invertebrate nAChRs are targets of anthelmintics as well as a major group of insecticides, the neonicotinoids. The honey bee, Apis mellifera, is one of the most beneficial insects worldwide, playing an important role in crop pollination, and is also a valuable model system for studies on social interaction, sensory processing, learning, and memory. We have used the A. mellifera genome information to characterize the complete honey bee nAChR gene family. Comparison with the fruit fly Drosophila melanogaster and the malaria mosquito Anopheles gambiae shows that the honey bee possesses the largest family of insect nAChR subunits to date (11 members). As with Drosophila and Anopheles, alternative splicing of conserved exons increases receptor diversity. Also, we show that in one honey bee nAChR subunit, six adenosine residues are targeted for RNA A-to-I editing, two of which are evolutionarily conserved in Drosophila melanogaster and Heliothis virescens orthologs, and that the extent of editing increases as the honey bee lifecycle progresses, serving to maximize receptor diversity at the adult stage. These findings on Apis mellifera enhance our understanding of nAChR functional genomics and provide a useful basis for the development of improved insecticides that spare a major beneficial insect species.

The honey bee, Apis mellifera, is an important beneficial insect in agriculture. In addition to producing honey and beeswax, the contribution of A. mellifera to crop pollination is valued at more than $14 billion dollars per year in the U.S. alone (United States Department of Agriculture http://www.ars.usda.gov/main/main.htm). Honey bees live in societies of considerable complexity and thus are studied as models for social behavior (Robinson et al. 1997).

The neonicotinoids are the newest major group of insecticides, which includes acetamiprid, clothianidin, dinotefuran, imidacloprid, nitenpyram, thiacloprid, and thiamethoxam (Tomizawa and Casida 2005). The worldwide annual sales of neonicotinoids amounts to ~1 billion dollars, and they are used against piercing-sucking pests (aphids, leafhoppers, and white-flies) of major crops. In France, the use of imidacloprid has been suspended over concerns that it may be having a drastic effect on bee populations (http://www.pan-uk.org/press/pr140604.htm), highlighting the importance that effective insecticides should also show selectivity within insects so that pollinators such as A. mellifera are spared. While the link between imidacloprid use and bee population decline has yet to be proven, studies have shown that imidacloprid is highly toxic to A. mellifera (Suchail et al. 2004) and at sublethal doses can alter honey bee foraging and learning (Guez et al. 2001; Lambin et al. 2001; Decourtye et al. 2004). Neonicotinoids act as agonists on their molecular targets, nicotinic acetylcholine receptors (nAChRs) (Matsuda et al. 2001), which are prototypical members of the cys-loop ligand-gated ion channel (LGIC) superfamily (Karlin 2002). The fast actions of acetylcholine (ACh) at synapses are mediated by nAChRs, which consist of five homologous subunits arranged around a central ion channel (Corringer et al. 2000; Unwin 2005). Analyses of completed genomes have revealed diverse nAChR gene families with mammals possessing 16 subunit genes, chicken, 17 (Millar 2003), Fugu rubripes, 28 (Jones et al. 2003), and Caenorhabditis elegans, at least 27 (Jones and Sattelle 2004). In contrast, Drosophila melanogaster and Anopheles gambiae have notably smaller nAChR gene families, each consisting of 10 subunits (Jones et al. 2005; Sattelle et al. 2005).

To date, four A. mellifera nAChR subunits (Apisα2, Apisα3, Apisα7-1, and Apisα7-2) have been identified (Thany et al. 2003, 2005), which are expressed in brain structures that play roles in learning and memory, olfactory signal processing, mechanosensory antennal input, and visual processing. These findings are consistent with ACh being a major excitatory neurotransmitter in the insect nervous system (Breer and Sattelle 1987; Lee and O'Dowd 1999). Patch clamp studies have demonstrated the existence of a distinct nAChR subtype in the honey bee nervous system that is blocked by the nAChR antagonists α-bungarotoxin (α-Btx), dihydroxy-β-erythroidine and methyllycaconitine, while nicotine and imidacloprid acted as partial agonists on this receptor (Goldberg et al. 1999; Déglise et al. 2002; Wustenberg and Grunewald 2004). Another study has shown the presence of two nAChR populations that differ in their responses to imidacloprid but not ACh (Nauen et al. 2001). The involvement of nAChRs in honey bee behavior has also been investigated. Injection of the nAChR agonist, nicotine, showed that potentiation of the cholinergic system improves short term memory (Thany and Gauthier 2005) and injection of the nAChR antagonist, mecamylamine, inhibited olfactory learning or memory recall depending upon the site of injection (Lozano et al. 1996, 2001). Recently it has been demonstrated that one distinct nAChR sub-type, which is α-Btx sensitive, is involved in long-term memory, whereas a second subtype, which is α-Btx insensitive, but is affected by mecamylamine, plays a role in retrieval processes (Dacher et al. 2005). Interestingly, this mirrors to a certain extent the mammalian central nervous system, where there are two predominant nAChR subtypes, the α7 and α4β2 receptors, that are α-Btx sensitive and insensitive, respectively, and both receptor subtypes play a role in memory (for review, see Hogg et al. 2003). Since individual nAChR subunits can confer distinct pharmacological properties on a receptor (Romanelli and Gualtieri 2003), the multiple nAChR subtypes present in the honey bee nervous system are likely to be determined by their subunit composition. Identifying the full complement of honey bee nAChR subunits represents a critical step in understanding the variety of roles played by nAChRs in the honey bee nervous system and the exquisite repertoire of bee behavior, as well as in identifying particular targets of chemical compounds. Here we have used the A. mellifera genome to describe the complete honey bee nAChR gene family.

