Drosophila gustatory receptors: from gene identification to functional expression

Journal of Insect Physiology 50 (2004) 469–477 www.elsevier.com/locate/jinsphys

Review

Drosophila gustatory receptors: from gene identification to functional expression Sylwester Chyb  Laboratory of Molecular Physiology, Imperial College London, Wye Campus, Wye, Kent TN25 5AH, UK Received 2 February 2004; accepted 22 March 2004

Abstract Recent years have seen long-awaited progress in understanding of the molecular mechanisms of taste perception in insects. The breakthrough came in the early 2000 with the identification of a novel family of candidate gustatory receptor (Gr) genes in the first release of the Drosophila melanogaster genome sequence. The 60 Gr genes are expressed in the subsets of gustatory neurons in the fly’s taste organs and, without exception, encode heptahelical G protein-coupled receptors (GPCRs). Here I review our current knowledge about Gr genes and their products focusing on the newly emerging information regarding the function of the Gr-encoded proteins. # 2004 Elsevier Ltd. All rights reserved. Keywords: Drosophila; Taste; G protein-coupled receptor; GAL4-UAS; Heterologous expression; Calcium imaging

1. Drosophila feeding and chemoreception Drosophila, like many other organisms, base their feeding decisions on the presence or absence of specific volatile and non-volatile chemicals. These signals are detected by chemosensilla, specialized hair-like structures. While the multiporous olfactory sensilla are located on the antennae and maxillary palps (primary and secondary olfactory organs), anatomically different gustatory sensilla are found in numerous locations on the head (primarily on the labellum but also within the food canal), tarsal leg segments, wing edges and, in the case of females, near the ovipositor (Stocker, 1994). All taste sensilla possess a terminal pore through which tastants diffuse into the lymph filling an internal canal. Majority of taste sensilla are innervated by four gustatory neurons, traditionally classified as S, L1, L2 or W responding to sweet, salt and water stimuli, respectively (Dethier, 1976). These bipolar neurons extend their single and unbranched dendrite towards the terminal pore and send an axon to the CNS where the processing of the taste information occurs. 

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0022-1910/$ - see front matter # 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.jinsphys.2004.03.012

Drosophila feed on a semi-liquid diet of fermented fruit and like other flies, exhibit a fascinating pattern of feeding behaviour (Fig. 1). As fruitflies walk on their foodsource, taste sensilla on the tarsi evaluate its chemical content. This is when the fly’s chemosensory apparatus makes the initial contact with potential tastants. If none are detected the fly walks away, but if phagostimulants are present the fly extends its proboscis. The second phase of the gustatory evaluation is performed by the labellar sensilla present near the tip of proboscis. This bilateral structure is formed by paired labella fused to form an opening leading to the food canal. Each labellum is endowed with around 35 taste sensilla arranged in four rows and surrounding the proboscis opening (Fig. 2). The quality of foodstuff is constantly monitored by the labellar sensilla during feeding and, in addition, ingested food makes contact with internal taste organs i.e. neurons of the cibarial organ where further chemosensory evaluation takes place.

2. Identification of Gr genes Until recently very little was known about the molecular basis of taste in Drosophila. Early in the year 2000, Peter Clyne and Carol Warr working in John

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Fig. 1. Drosophila feeding behaviour. Stimulation of tarsal gustatory receptors (left) by sugars present in a semi-liquid diet evokes extension of proboscis and allows contact with labellar sensilla (right) resulting in the opening of the food canal. (Images courtesy of Maryann Frazier and Marta Chyb.)

Carlson’s lab at Yale University used an ingenious computer algorithm to search the 60% complete Drosophila melanogaster genome database. Instead of looking for novel genes sharing homology with the already identified genes their quasi periodic feature classifier (QFC) has been trained to pick novel genes encoding proteins with particular structural features, in this case multiple transmembrane domains. In fact, QFC has been previously used with much success to identify Drosophila olfactory receptor (Or) genes (Clyne et al., 1999 but also see Vosshall et al., 1999). This time, in addition to numerous hormone and neurotransmitter receptors, ion channels, transporters and the Or genes, Carlson’s group identified 42 previously unknown sequences (19 full-length and 23 partial; Clyne et al., 2000). In order to be regarded as taste receptor genes newly identified sequences had to fulfil a second crucial criterion i.e. expression of these genes should be exclusive to taste neurons. In contrast to Or genes which can be readily visualized by the standard in situ hybridization technique, most new probes gave negative results. There were several possible explanations—perhaps lack of detection was due to low transcript numbers, relative inaccessibility of taste neurons or maybe the wrong developmental stage was probed. To overcome this difficulty Clyne et al. (2000) isolated gene transcripts from major taste organs and performed reverse transcription followed by gene amplification, the procedure known as RT-PCR. Almost all (18 out of 19) full-length sequences were expressed exclusively in the labella and/ or tarsi. No signal was detected from pox-n mutants which lack all chemosensilla transformed to mechanosensilla during the development (Nottebohm et al., 1992).

