Visual arrestins in olfactory pathways of Drosophila and the malaria vector mosquito Anopheles gambiae C. E. Merrill*, J. Riesgo-Escovar†‡, R. J. Pitts*, F. C. Kafatos§, J. R. Carlson†, and L. J. Zwiebel*§¶ *Department of Biological Sciences, Program in Developmental Biology and Center for Molecular Neuroscience, VU Station B35-1812, Vanderbilt University, Nashville, TN 37235-1812; †Department of Molecular, Cellular, and Developmental Biology, Yale University, New Haven, CT 06520; ‡Department of Developmental Neurobiology, Centro de Neurobiologı´a, Apdo. Postal 1-1141, Quere´taro, Qro 76001, Mexico; and §European Molecular Biology Laboratory, Meyerhofstrasse 1, D-69117 Heidelberg, Germany Edited by John H. Law, University of Arizona, Tucson, AZ, and approved November 7, 2001 (received for review September 24, 2001)
T
he events that initiate and terminate chemosensory signal transduction have received considerable attention in recent years. Olfactory transduction is mediated by G protein-coupled receptor (GPCR) second messenger pathways (1, 2). Olfaction exhibits not only activation but also adaptation, whereby a progressively weaker response is generated to repeated or persistent stimuli. At the molecular level, desensitization of GPCR signaling is brought about by reduced coupling between the receptor and heterotrimeric G proteins. After receptor phosphorylation by G protein-receptor kinases, binding of an arrestin uncouples GPCRs from the signaling cascade (3). Arrestins also have been shown to be involved in GPCR internalization, an integral component of GPCR resensitization in many systems (4). Most arrestins characterized to date have been divided into two broad categories that are largely reflective of their presumptive functional roles (5). Visual arrestins have been reported to display nearly exclusive expression in photoreceptors and interact with rhodopsin to terminate visual stimulation (6, 7). The second general arrestin subtype category comprises the nonvisual arrestins that are expressed in a wide variety of tissues (excluding photoreceptors) where they are presumed to interact with a diverse array of GPCRs (8). In light of the rapid progress in understanding the mechanisms that govern olfaction (9), we have initiated molecular studies to extend this understanding to insect disease vectors. Olfaction principally mediates host preference, a trait that strongly contributes to the ability of Anopheline mosquitoes to act as vectors for malaria or other serious human diseases (10). We now report the cloning and characterization of AgArr1, a novel arrestin family member from Anopheles gambiae sensu stricto (hereafter A. gambiae). Additionally, we show its expression in both olfactory and visual systems. Such an overlap in sensory system expression is verified by studies of two arrestins in Drosophila melanogaster previously thought to be exclusively visual in nature; both Arrestin1 and Arrestin2 (DmArr1 and DmArr2) are expressed not only in photoreceptors, but also in olfactory neurons. By using wellestablished olfactory paradigms in the Drosophila model system, we directly demonstrate the function of arrestins in olfactory signal www.pnas.org兾cgi兾doi兾10.1073兾pnas.022505499
transduction systems. Taken together this report establishes that particular insect arrestins are important components in both visualand nonvisual-signaling pathways. Experimental Procedures Generation and Screening of a Subtracted cDNA Library. Total RNA
was prepared from hand-dissected olfactory tissue (antennae and maxillary palps) or carcasses (bodies stripped of heads including antennae, maxillary palps, and all other appendages) of A. gambiae by using RNeasy kit (Qiagen, Chatsworth, CA). Selective amplification via biotin and restriction-mediated enrichment (SABRE) (11) was used to prepare an ‘‘antennae minus carcass’’ subtracted library. The original protocol was modified by using SuperScript II reverse transcriptase (GIBCO兾BRL) for first-strand cDNA synthesis and by redesigning PCR primers. Purified antennal cDNAs were amplified and subtracted an additional time or subcloned directly into pBluescript II-KS (Stratagene). Initial sampling of 53 insert-containing plasmids by DNA sequencing showed that eight cDNA fragments had significant similarly to arrestins by BLAST (12) analysis. One fragment was chosen for further study based on its strong similarly to a 94-aa stretch of antennal-specific arrestins. Specific primers were designed for both 5⬘ and 3⬘ rapid amplification of cDNA ends (RACE; ref. 13) by using a Marathon cDNA amplification kit (CLONTECH). These were designated Arr5⬘-RACE (5⬘-CTAGTCTCCAGCGATGCCACTGTGTT-3⬘) and Arr3⬘-RACE (5⬘-CAGCTGGGTGTGTGGATGTGGTGC-3⬘). Finally, the complete cDNA representing this A. gambiae arrestin gene was engineered by joining 5⬘ and 3⬘ RACE products before final ligation into pBlueScript II (KS) (Stratagene) and was designated AgArr1.