Results

Existence of II candidate nAChR subunit genes in the A. mellifera genome

Using TBLASTN, we identified 11 candidate nAChR subunits in the A. mellifera genome. The complete open-reading frames of each subunit were confirmed and completed RT-PCR and RACE–PCR. An alignment of their protein sequences shows that the honey bee nAChR candidate subunits possess features common to members of the cys-loop LGIC superfamily (Fig. 1

). These include an N-terminal signal peptide sequence, an extracellular N-terminal region with conserved residues in loops A–F that are involved in ACh binding, the dicysteine loop (cys-loop) consisting of two disulphide-bond forming cysteines separated by 13 amino-acid residues, four transmembrane regions (TM1–TM4), and a highly variable intracellular loop between TM3 and TM4. As with other LGIC subunits, the Apis nAChR subunits also possess potential N-glycosylation sites within the extracellular N-terminal domain and phosphorylation sites within the TM3– TM4 intracellular loop. In nine of the candidate subunits, two adjacent cysteine residues that are required for ACh binding (Kao and Karlin 1986) are present in loop C, defining them as α subunits. Due to the absence of the vicinial subunits, the remaining two candidates are designated β subunits.

Figure 1.

Protein sequence alignment of A. mellifera nAChR subunits. Dα1 of D. melanagaster is included for comparison. N-terminal signal leader peptides are underlined, and the loops implicated in ligand binding (LpA-F), as well as the four transmembrane (more ...)

The honey bee nAChR subunits show substantial sequence similarity with known nAChR subunits, in particular, those of other insects. As shown in Table 1, Apis and Drosophila nAChR subunits can share up to 84% amino-acid identity. With regard to vertebrate nAChR subunits, they show 25%–38% identity. A phylogenetic tree demonstrating the relationship between Apis nAChR subunits and those of Drosophila and Anopheles indicates orthologous relationships between the honey bee and fruit fly/ mosquito subunits (Fig. 2

). Previously characterized Apis nAChR subunits were named based on their closest human homolog (e.g., Apisα3 after the human α3 subunit [Thany et al. 2003]). However, several honey bee subunits do not have clear human homologs. Thus, to facilitate comparison of insect nAChR gene families and maintain consistency in insect subunit nomenclature, all honey bee nAChR subunits have been named after their Drosophila and Anopheles counterparts (any alternative nomenclature of Apis subunits is given in Table 1). As with Anopheles (Jones et al. 2005), the Apis counterpart of Dβ2 is of the α type (Amelα8). As is the case for Drosophila and Anopheles (Grauso et al. 2002; Jones et al. 2005; Thany et al. 2005), three Apis subunits (Amelα5, Amelα6, and Amelα7) show close homology with the vertebrate α7 subunit, sharing 34%, 44%, and 40% identity, respectively. Whereas Amelα6 is clearly orthologous to Dα6 (Fig. 2

), the Apis subunits analogous to Dα5 and Dα7 were not so easy to determine. We designated Amelα7 as the Dα7 counterpart based on the fact that its N-terminal extracellular domain resembles that of Dα7 more closely than Dα5, showing 90% and 88% identity, respectively. The third α7-like subunit, Amelα5, is considerably distant from Dα5, as indicated by the long branch in the phylogenetic tree (Fig. 2

Table 1.

Percentage identity/similarity between A. mellifera and D. melanogaster nAChR subunit protein sequences

Figure 2.

Tree showing relationships of D. melanogaster, A. gambiae, and A. mellifera nAChR subunit protein sequences. Numbers at each node signify bootstrap values with 1000 replicates and the scale bar represents substitutions per site. The D. melanogaster GABA (more ...)

Features particular to certain Drosophila and Anopheles nAChR subunits are also evident in their Apis counterparts (Fig. 1

). For instance, as with Dα1, Dα2, Dα3, Dα4, and Dβ2, the corresponding Apis subunits (Amelα1–Amelα4 and Amelα8) have an insertion in loop F, which, interestingly, may contribute to imidacloprid interactions (Shimomura et al. 2004). The Dα1, Dα2, and Dβ2 genes, as well as the Anopheles orthologs, Agamα1, Agamα2, and Agamα8, are similarly arranged and tightly clustered within 200 and 220 kb, respectively (Jones et al. 2005). In the Apis genome, only Amelα1 and Amelα2 are clustered, being situated within 120 kb of each other. Immunohistochemical and coimmunoprecipitation studies show that Dα1, Dα2, and Dβ2 are integral components of certain nAChR subtypes, leading to the suggestion that clustering may facilitate coordinate expression and coassembly of the nAChR subunits (Chamaon et al. 2002). The separation of the Apis Dβ2 ortholog Amelα8 from the cluster may thus result in diversification of receptor expression and coassembly. In line with this potential broadening of receptor complexity, studies indicate that Dβ2 may also be part of a receptor subtype that includes Dβ1 but not Dα1 and Dα2 (Chamaon et al. 2002). Two other subunits, Amelα7 and Amelβ1, lie in close proximity to each other in the A. mellifera genome, both located in linkage group 14.14. This is also the case for the Anopheles orthologs Agamα7 and Agamβ1, which are both on chromosome X at map positions 5D and 5C, respectively, whereas the Drosophila orthologs Dα7 and Dβ1 are located on different chromosomes, X and 3L, respectively (Jones et al. 2005).