The new family has been named the gustatory receptor (Gr) genes. All sequences are predicted to encode proteins with 7 transmembrane (7TM) domains, a signature motif characteristic of G protein-coupled receptors (GPCR). This finding was not unexpected as there were numerous lines of evidence, which indicated that G proteins participate in insect taste transduction (e.g. Idei et al., 1996). GPCRs are involved in numerous other signalling pathways in many different systems e.g. rhodopsins in visual transduction in mammals and Drosophila (e.g. Zuker, 1996) and in mammalian taste transduction (Adler et al., 2000; Chandrashekar et al., 2000). By all counts, GPCRs constitute one of the larger if not the largest protein families in sequenced genomes of model organisms (over 5% of the nematode C. elegans genes encode GPCRs; Mombaerts, 1999). Two papers published in quick succession in 2001 extended the new family to 54 (Scott et al., 2001) and 56 genes (Dunipace et al., 2001). Additional Gr genes were identified by reiterative BLAST searches of the subsequent releases of Drosophila melanogaster genome. The current complement consisting of 60 Gr genes encoding 68 gustatory receptor (GR) proteins is shown in Table 1. Gr genes are spread around the Drosophila genome (Fig. 3), which along with the relatively conserved position of introns, is indicative of their ancient evolutionary lineage. In fact, the origin of this gene family may predate the origin of the arthropods (Robertson, 2001). However, some Gr genes are clustered in tandem arrays suggesting more recent episodes of gene duplication (Robertson et al., 2003). The nomenclature of Gr genes follows a set of guidelines proposed for the Or genes (Drosophila Odorant Receptor Nomenclature Committee, 2000), which, inciden-

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Fig. 2. Gustatory sensilla on Drosophila labellum. Majority of taste sensilla are found on the tip of proboscis and arranged in four rows (top left panel). Each sensillum houses between two and four gustatory bipolar taste neurons (expressing green fluorescent protein, GFP, top right; confocal image of Voila-GFP courtesy of Dr. Reinhold Stocker). Bottom row shows frontal view of the proboscis tip when food canal opening is closed (left) and during ingestion (right).

tally, is now believed to be a single but highly branched and evolutionarily newer lineage within the Gr family (Robertson et al., 2003). Each Gr gene is named after its numbered cytogenetic location on the Drosophila melanogaster chromosomal map followed by the small letter—‘‘a’’ for single genes (e.g. Gr5a) or ‘‘a’’, ‘‘b’’, ‘‘c’’ and so on for the clustered genes (e.g. Gr6 4a, Gr64b, Gr64c). 3. Gr expression patterns and central projections To overcome difficulties of in situ hybridization and relatively low spatial resolution of RT-PCR, expression patterns of Gr genes have been studied using an elegant technique of GAL4/UAS (Brand and Perrimon, 1993).

This genetic approach is widely used in Drosophila research to target and drive ectopic gene expression. Central to it GAL4 and UAS have been ‘‘borrowed’’ from yeast where they control gene expression: GAL4 is a transcription factor, which interacts with upstream activating sequence (UAS) and together they drive expression of any gene that happens to be downstream of the UAS. A simple genetic cross is performed between two transgenic Drosophila strains, one carrying a construct containing a promoter of choice controlling GAL4 gene (promoter-GAL4), the other containing a UAS sequence fused to a transgene of choice (UAStarget gene). As a result, in the progeny of such a cross, expression of GAL4 driven in a spatial and temporal pattern typical to the promoter used, turns on a pattern

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Table 1 Drosophila gustatory receptor genes, Gr, and their products showing name of the gene, transcript, chromosome location, number of introns, number of amino acid residues, molecular weight, length of the N-terminus sequence and membership of the Gr subfamily (if known) Name