Sequence Analysis. Sequence comparisons with the DNA Data Base
in Japan兾European Molecular Biology Laboratory (EMBL)兾 GenBank and SWISSPROT databases were performed by using the GCG software (14), and protein alignment was performed by using the CLUSTAL W software package (15).
Reverse Transcription–PCR (RT-PCR). RNA for developmental pro-
files was made as above by using whole bodies at various stages: (i) embryos refer to freshly laid fertilized eggs; (ii) early larvae are 1–2 instars, (iii) late larvae are 3–4 instars, and (iv) pupae refers to pupae taken at least 6 h before eclosion. Total adult A. gambiae RNA was prepared as described above from either female or male
This paper was submitted directly (Track II) to the PNAS office. Abbreviations: GPCR, G protein-coupled receptor; RACE, rapid amplification of cDNA ends; EAG, electroantennogram; EPG, electropalpogram; EA, ethyl acetate; SH3, Src homology 3; BU, butanol; RT-PCR, reverse transcription–PCR. Data deposition: The sequence reported in this paper has been deposited in the GenBank database (accession no. AY017417 for AgArr1). See commentary on page 1113. ¶To
whom reprint requests should be addressed. E-mail:
[email protected].
The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. §1734 solely to indicate this fact.
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Arrestins are important components for desensitization of G proteincoupled receptor cascades that mediate neurotransmission as well as olfactory and visual sensory reception. We have isolated AgArr1, an arrestin-encoding cDNA from the malaria vector mosquito, Anopheles gambiae, where olfaction is critical for vectorial capacity. Analysis of AgArr1 expression revealed an overlap between chemosensory and photoreceptor neurons. Furthermore, an examination of previously identified arrestins from Drosophila melanogaster exposed similar bimodal expression, and Drosophila arrestin mutants demonstrate impaired electrophysiological responses to olfactory stimuli. Thus, we show that arrestins in Drosophila are required for normal olfactory physiology in addition to their previously described role in visual signaling. These findings suggest that individual arrestins function in both olfactory and visual pathways in Dipteran insects; these genes may prove useful in the design of control strategies that target olfactory-dependent behaviors of insect disease vectors.
head appendages (antennae, palps, and proboscis), heads stripped of appendages, or carcasses (bodies stripped of heads). PCRs were carried out with AgArr1 oligonucleotide primers ARR5.0 (5⬘TTAAGGCCATGGTCCAGCAGGGTG-3⬘) and ARR-R5 (5⬘CGTTGCGTCGTCTATTCAAA-3⬘). Expression of ribosomal protein s7 (16) was assayed in parallel reactions by using sequencespecific primers: S7a (5⬘-GGCGATCATCATCTACGTGC-3⬘) and S7b (5⬘-GTAGCTGCTGCAAACTTCGG-3⬘). D. melanogaster RT-PCR reactions were performed on adult RNA generated as above from acetone-fixed, hand-dissected third antennal segments, heads stripped of antennae, and bodies with heads removed. The reactions to amplify DmArr1 used primers DmArra1 (5⬘CAACTCCAACAAGGTGGTGA-3⬘) and DmArra3 (5⬘-GCTTCCAGTTGGGCCTTG-3⬘); DmArr2 used primers DmArrb1 (5⬘-ATGGTGAACGCCCAGTTTAG-3⬘) and DmArrb2 (5⬘GGCGAAGTCCTCGAATACAA-3⬘). PCR for DmKrz cDNA was performed with krzL1 (5⬘-GGTGGTGGTGGAGGAAGTG3⬘) and krzR1 (5⬘-GCTGCTCACCGACTTTGGAT-3⬘) (17). In all reactions, primers for the ribosomal protein 49 gene (rp49a, 5⬘GTATCGACAACAGAGTCGGTCGC-3⬘ and rp49b, 5⬘-TTGGTGAGCGGACCGACAGCTGC-3⬘) were included to simultaneously amplify a constitutive message. Ab Production. Full-length AgArr1 cDNA was cloned into pET15b (Novagen), transformed into BL21 (DE3)pLysS bacteria (Novagen), and purified by using His-Bind affinity resin (Novagen). Polyclonal Abs were generated by using standard immunization protocols. Affinity-purified antisera was obtained by passing a Protein G (Sigma)-purified IgG fraction over an affinity column constructed from purified soluble rAgARR1 immobilized on resin (AffiGel 10兾15, Bio-Rad) according to the