Analysis of Drosophila and Anopheles nAChRs shows that each insect possesses a distantly related subunit sharing relatively low-sequence identity with other nAChR subunits. In the case of Drosophila, the subunit is of the non-α type (Dβ3) (Lansdell and Millar 2002), whereas in Anopheles it is an α subunit (Agamα9) (Jones et al. 2005). Interestingly, the honey bee has two distantly related subunits, one α (Amelα9) and the other non-α (Amelβ2), which are designated members of the “Dβ3 Group” (Fig. 2

). It is interesting to speculate that duplication of a common ancestor gave rise to an α and a β subunit with the α subunit being lost in the Drosophila lineage, the β subunit disappearing in the Anopheles lineage, and both subunit types being retained in Apis. Indeed, the Amelα9 and Amelβ2 genes lie only within 10 kb of each other in the honey bee genome, suggesting that both subunits arose from an evolutionary recent duplication event from a common gene.

A comparison of Apis and Drosophila nAChR gene structures shows that only one ortholog pair (Dα6 and Amelα6) shares an identical set of exon–intron boundaries (Fig. 3

). This conservation in gene structure is further highlighted by the Anopheles ortholog, Agamα6, also possessing the same exon composition (Jones et al. 2005). In other cases, Apis nAChR genes possess fewer introns than their Drosophila counterparts (e.g., Amelα1 and Amelβ1, which both possess two less than Dα1 and Dβ1, respectively), more introns (e.g., Amelα3, which has two more than Dα3), or the same number of introns (e.g., Amelα4). Both Amelα5 and Amelβ2 possess an uncommon exon–intron boundary within TM1. It is interesting to observe that in addition to having amino-acid sequences closely resembling the vertebrate α7 subunit, Amelα5, Amelα6, and Amelα7 possess exon–intron junctions found in mammalian, bird, and fish α7, as well as the closely related α8 subunits (Fig. 3

) (Jones et al. 2003), indicating an ancient lineage for this receptor subtype.

Figure 3.

Exon composition of D. melanogaster and A. mellifera nAChR subunits. The N-terminal signal peptide is shown as a bar, the cys-loop is denoted by a star, and the four transmembrane regions are marked as white boxes. Conserved exon–intron boundaries (more ...)

Splice variants increase Apis nicotinic receptor diversity

Two Apis nAChR subunits, Amelα4 and Amelα6, have alternatively spliced exons most likely arising from tandem exon duplication (Kondrashov and Koonin 2001). As with Dα4 and Agamα4 (Lansdell and Millar 2000; Jones et al. 2005), Amelα4 possesses two alternatives for exon 4 (denoted exon4 and exon4′) (Fig. 4A

). However, whereas Dα6 and Agamα6 have two alternatives for exon 3 (Grauso et al. 2002; Jones et al. 2005), Amelα6 has only a single exon. For α6 exon 8, both Apis and Anopheles have two alternatives, while Drosophila has three, although the mosquito possesses exons analogous to Dα6 8b and 8c (Jones et al. 2005), while the honey bee clearly possesses 8a and 8b-like exons (Fig. 4A

Figure 4.

Splice variants of A. mellifera nAChR subunits. (A) Comparison of alternative exons of D. melanogaster and A. mellifera. Apis residues that differ from those of the orthologous Drosophila exon are underlined. (B) Conservation of truncated nAChR transcripts (more ...)

As previously observed for Drosophila nAChRs, alternative splicing introduces amino-acid changes in functionally significant regions (Lansdell and Millar 2000; Grauso et al. 2002). For the two versions of Amelα6 exon 8, residues in the region linking TM2 with TM3 are altered (Fig. 4A

). Since studies using chimeric vertebrate α7/α3 receptors as well as site-directed mutagenesis in α7 have shown that this region is involved in coupling agonist binding to ion channel gating (Campos-Caro et al. 1996), alternative splicing of Amelα6 exon 8 may alter the response of ion channel function upon agonist binding.

Both Dα3 and Agamα3 possess extraordinarily long intra-cellular domains between TM3 and TM4 (Schulz et al. 1998; Jones et al. 2005). However, the Apis ortholog, Amelα3, does not have such an extended region (Fig. 1

), although use of different splice sites gives rise to two variants, Amelα3L (long variant) and Amelα3S (short variant), which have intracellular domains differing in size by 13 amino-acid residues (Fig. 4C

). It is worth noting that Amelα3L has two extra phosphorylation sites. Since phosphorylation of the intracellular loop is involved in regulating several aspects of receptor function such as desensitization and aggregation (Hopfield et al. 1988; Borges and Ferns 2001), the two splice variants have the potential to alter several receptor properties (Schulz et al. 1998; Jones et al. 2005).

Truncated transcripts for several Drosophila nAChR subunits have also been described. For instance, Dα4 cDNAs lacking exon 2 (Dα4^Δexon2) have been identified (Lansdell and Millar 2000), while in other cases, omission of exon 4 from Dα4 (Dα4^Δexon4) and exon 5 from Dα5 (Dα5^Δexon5) result in frameshifts and the introduction of premature stop codons (Lansdell and Millar 2000; Grauso et al. 2002). For Dα7, a premature stop codon is introduced by lack of splicing intron 5 (Grauso et al. 2002). RT– PCR was performed to determine whether similar truncated transcripts could be detected for the corresponding Apis nAChR subunits. As with Anopheles (Jones et al. 2005), truncated honey bee cDNAs analogous to Dα4^Δexon2 and Dα5^Δexon5 were not detected, whereas Amel4^Δexon4 and truncated Amelα7 transcripts were identified, both having premature stop codons (Fig. 4B

). In addition, RT–PCR analysis revealed a novel truncated variant, where lack of splicing intron 9 in Amelα3 results in the introduction of a premature stop codon (Fig. 4C