Transcript

Location

Introns

Residues

Molecular weight

N-term

Subfamily

Gr2a Gr5a Gr8a Gr9a Gr10a Gr10b Gr21a Gr22a Gr22b Gr22c Gr22d Gr22e Gr22f Gr23a

CG18531 CG15779 CG15371 CG32693 CG32664 CG12622 CG13948 CG31662 CR31931 CG31929 CR31930 CG31936 CG31932 CG15396

2A3 5A12 8D9 9B1 10B10 10B10 21D2 22B2 22B2 22B2 22B2 22B2 22B2 23A2

3 6 3 2 1 2 3 1 1 1 1 1 1 3

Gr28a Gr28b

CG13787 CG13788

27F6 27F6

2 2

Gr32a Gr33a Gr36a Gr36b Gr36c Gr39a

CG14916 CG17213 CG31747 CG31744 CG31748 CG31622

32D4 33D1 36C2 36C2 36C2 39C1

3 4 2 2 2 3

Gr39b Gr43a Gr47a Gr47b Gr57a Gr58a Gr58b Gr58c Gr59a Gr59b Gr59c Gr59d Gr59e Gr59f Gr61a Gr63a Gr64a Gr64b Gr64c Gr64d Gr64e Gr64f Gr66a Gr68a Gr77a Gr85a Gr89a Gr92a Gr93a Gr93b Gr93c

CG31620 CG1712 CG12906 CG30030 CG13441 CG30396 CG13495 CG13491 CG30189 CG30191 CG30186 CG30330 CG33151 CG33150 CG13888 CG14979 CG32261 CG32257 CG32256 CG33157 CG33157 CG14988 CG7189 CG7303 CG32433 CG31405 CG14901 CG31208 CG13417 CG31336 CG31173

39D3 43B2 47A2 47F5 57B1 58B1 58B1 58A4 59C3 59C3 59C4 59C4 59E3 59E3 61D2 63F5 64A4 64A4 64A4 64A4 64A4 64A4 66C5 68D4 77E4 85F5 89D2 92D1 93F6 93F7 93F7

2 8 1 2 1 1 1 1 1 1 1 1 2 3 7 2 6 7 4 4 8 6 1 0 1 1 1 1 1 1 1

414 444 385 341 408 373 447 394 386 383 387 389 378 370 374 450 452 443 470 440 447 461 475 391 391 390 372 381 381 371 369 427 361 408 416 395 408 412 367 366 397 390 399 406 436 512 456 406 419 429 460 469 530 389 449 397 382 386 419 401 397

47,005 51,155 44,484 39,131 48,391 44,030 51,907 46,131 45,462 45,399 44,970 45,560 44,421 41,960 43,587 51,856 51,586 50,492 53,598 53,598 50,491 53,885 55,545 45,372 44,890 45,264 42,820 44,517 44,591 42,447 42,334 48,533 41,541 48,391 47,529 46,262 47,852 48,375 43,667 43,285 46,164 44,823 45,796 47,559 49,919 57,465 52,822 47,488 48,889 50,228 54,564 55,681 59,442 43,779 51,954 46,080 42,164 45,384 48,833 46,851 46,253

44 56 38 16 20 8 107 16 16 15 11 14 13 26 6 47 28 53 76 51 55 57 41 3 3 5 32 39 44 43 32 2 64 20 16 32 44 37 6 37 3 38 33 36 41 129 61 47 15 23 100 97 49 42 32 29 20 16 62 58 7

I II – – – – – III III III III III III IV IV V V V V V V I – VI VI VI I I I I – – – – – VII VII VII VII VII VI VI VIII VIII II – II II II II II II V I – – – IX IX IX IX

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Table 1 (continued ) Name

Transcript

Location

Introns

Residues

Molecular weight

Gr93d Gr94a Gr97a Gr98a Gr98b Gr98c Gr98d

CG31335 CG31280 CG33083 CG13976 CG31059 CG31060 CG31061

93F7 94D10 97D11 98A2 98A4 98A4 98A4

1 0 1 2 3 3 3

381 404 425 391 403 408 412

44,954 46,857 49,688 45,536 46,851 47,392 47,722

of expression of the chosen target gene. What makes this system so powerful is the fact that promoter-GAL4 and UAS-target gene constructs are initially separated into two distinct transgenic lines: in the GAL4 line, the

N-term 39 11 26 31 45 42 42

Subfamily IX – – – – – –

GAL4 activator gene is present but the target gene is absent, whereas the UAS line contains the target gene but lacks the GAL4. The modular nature of GAL4/ UAS technology allows many different constructs, and

Fig. 3. Localization of Gr genes on Drosophila chromosomes. Gr genes, like Or genes, are spread around the Drosophila genome on chromosomes X, second and third with some Gr genes clustered (e.g. Gr64 cluster contains genes with homology to Gr5a and Gr61a).