manufacturer’s instructions. Immunoblots. Frozen male and female adult A. gambiae (G3 strain) antennae兾palps, heads, and carcasses were hand-dissected over dry ice for preparation of crude total protein extracts by homogenization in 50 l of buffer HmB (PBS兾0.1% SDS兾0.1 M 2-mercaptoethanol); samples were mixed with 10 l of SDS兾PAGE sample buffer (62.5 mM Tris䡠HCl, pH 6.8兾10% glycerol兾2% SDS兾5% 2-mercaptoethanol兾0.05% bromophenol blue) and boiled for 5 min. Samples were then electrophoresed on a 10% SDS兾PAGE gel and transferred to poly(vinylidene difluoride) by using semidry techniques (Bio-Rad). Blots were probed with polyclonal rabbit anti-AgARR1 sera (1:5,000) or rabbit anti-actin sera (1:250, Sigma). Detection was accomplished with alkaline phosphataseconjugated secondary Abs and stained with AP color substrate per the manufacturer’s instructions (Bio-Rad). Duplicate immunoblots were prepared simultaneously for use with either antisera. Immunohistochemistry. Female A. gambiae adults were coldanaesthetized, and heads plus antennae were dissected and fixed 30⬘ in 4% paraformaldehyde兾PBS, washed in PBST (PBS兾0.2% Tween-20), and cryoprotected in 12% sucrose overnight. Heads and antennae were embedded in Tissue-Tek (Lab-Tek) and frozen in liquid nitrogen. Tissue was collected in 9-m sections onto pretreated slides (VWR Scientific), dried at least 30 min, fixed in 4% paraformaldehyde兾PBS 30 min, and washed in PBST. Tissue was blocked with PBSG5 (PBS, 0.2% Tween-20兾3% BSA兾5% normal goat serum) for 1 h under coverslips. Either preimmune or affinity-purified polyclonal serum against full-length rAgARR1 was diluted 1:50 in PBSG5. Antisera were added under a coverslip and incubated overnight at 4°C in humid chambers. Detection was done with VectaStain ABC reagents (Vector Laboratories) per the manufacturer’s instructions and visualized with either a 5-bromo4-chloro-3-indolyl phosphate兾nitroblue tetrazolium substrate (Boehringer Mannheim) for a dark blue stain or with VectorRed for a red precipitate (Vector Laboratories). Microscopy was performed by using an Olympus BX60 microscope connected to the 1634 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.022505499
SPOT camera system (Diagnostic Instruments, Sterling Heights, MI). Drosophila stainings were done as in ref. 19 by using Abs directed against DmArr1 and DmArr2 kindly provided by P. Dolph (Dartmouth University, Hanover, NH) and C. Zuker (University of California, San Diego) (20). Sections and whole mounts were incubated with horseradish peroxidase-conjugated secondary Abs, developed, mounted, and photographed by using a Zeiss Axiophot microscope. Electrophysiology. Electrophysiological recordings of antennae and maxillary palps in D. melanogaster were done with two different setups, as described in refs. 19 and 21. To generate recordings, glass microelectrodes were positioned on the surfaces of these olfactory organs until electrical contact was made. Reference electrodes were inserted into the head capsule. Most experiments were carried out independently in both setups and gave similar results. Dilutions of odorants were in paraffin oil. Insect Stocks and Culture. A. gambiae G3 strain embryos were kindly
provided by Mark Benedict (Centers for Disease Control and Prevention, Atlanta) and reared to adult stage under standard conditions. Adult mosquitoes were maintained in plastic containers at 27°C with 75% relative humidity under a 12:12 photoperiod. For Drosophila, the DmArr1 and DmArr2 mutant alleles and the Df(2L)I131 stock were obtained from P. Dolph and C. Zuker and are described in ref. 20. Dp(2:Y)H3 was obtained from the Indiana University, Bloomington, stock center and is a duplication that covers the DmArr1 locus. Controls were flies heterozygous for the mutations in the cases in which the effects were recessive, or flies heterozygous for the mutation (in the case of arr12) or deficiency, and carrying an extra copy of the gene in a duplication, and the wild-type stock Canton S. All other stocks and balancers used are described in ref. 22.