). It remains to be determined whether these truncated transcripts are removed by a process such as nonsense-mediated decay, which rapidly degrades mRNAs with premature stop codons (Hillman et al. 2004). Otherwise, if the truncated transcripts are translated, it would be of interest to determine the functional role of the resulting proteins. Perhaps they regulate receptor expression in a similar manner to a truncated variant of the mouse α7 subunit, which acts as a dominant negative when cotransfected with full-length α7 in HEK 293 cells (Saragoza et al. 2003). Alternatively, the truncated receptors may be modulating cholinergic synaptic transmission by acting as an ACh “sponge” in a manner similar to that of the molluscan ACh-binding protein (Smit et al. 2003). However, the truncated receptors lack some of the loops important for ligand binding, most notably loop C, which is crucial for ACh interaction; thus, their ability to bind ACh is questionable and remains to be determined. The novel truncated Amelα3 transcript (Fig. 4C

), however, is the first abbreviated insect variant reported to possess all ligand-binding loops as well as the first three trans-membrane domains. With the complete N-terminal extracellular domain as well as TM2, which lines the ion channel, the truncated transcript may well assemble with other subunits to form a functional receptor. Since analysis of subunit mutants suggests a role for TM4 in channel gating (Mitra et al. 2004), it is likely that the Amelα3 variant would have a profound effect on ion channel properties. The Amelα3 truncation also possesses four putative phosphorylation sites (Fig. 4C

); thus, it may serve to diversify several characteristics of receptor function.

The two Amelα4 splice variants are differentially expressed

RT–PCR was performed to determine which of the 11 Apis nAChR subunits as well as all splice variants are transcribed at different stages of honey bee development, including four larval stages (L₀–L₃), three pupal stages (P₁, P₃, and P₄), and the following tissues from adults: mushroom bodies, optic lobes, brains, head, and whole bodies. All 11 subunits, as well as all splice variants, are transcribed in each developmental stage and tissue tested (see Supplemental material) with one exception. Amelα4 exon4 transcripts were detected in all stages and tissues, whereas transcripts of Amelα4 exon4′ splice variants were not observed in larvae and were particularly more abundant in the mushroom bodies, optic lobes, and brain (see Supplemental material). Since alternative splicing of Amelα4 exon4′ substitute residues in the vicinity of the cys-loop, which has been shown to be important for complete receptor assembly (Green and Wanamaker 1997) and radio-ligand-binding assays, indicate that Dα4 with exon 4′ assembles less efficiently than with exon4 (Lansdell and Millar 2000), Amelα4 exon 4′ subunits may serve to modulate receptor assembly during the later stages of honey bee development and in tissues rich in neural activity such as the mushroom bodies and optic lobes.

Amelα6 undergoes A-to-I pre-mRNA editing

Pre-mRNA A-to-I editing modifies select adenosine (A) residues to inosine (I) in transcripts, which is interpreted as guanosine (G), thereby generating mRNA with a nucleotide composition that differs from the corresponding genomic DNA (Seeburg 2002). RNA editing has been observed in several Drosophila nAChR subunits, including two sites in loop D of Dβ1, one site in TM2 of Dβ2, one site in TM3, three sites in TM4 of Dα5, and seven sites in loops E to F in Dα6 (Grauso et al. 2002; Hoopengardner et al. 2003; Sattelle et al. 2005). To determine whether orthologous Apis nAChR subunits are also RNA edited, the equivalent regions of Amelβ1, Amelα8, Amelα5, and Amelα6 were amplified with high-fidelity proofreading DNA polymerase. For Amelβ1, Amelα8, and Amelα5, the sequences of the resulting amplification products were identical to those of genomic DNA with no indication of A-to-G changes (data not shown), showing that these regions of the three subunits are not RNA edited.

For Amelα6, however, six RNA-editing sites were observed, two of which are conserved in the Drosophila and Heliothis virescens orthologs, Dα6 and Hvα7-2, respectively (Grauso et al. 2002) (Fig. 5

). The genomic DNA and adult cDNA sequence traces shown in Figure 5

were taken from the same individual bee, indicating that sequence variation likely arose at the RNA level, thereby eliminating the possibility that they are polymorphisms. Editing at five of the six sites alters amino acid residues, all of which are situated in functionally significant regions. For instance, an N-glycosylation site in loop E is eliminated by one case of editing.

Figure 5.

RNA editing of Amelα6. Sequencing traces of RT-PCR products from larval (L₀), pupal (P₁), and adult stages are compared with amplified genomic DNA. Editing is shown by mixed A and G signals. Amino acids also affected by editing in D. melanogaster (more ...)

Since loop E contributes to ligand binding and N-glycosylation has also been linked to ligand binding as well as channel desensitization and conductance (Corringer et al. 2000; Nishizaki 2003), editing at this site has considerable potential to alter receptor function. In the remaining cases, editing alters residues near or within the cys-loop, which, like alternative splicing of Amelα4 exon4, may affect receptor assembly. Analysis of Amelα6 editing at different stages of honey bee development shows that in larvae, five of the six sites undergo editing, the extent of which increases throughout development so that in adults, four sites are predominantly edited. In pupae, from P₃ onward, editing was observed at the sixth site, which increases considerably the potential diversity of subunit isoforms, as a lysine residue can be converted to either arginine, glutamic acid, or glycine. Interestingly, the elevated editing in the later stages of development is consistent with findings that RNA editing is particularly important in the nervous system function of Drosophila adults (Palladino et al. 2000) and that the highest levels of RNA editing are seen in adult flies (Keegan et al. 2005).