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Fig. 4. Research strategy used in functional characterisation of Drosophila Gr genes. Studies of promoter region can lead to insights into spatial and temporal expression, allow genetic or functional ablation of cells expressing given Gr gene as well as enable gene downregulation using RNA interference. Gene transcript can be used to isolate full-length coding cDNA which can be used for heterologous expression and subsequent functional characterisation of the gene product.

in fact more than one at a time, to be expressed in many different restricted sets of tissues (Fig. 4 left). In addition to visualizing spatial and temporal expression pattern using reporter genes encoding GFP (e.g. Fig. 2 top right panel) or b-galactose (lacZ), the GAL4–UAS system is quite commonly used to achieve a number of other effects e.g. genetic or functional ablation (Sweeney et al., 2000) as well as downregulation of gene expression using RNAi (Kalidas and Smith, 2002). However, one should note that identification of the promoter sequences is not straightforward and that the pattern of GAL4 expression does not always match results from in situ hybridization, possibly due to lack of enhancer sequences in the putative promoter region. Gr genes have been found to be expressed in spatially restricted subsets of gustatory neurons (Scott et al., 2001; Dunipace et al., 2001). No Gr gene studied so far is expressed in all taste neurons and Gr genes generally exhibit one of two distinct patterns of expression. The majority of Gr are expressed in a very small number of taste neurons—usually 1–4% of cells (e.g. Gr2a is expressed in only one labellar taste neuron, Gr22f in 3–4 sensilla; Scott et al., 2001). In contrast, a very few genes show a much broader expression pattern e.g. Gr47a and Gr66a expressed in 20% of taste neurons. Some Gr genes, up to 1/3 according to one estimate, are not expressed in taste neurons at all (Dunipace et al., 2001), while at least one gene, Gr22e, is also expressed in the olfactory organs (Scott et al., 2001). According to the estimate provided by Dunipace et al. (2001) each taste sensillum would express at least two Gr genes but given that there are between two and four gustatory neurons associated with each sensillum, the ‘‘one neuron–one

Gr’’ rule could still hold. As expected, all labellar gustatory neurons send axons to the processing centres in the suboesophageal ganglion (SOG) while tarsal and wing taste neurons project to thoracic ganglia (Scott et al., 2001). But while olfactory neurons project to discrete parts of the glomeruli within the antennal lobes, the axons of taste neurons terminate in less defined domains of the SOG (Scott et al., 2001). 4. Functional studies of insect chemoreceptor genes All Gr products are without exception 7TM proteins composed of a single polypeptide chain of between 341 and 530 residues (with predicted molecular weight of between 39 and 59 kDa). Based on their structure, GR proteins encode a novel family of heptahelical receptors exhibiting a considerable diversity. Previous classifications of GPCR families into rhodopsin-like receptor family (class A), secretin/glucagon receptor family (class B), metabotropic glutamate receptor/calcium sensor family (class C) were based on the length of the N-terminus and the site of agonist-binding domain (Bockaert and Pin, 1999; Rang et al., 2003). Within each of these families, proteins exhibit considerable sequence homology while very little homology is seen between families. In contrast, GR proteins have variable but usually relatively short (from 2 to 100 amino acid residues) extracellular N-terminal sequences. There is also a significant level of structural diversity as GR proteins have only between 7% and 40% identical amino acid residues (greater sequence similarities are characteristic for products of clustered genes). The most conserved region of GR proteins is a 33 residue