Discussion

We have used the available A. mellifera genome information to complete the characterization of the honey bee nAChR gene family, thus describing the first complete set of Hymenoptera nAChR subunits and the third insect nAChR gene family following those of the two Diptera, A. gambiae (Jones et al. 2005) and D. melanogaster (Sattelle et al. 2005). The three insect species represent ~280 million years of evolution (Carpenter and Burnham 1985; De Gregorio and Lemaitre 2002) where the nAChR gene family has remained compact with A. mellifera having 11 genes encoding nAChR subunits, whereas both D. melanogaster and A. gambiae possess 10 genes (Jones et al. 2005; Sattelle et al. 2005). The nAChR subunit composition of Apis most closely resembles that of Anopheles in that both possess nine α and one β subunit, while Drosophila has seven α and three β. The extra honey bee subunit is a β subunit (Amelβ2) making A. mellifera only the second insect known to possess more than one non-α type subunit.

The characterization of the full complement of honey bee nAChR subunits presents an important basis for associating particular nAChR subtypes with key aspects of behavior, identifying receptor subtypes targeted by neonicotinoids as well as developing insecticides with improved selectivity. Indeed, comparison of complete insect nAChR gene families has identified a highly divergent subunit group (the Dβ3 group) as well as species-specific proteome diversification arising from alternative splicing and RNA editing, all of which represent promising subunit differences to target for future rational insecticide design. While studies using heterologous expression systems such as Xenopus laevis oocytes have proven instructive in characterizing vertebrate nAChRs (Corringer et al. 2000) and low levels of functional expression of an insect α subunit, αL1, have been observed in Xenopus oocytes (Marshall et al. 1990), expression of functional insect nAChRs has so far proven elusive (Sattelle et al. 2005). Nevertheless, Drosophila nAChR α subunits can form robust functional channels when coexpressed with a vertebrate β2 subunit (Bertrand et al. 1994) and studies on such hybrid receptors have provided insights into the selectivity of neonicotinoids for insect nAChRs over those of vertebrates (Matsuda et al. 1998; Ihara et al. 2003), regions of subunit proteins involved in neonicotinoid interactions (Shimomura et al. 2002, 2003, 2004), and the actions of different neonicotinoids (Ihara et al. 2004). Also, computer models of insect nAChRs have been recently generated, which permit docking experiments to assess interactions with compounds of interest (Sattelle et al. 2005). Similar studies combining functional expression with molecular modeling of Apis nAChRs are likely to prove useful in screening for novel compounds that show low selectivity for honey bee receptors and in dissecting the mechanisms of insecticide actions and selectivity on nAChRs.

Methods

Identification of nAChR subunits in the A. mellifera genome

To identify putative nAChR subunits, we screened the A. mellifera genome (database version 34.2b available at http://www.ensembl.org/Apis_mellifera/index.html and assembly version 3.0 available at http://www.ensembl.org/Apis_mellifera/) with each of the 10 D. melanogaster nAChR subunit cDNA sequences using the TBLASTN algorithm (Altschul et al. 1990). Candidate honey bee nAChR subunits were identified based on their considerable sequence homology with previously characterized nAChR subunits (sequences with lowest similarity had E-value 1e-21), particularly at the N-terminal ligand-binding domain and the four transmembrane regions. RT–PCRs were performed to verify the open-reading frame sequences of each subunit. Since BLAST was unable to identify the highly variable N-terminal signal peptides, 5′-RACE, using the Roche 5′/3′ RACE kit, was performed to complete the nAChR subunit sequences.

The multiple protein-sequence alignment was constructed with CLUSTALX (Thompson et al. 1997) using the slow-accurate mode with a gap-opening penalty of 10 and a gap-extension penalty of 0.1 as well as applying the Gonnet 250 protein weight matrix (Benner et al. 1994). The protein alignment was viewed using GeneDoc (http://www.psc.edu/biomed/genedoc). The neighbor-joining method (Saitou and Nei 1987) and bootstrap resampling (Felsenstein 1985), available with the CLUSTALX program, were used to construct a phylogenetic tree, which was then displayed using the TreeView application (Page 1996). Signal peptide cleavage sites were predicted using the SignalP 3.0 server (Dyrlov Bendtsen et al. 2004) and membrane-spanning regions were predicted using the TMpred program (available at http://www.ch.embnet.org/software/TMPRED_form.html). The PROSITE database (Falquet et al. 2002) was used to identify potential cyclic AMP (cAMP), protein kinase C (PKC), CK2, and potential kinase phosphorylation sites.

Dissection of A. mellifera tissues

Honey bee pupae and larvae were taken from the hive. Their developmental stage was determined using pigmentations of eyes, joints, and legs as described by Winston (1987). Adult honey bees were collected at the entrance of the hive. Bees were anaesthetized on ice and dissection of A. mellifera tissues were performed under a stereomicroscope in sterile 1X PBS. The brain was removed from the capsule head free of cuticle and trachea. When necessary, brain parts were separated manually. The tissues were then frozen in liquid nitrogen before RNA and genomic DNA extraction.

Reverse transcription and polymerase chain reaction

Genomic DNA was extracted from adult bees using the DNeasy Tissue Kit (Qiagen) and total RNA was extracted from various developmental stages and tissues using the RNeasy Mini Kit (Qiagen). First-strand cDNA was synthesized from 1 μg of total RNA using Superscript III First-Strand Synthesis Super Mix (Invitrogen). Nested RT–PCR reactions were performed to detect transcript of all honey bee nAChR subunits and variants. Primer pairs that recognize different exons were used to allow identification of cDNA-specific products (see Supplemental material for PCR primer sequences). The PCR reactions were performed in a total volume of 50 μL composed of Taq polymerase and 1X PCR buffer (Sigma), 0.2 mM dNTP mix (Roche), 0.4 μM each primer, and 2 μL first-strand cDNA template. The nested PCR approach involved two reactions each with 30 cycles of 95°C for 30 sec, 55°C for 30 sec, and 72°C for 30 sec/500 bp amplified. The first PCR was used at a final dilution of one in 5000 as template for the second nested PCR reaction. For RNA-editing analysis, nested PCR using the proofreading Pfu Turbo DNA polymerase (Stratagene) in 2 × 30-cycle reactions was performed on at least two independently made first-strand cDNAs. PCR products were analyzed by electrophoresis in a TAE gel and then purified using the QIAquick Gel Extraction Kit (Qiagen) before being sequenced by the dye termination method at the Biochemistry Sequencing Facility, University of Oxford.