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signature motif in the 7th transmembrane domain whereas Or genes have additional conserved motifs not present in the GRs (Scott et al., 2001). The 68 GR proteins encoded by the 60 Gr genes may well represent the final count and attention is now turning to the functional characterisation of the Gr-encoded taste receptors. While we have expression data for many Gr genes, to date there have been only a few attempts to functionally express and characterise insect chemoreceptor genes. An approach used with much success in studies of mammalian gustatory receptors is their heterologous expression in cultured cells (Nelson et al., 2001, 2002). A number of conditions have to be fulfilled in the study of a GPCR membrane receptor protein. First, the candidate receptor protein gene has to be subcloned into an appropriate expression vector to be used in the transfection of cultured cells. Second, the receptor protein has to be targeted to the plasma membrane where it needs to achieve proper topology. Last, in order to generate measurable Ca2+ signals, the receptor protein has to couple efficiently to the Gq–mediated PI pathway in the chosen cell type (Fig. 4, right). Full-length coding sequence is usually isolated using a variant of PCR, RACE PCR, and engineered into an expression vector suitable for transfecting chosen cell type. While it is possible to use cells transiently expressing the transfected gene, quite often the use of stable clonal cell lines may be preferred. This however involves initially co-transfecting cells with another vector conferring antibiotic selection, subjecting transfected cells to selective media and selecting cell clones. The level of gene expression can be verified using standard PCR. Targeting of the GR receptor protein to the plasma membrane can be verified by utilising epitope tagging i.e. the addition of an antigenic epitope to a protein. One widely used epitope tag is 11-amino acid long myc-tag, which can be added to either N- or Cterminus. Promiscuous G proteins e.g. Ga15 or Ga16 can be used in cases where the GPCR in question does not couple to the Gq protein responsible for PI pathway and Ca2+ release (Offermanns and Simon, 1995; Milligan et al., 1996). However, it is preferable to do this in transiently expressing cells and/or under the control of an inducible promoter as otherwise celllethal constitutive calcium influx may result. The final challenge is the choice of ligands to be tested. Usually it is a panel of chemicals with clear behavioural effects (Wetzel et al., 2001) but a more comprehensive ligand library with potential agonists grouped according to the number of carbons, degree of saturation, branching and substitution may be tested as well (Araneda et al., 2000). A systematic screen of tastants identified in the literature could provide a good starting point for identification of ligands for Drosophila gustatory receptors.

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5. Drosophila Gr5a encodes trehalose receptor Heterologous expression approach has been recently used by us to characterise Drosophila Gr5a receptor (Chyb et al., 2003). The Gr5a gene is located near the distal tip of the X chromosome where the locus affecting trehalose sensitivity has been previously mapped (Tanimura et al., 1982, 1988). Trehalose is a disaccharide used primarily for storage and transport in many insect species. First, it is present in millimolar concentration in hemolymph where it plays an important role in osmoregulation. Second, as a non-reducing sugar, it prevents glycosylation of hemolymph’s other components. Finally, trehalose is used to transport glucose from fat body to the tissues while in muscles it itself constitutes fuel reserve for (possibly long-range) flight. Interestingly, in Drosophila trehalose seems also to play a significant role in feeding: although their natural diet is fermented fruit, fruitflies ingest the fungi responsible for fermentation. It is therefore no surprise that trehalose, an abundant component of yeast and other fungi, may be a feeding stimulant for these insects. Two groups working independently established that gene Gr5a is required for trehalose detection. First, Ueno et al., (2001) generated deletion lines lacking trehalose sensitivity from a fly strain, EP(X)496 (Ishimoto et al., 2000) by inducing imprecise excision of a transposable element inserted between the Gr5a and Tre1 genes. Two deletion lines obtained from the parental strain, DEP5 and DEP19, show major trehalose feeding defects in a behavioural biossay where flies are offered a choice between trehalose and sucrose. Dahanukar et al. (2001) using Ueno et al.’s deletion lines confirmed the taste defect in both the behavioural assay and by performing electrophysiological recordings from single taste sensilla (tip-recording). In addition, Carlson’s group generated transgene constructs containing Tre1 and/or Gr5a gene(s) and showed that only Gr5a can rescue trehalose sensitivity (incidentally, Tre1 encodes GPCR which has now been implicated in germ cell migration, Kunwar et al., 2003). Persuasive but indirect evidence therefore indicated that Gr5a encodes a candidate taste receptor for trehalose. More direct evidence came from heterologous expression studies where Gr5a has been expressed in Drosophila S2 cells (Chyb et al., 2003). These authors reported two significant findings. Firstly, Gr5a encodes a functional taste receptor protein tuned narrowly and perhaps exclusively to trehalose. The only other functionally characterised sweet taste receptors are mammalian T1R proteins responding to a broad range of sugars, sweet amino acids and sweeteners (Nelson et al., 2002). Interestingly, mammalian sweet taste receptors form heterodimers composed of products of two genes: T1R2 and T1R3. There is no evidence suggesting Gr5a is coexpresed with any other Gr gene but it is possible

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that GR5a forms homodimers. Secondly, Gr5a has a broad expression pattern in labella—no other Gr gene studies so far is expressed in so many labellar taste neurons. Both of these findings have important implications for the logic of taste coding suggesting an existence of a highly specific coding mechanism for trehalose.

6. Perspective A number of challenges await workers in the field of insect taste in the postgenomic era. On one hand, attention seems to be shifting from the molecular identification of taste receptor genes to finding physiological ligands for these receptor proteins. At the same time, however, further molecular studies can help to determine ligand binding sites, transduction cascades and modes of taste receptor protein inactivation. With few exceptions we have very little functional data about Gr receptor proteins (and the situation is not better with Or genes but see Wetzel et al., 2001; Sto¨rtkuhl and Kettler, 2001). In most cases, relevant ligands have not yet been identified and most GR proteins are thus classified as ‘‘orphan receptors’’. Determining the ligand profile (‘‘deorphaning’’ of the receptor) is therefore a major task. Gr5a, the first functionally expressed Drosophila taste receptor gene, shares relatively high similarity (at least by the standards of the Gr family) with Gr61a and the six genes at position 64 (Gr64a–f). In fact, Gr5a is believed to have only recently (on an evolutionary timescale) transposed from chromosome arm 3L to X chromosome (Robertson et al., 2003). Gr5a is 43% identical to Gr64f and 35% identical to Gr64e and regions of extensive homology are found between TM4–7 and second intracellular loop. Given that Gr5a encodes a trehalose receptor could the remaining seven members of this Gr subfamily II encode other sugar receptors? Two more examples of functional analysis of GR proteins come from De Bruyne et al. (2003) and Bray and Amrein (2003). The first group studied Gr21a, one of the Gr genes expressed in antenna. Using GAL4/UAS mediated expression of a toxigene (reaper, rpr) they performed genetic ablation of olfactory neurons expressing Gr21a and found a reduction in the responses to CO2 (but not to any other odorant tested). Thus Gr21a may encode a specific carbon dioxide receptor involved in the detection of sites of fermentation and possibly in protection against well known noxious effects of CO2 (which in the laboratory is preferred to ether for the purpose of anaesthetising flies). The second group studied Gr68a and found that this gene encodes a sex pheromone receptor expressed in the taste neurons of approximately 20 male-specific chemosensilla on the tarsi of the forelegs (Bray and Amrein, 2003). Male

courtship behaviour is significantly affected when these neurons are functionally ablated or the Gr68a transcript is downregulated using RNAi. We have no detailed information as to where the ligand binding domain is located for any GR protein. The N-terminal sequence is generally quite short in these GPCRs and therefore unlikely to contain the agonist binding site. In contrast, vertebrate sweet taste receptors belong to C family of GPCR characterised by a long N-terminal domain where the binding site is believed to be located (e.g. the metabotropic glutamate receptor after which the GPCR family C is named, has 601 amino acids long extracellular sequence containing ligand-binding domain; its total length is 976 residues). It is however quite likely that the N-terminal chain contributes to the formation of the site along with one or more of the extracellular loops. Evidence from other GPCRs reveals that the cytoplasmic loops (and Ctermini) change conformation in response to ligand binding. Comparison between the two common wildtype stocks of Drosophila, Canton-S and Oregon-R, provides the first clue. These flies differ quite markedly in trehalose sensitivity with Oregon-R markedly less sensitive to trehalose than Canton-S. Isono et al. (2002) found that a single nucleotide polymorphism (SNP) in position 218 involving substitution of alanine (A) to threonine (T) correlates with reduced trehalose sensitivity of Oregon R. According to their model, residue 218 is found on the second intracellular loop where it is unlikely to participate in ligand binding but may be involved in interactions with a G protein. At the moment we know very little about G protein selectivity or the mechanism of the signal termination including any role of the phosphorylation in this process. Finally, will the recent progress in Drosophila lead to identification of chemosensory receptors in other insects including pest species? Can we identify taste receptor genes in other insects including economically significant pests or disease-transmitting insect vectors using homology cloning given how much diversity is there within the Gr family? Bioinformatic approaches have been successfully utilized to identify putative odorant and taste receptors in Anopheles gambiae (Hill et al., 2002) and candidate olfactory receptors in the moth, Heliothis virescens (Krieger et al., 2002) and more insect genomes are currently being sequenced. One thing therefore is certain—there will be further new chemosensory receptors awaiting characterisation which makes it a very exciting time to be an insect physiologist.

Acknowledgements I thank J.R. Carlson and H.M. Robinson for discussions and access to unpublished work, and anonymous

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referees and members of my lab for comments on an earlier version of this paper. The work was supported by the BBSRC and Royal Society.

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