Acknowledgments

We are indebted to the A. mellifera Genome Project (Human Genome Sequencing Center), which provided the starting point for this study. We thank Sandrine Paute for technical support. We also thank Ryszard Maleszka and Chris Ponting for encouragement, support, and helpful comments on the manuscript.

Footnotes

[Supplemental material is available online at www.genome.org. Sequence data from this article have been deposited with the EMBL/GenBank Data Libraries under accession nos. DQ026031–DQ026039.]

Article published online before print. Article and publication date are at http://www.genome.org/cgi/doi/10.1101/gr.4549206.

References

Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J. Basic local alignment search tool. J. Mol. Biol. 1990;215:403–410. [PubMed]
Benner, S.A., Cohen, M.A., Gonnet, G.H. Amino acid substitution during functionally constrained divergent evolution of protein sequences. Protein Eng. 1994;7:1323–1332. [PubMed]
Bertrand, D., Ballivet, M., Gomez, M., Bertrand, S., Phannavong, B., Gundelfinger, E.D. Physiological properties of neuronal nicotinic receptors reconstituted from the vertebrate β2 subunit and Drosophila α subunits. Eur. J. Neurosci. 1994;6:869–875. [PubMed]
Borges, L.S., Ferns, M. Agrin-induced phosphorylation of the acetylcholine receptor regulates cytoskeletal anchoring and clustering. J. Cell Biol. 2001;153:1–12. [PubMed]
Breer, H., Sattelle, D.B. Molecular properties and functions of insect acetylcholine receptors. J. Insect Physiol. 1987;33:771–790.
Campos-Caro, A., Sala, S., Ballesta, J.J., Vicente-Agullo, F., Criado, M., Sala, F. A single residue in the M2-M3 loop is a major determinant of coupling between binding and gating in neuronal nicotinic receptors. Proc. Natl. Acad. Sci. 1996;93:6118–6123. [PubMed]
Carpenter, F., Burnham, L. The geological record of insects. Annu. Rev. Earth Planet. Sci. 1985;13:297–314.
Chamaon, K., Smalla, K.H., Thomas, U., Gundelfinger, E.D. Nicotinic acetylcholine receptors of Drosophila: Three subunits encoded by genomically linked genes can co-assemble into the same receptor complex. J. Neurochem. 2002;80:149–157. [PubMed]
Corringer, P.J., Le Novere, N., Changeux, J.P. Nicotinic receptors at the amino acid level. Annu. Rev. Pharmacol. Toxicol. 2000;40:431–458. [PubMed]
Dacher, M., Lagarrigue, A., Gauthier, M. Antennal tactile learning in the honeybee: Effect of nicotinic antagonists on memory dynamics. Neuroscience. 2005;130:37–50. [PubMed]
Decourtye, A., Devillers, J., Cluzeau, S., Charreton, M., Pham-Delegue, M.H. Effects of imidacloprid and deltamethrin on associative learning in honeybees under semi-field and laboratory conditions. Ecotoxicol. Environ. Saf. 2004;57:410–419. [PubMed]
Déglise, P., Grunewald, B., Gauthier, M. The insecticide imidacloprid is a partial agonist of the nicotinic receptor of honeybee Kenyon cells. Neurosci. Lett. 2002;321:13–16. [PubMed]
De Gregorio, E., Lemaitre, B. The mosquito genome: The post-genomic era opens. Nature. 2002;419:496–497. [PubMed]
Dyrlov Bendtsen, J., Nielsen, H., Von Heijne, G., Brunak, S. Improved prediction of signal peptides: SignalP 3.0. J. Mol. Biol. 2004;340:783–795. [PubMed]
Falquet, L., Pagni, M., Bucher, P., Hulo, N., Sigrist, C.J., Hofmann, K., Bairoch, A. The PROSITE database, its status in 2002. Nucleic Acids Res. 2002;30:235–238. [PubMed]
Felsenstein, J. Confidence limits on phylogenies: An approach using the bootstrap. Evolution Int. J. Org. Evolution. 1985;39:783–791.
Goldberg, F., Grunewald, B., Rosenboom, H., Menzel, R. Nicotinic acetylcholine currents of cultured Kkenyon cells from the mushroom bodies of the honey bee Apis mellifera . J. Physiol. 1999;514:759–768. [PubMed]
Grauso, M., Reenan, R.A., Culetto, E., Sattelle, D.B. Novel putative nicotinic acetylcholine receptor subunit genes, Dα5, Dα6 and Dα7, in Drosophila melanogaster identify a new and highly conserved target of adenosine deaminase acting on RNA-mediated A-to-I pre-mRNA editing. Genetics. 2002;160:1519–1533. [PubMed]
Green, W.N., Wanamaker, C.P. The role of the cystine loop in acetylcholine receptor assembly. J. Biol. Chem. 1997;272:20945–20953. [PubMed]
Guez, D., Suchail, S., Gauthier, M., Maleszka, R., Belzunces, L.P. Contrasting effects of imidacloprid on habituation in 7- and 8-day-old honeybees (Apis mellifera). Neurobiol. Learn. Mem. 2001;76:183–191. [PubMed]
Hillman, R.T., Green, R.E., Brenner, S.E. An unappreciated role for RNA surveillance. Genome Biol. 2004;5:R8. [PubMed]
Hogg, R.C., Raggenbass, M., Bertrand, D. Nicotinic acetylcholine receptors: From structure to brain function. Rev. Physiol. Biochem. Pharmacol. 2003;147:1–46. [PubMed]
Hoopengardner, B., Bhalla, T., Staber, C., Reenan, R. Nervous system targets of RNA editing identified by comparative genomics. Science. 2003;301:832–836. [PubMed]
Hopfield, J.F., Tank, D.W., Greengard, P., Huganir, R.L. Functional modulation of the nicotinic acetylcholine receptor by tyrosine phosphorylation. Nature. 1988;336:677–680. [PubMed]
Ihara, M., Matsuda, K., Otake, M., Kuwamura, M., Shimomura, M., Komai, K., Akamatsu, M., Raymond, V., Sattelle, D.B. Diverse actions of neonicotinoids on chicken α7, α4β2 and Drosophila-chicken SADβ2 and ALSβ2 hybrid nicotinic acetylcholine receptors expressed in Xenopus laevis oocytes. Neuropharmacology. 2003;45:133–144. [PubMed]
Ihara, M., Matsuda, K., Shimomura, M., Sattelle, D.B., Komai, K. Super agonist actions of clothianidin and related compounds on the SAD β 2 nicotinic acetylcholine receptor expressed in Xenopus laevis oocytes. Biosci. Biotechnol. Biochem. 2004;68:761–763. [PubMed]
Jones, A.K., Sattelle, D.B. Functional genomics of the nicotinic acetylcholine receptor gene family of the nematode, Caenorhabditis elegans . Bioessays. 2004;26:39–49. [PubMed]
Jones, A.K., Elgar, G., Sattelle, D.B. The nicotinic acetylcholine receptor gene family of the pufferfish, Fugu rubripes . Genomics. 2003;82:441–451. [PubMed]
Jones, A.K., Grauso, M., Sattelle, D.B. The nicotinic acetylcholine receptor gene family of the malaria mosquito, Anopheles gambiae . Genomics. 2005;85:176–187. [PubMed]
Kao, P.N., Karlin, A. Acetylcholine receptor binding site contains a disulfide cross-link between adjacent half-cystinyl residues. J. Biol. Chem. 1986;261:8085–8088. [PubMed]
Karlin, A. Emerging structure of the nicotinic acetylcholine receptors. Nat. Rev. Neurosci. 2002;3:102–114. [PubMed]
Keegan, L.P., Brindle, J., Gallo, A., Leroy, A., Reenan, R.A., O'Connell, M.A. Tuning of RNA editing by ADAR is required in Drosophila . EMBO J. 2005;24:2183–2193. [PubMed]
Kondrashov, F.A., Koonin, E.V. Origin of alternative splicing by tandem exon duplication. Hum. Mol. Genet. 2001;10:2661–2669. [PubMed]
Lambin, M., Armengaud, C., Raymond, S., Gauthier, M. Imidacloprid-induced facilitation of the proboscis extension reflex habituation in the honeybee. Arch. Insect Biochem. Physiol. 2001;48:129–134. [PubMed]
Lansdell, S.J., Millar, N.S. Cloning and heterologous expression of Dα4, a Drosophila neuronal nicotinic acetylcholine receptor subunit: Identification of an alternative exon influencing the efficiency of subunit assembly. Neuropharmacology. 2000;39:2604–2614. [PubMed]
Lansdell, S.J., Millar, N.S. Dβ3, an atypical nicotinic acetylcholine receptor subunit from Drosophila: Molecular cloning, heterologous expression and coassembly. . J. Neurochem. 2002;80:1009–1018. [PubMed]
Lee, D., O'Dowd, D.K. Fast excitatory synaptic transmission mediated by nicotinic acetylcholine receptors in Drosophila neurons. J. Neurosci. 1999;19:5311–5321. [PubMed]
Lozano, V.C., Bonnard, E., Gauthier, M., Richard, D. Mecamylamine-induced impairment of acquisition and retrieval of olfactory conditioning in the honeybee. Behav. Brain Res. 1996;81:215–222. [PubMed]
Lozano, V.C., Armengaud, C., Gauthier, M. Memory impairment induced by cholinergic antagonists injected into the mushroom bodies of the honeybee. J. Comp. Physiol. [A]. 2001;187:249–254.
Marshall, J., Buckingham, S.D., Shingai, R., Lunt, G.G., Goosey, M.W., Darlison, M.G., Sattelle, D.B., Barnard, E.A. Sequence and functional expression of a single α subunit of an insect nicotinic acetylcholine receptor. EMBO J. 1990;9:4391–4398. [PubMed]
Matsuda, K., Buckingham, S.D., Freeman, J.C., Squire, M.D., Baylis, H.A., Sattelle, D.B. Effects of the α subunit on imidacloprid sensitivity of recombinant nicotinic acetylcholine receptors. Br. J. Pharmacol. 1998;123:518–524. [PubMed]
Matsuda, K., Buckingham, S.D., Kleier, D., Rauh, J.J., Grauso, M., Sattelle, D.B. Neonicotinoids: Insecticides acting on insect nicotinic acetylcholine receptors. Trends Pharmacol. Sci. 2001;22:573–580. [PubMed]
Millar, N.S. Assembly and subunit diversity of nicotinic acetylcholine receptors. Biochem. Soc. Trans. 2003;31:869–874. [PubMed]
Mitra, A., Bailey, T.D., Auerbach, A.L. Structural dynamics of the M4 transmembrane segment during acetylcholine receptor gating. Structure. 2004;12:1909–1918. [PubMed]
Nauen, R., Ebbinghaus-Kintscher, U., Schmuck, R. Toxicity and nicotinic acetylcholine receptor interaction of imidacloprid and its metabolites in Apis mellifera (Hymenoptera: Apidae). Pest Manag. Sci. 2001;57:577–586. [PubMed]
Nishizaki, T. N-glycosylation sites on the nicotinic ACh receptor subunits regulate receptor channel desensitization and conductance. Brain Res. Mol. Brain Res. 2003;114:172–176. [PubMed]
Page, R.D. TreeView: An application to display phylogenetic trees on personal computers. Comput. Appl. Biosci. 1996;12:357–358. [PubMed]
Palladino, M.J., Keegan, L.P., O'Connell, M.A., Reenan, R.A. A-to-I pre-mRNA editing in Drosophila is primarily involved in adult nervous system function and integrity. Cell. 2000;102:437–449. [PubMed]
Robinson, G.E., Fahrbach, S.E., Winston, M.L. Insect societies and the molecular biology of social behavior. Bioessays. 1997;19:1099–1108. [PubMed]
Romanelli, M.N., Gualtieri, F. Cholinergic nicotinic receptors: Competitive ligands, allosteric modulators, and their potential applications. Med. Res. Rev. 2003;23:393–426. [PubMed]
Saitou, N., Nei, M. The neighbor-joining method: A new method for reconstructing phylogenetic trees. Mol. Biol. Evol. 1987;4:406–425. [PubMed]
Saragoza, P.A., Modir, J.G., Goel, N., French, K.L., Li, L., Nowak, M.W., Stitzel, J.A. Identification of an alternatively processed nicotinic receptor α7 subunit RNA in mouse brain. Brain Res. Mol. Brain Res. 2003;117:15–26. [PubMed]
Sattelle, D.B., Jones, A.K., Sattelle, B.M., Matsuda, K., Reenan, R., Biggin, P.C. Edit, cut and paste in the nicotinic acetylcholine receptor gene family of Drosophila melanogaster . Bioessays. 2005;27:366–376. [PubMed]
Schulz, R., Sawruk, E., Mulhardt, C., Bertrand, S., Baumann, A., Phannavong, B., Betz, H., Bertrand, D., Gundelfinger, E.D., Schmitt, B. Dα3, a new functional α subunit of nicotinic acetylcholine receptors from Drosophila . J. Neurochem. 1998;71:853–862. [PubMed]
Seeburg, P.H. A-to-I editing: New and old sites, functions and speculations. Neuron. 2002;35:17–20. [PubMed]
Shimomura, M., Okuda, H., Matsuda, K., Komai, K., Akamatsu, M., Sattelle, D.B. Effects of mutations of a glutamine residue in loop D of the α7 nicotinic acetylcholine receptor on agonist profiles for neonicotinoid insecticides and related ligands. Br. J. Pharmacol. 2002;137:162–169. [PubMed]
Shimomura, M., Yokota, M., Okumura, M., Matsuda, K., Akamatsu, M., Sattelle, D.B., Komai, K. Combinatorial mutations in loops D and F strongly influence responses of the α7 nicotinic acetylcholine receptor to imidacloprid. Brain Res. 2003;991:71–77. [PubMed]
Shimomura, M., Yokota, M., Matsuda, K., Sattelle, D.B., Komai, K. Roles of loop C and the loop B-C interval of the nicotinic receptor α subunit in its selective interactions with imidacloprid in insects. Neurosci. Lett. 2004;363:195–198. [PubMed]
Smit, A.B., Brejc, K., Syed, N., Sixma, T.K. Structure and function of AChBP, homologue of the ligand-binding domain of the nicotinic acetylcholine receptor. Ann. N.Y. Acad. Sci. 2003;998:81–92. [PubMed]
Suchail, S., Debrauwer, L., Belzunces, L.P. Metabolism of imidacloprid in Apis mellifera . Pest Manag. Sci. 2004;60:291–296. [PubMed]
Thany, S.H., Gauthier, M. Nicotine injected into the antennal lobes induces a rapid modulation of sucrose threshold and improves short-term memory in the honeybee Apis mellifera . Brain Res. 2005;1039:216–219. [PubMed]
Thany, S.H., Lenaers, G., Crozatier, M., Armengaud, C., Gauthier, M. Identification and localization of the nicotinic acetylcholine receptor α3 mRNA in the brain of the honeybee, Apis mellifera . Insect Mol. Biol. 2003;12:255–262. [PubMed]
Thany, S.H., Crozatier, M., Raymond-Delpech, V., Gauthier, M., Lenaers, G. Apisα2, Apisα7–1 and Apisα7-2: Three new neuronal nicotinic acetylcholine receptor α-subunits in the honeybee brain. Gene. 2005;344:125–132. [PubMed]
Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., Higgins, D.G. The CLUSTAL_X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 1997;25:4876–4882. [PubMed]
Tomizawa, M., Casida, J.E. Neonicotinoid insecticide toxicology: Mechanisms of selective action. Annu. Rev. Pharmacol. Toxicol. 2005;45:247–268. [PubMed]
Unwin, N. Refined structure of the nicotinic acetylcholine receptor at 4A resolution. J. Mol. Biol. 2005;346:967–989. [PubMed]
Winston, M.L. The biology of the honeybee. Harvard University Press; Boston, MA: 1987. Development and nutrition; pp. 46–71.
Wustenberg, D.G., Grunewald, B. Pharmacology of the neuronal nicotinic acetylcholine receptor of cultured Kenyon cells of the honeybee, Apis mellifera . J. Comp. Physiol. A Neuroethol. Sens. Neural. Behav. Physiol. 2004;190:807–821. [PubMed]

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