Results A putative arrestin gene, AgArr1, was isolated as part of a broad screen for olfactory components in A. gambiae, whereby a pool of cDNA fragments representing enhanced antennal expression were subcloned and surveyed by random sequencing. We subsequently used 5⬘ and 3⬘ RACE to complete the sequence of this gene. The total length of AgArr1 encompasses 1,964 bp, of which 1,152 bp constitutes a 383-aa residue ORF followed by a 553-bp 3⬘ untranslated region, including a 40-bp polyadenosine [(poly)A⫹] tract. Multiple 3⬘ RACE clones were sequenced and found to be identical throughout the region preceding their (poly)A⫹ tract, supporting the hypothesis that the reported AgArr1 3⬘ untranslated region sequence is complete. The ORF is preceded by a 5⬘ untranslated region of at least 259 bp, identical in the three independent 5⬘ RACE clones that were sequenced. Although this result provides a high degree of confidence that the common 5⬘ end represents the true start of the AgArr1 mRNA, we cannot exclude the possibility that the actual start is further upstream. The predicted AgARR1 protein has a molecular mass of 42.8 kDa and a pI of 8.0, consistent with all known arrestin subtypes. Sequence Comparisons. A comparison of the deduced amino acid sequence of AgARR1 with other invertebrate and all human arrestin subtypes is shown in Fig. 1. The highest (72%) identity is seen with an arrestin isolated from an antennal cDNA library from the moth Heliothis virescens (HvARRH; ref. 23), followed by 68% identity to the visual subtype DmARR1 (24) and 65% to CvARRA (25), a visual arrestin from the blowfly Calliphora vicina. Following this cluster of homology, the identity between AgARR1 and other invertebrate arrestins ranges from 40% to 48%. When compared to all human arrestin subtypes, AgARR1 remains 38–42% identical. The AgARR1-deduced amino acid sequence contains several domains implicated in arrestin function. These motifs include Merrill et al.
Fig. 1. Arrestin alignment. The CLUSTAL W alignment (12) of all invertebrate and human arrestins. Proteins included in alignment: AgARR1 (AY017417); HvARRH [P55274; (23)]; CvARRA [P51486; (25)]; DmARR1 [P15372; (24)]; DmARR2 [P19107; (37)]; DmiARRB [P19108; (38)]; LmARRH [P32122; (23)]; LpARRA [P51484; (39)]; DmKRZ [AAF32365; (17)]; HsARRC [XP010397; (40)]; HsARRS [P10523; (41)]; HsARRB1 [AAH03636; (18)]; and HsARRB2 [AAH07427; (18)]. Overall homology is indicated by the degree of shading where darker regions denote more conserved domains. Three characters are used above the alignment to mark strongly conserved positions: * indicates a single, fully conserved residue; : indicates one of these strong groups is conserved: STA; NEQK; NHQK; NDEQ; QHRK; MILV; MILF; HY; and FYW whereas a 䡠 indicates one of these weak groups is conserved: CSA; ATV; SAG; STNK; STPA; SGND; SNDEQK; NDEQHK; NEQHRK; FVLIM; and HFY. SH3, SH3 consensus-binding sequences; CB, clathrin-binding domain; arrows, AP-2 basic residues.
potential consensus Src homology 3 (SH3)-binding sites (Fig. 1) (26). Because these regions mediate binding of c-Src to nonvisual arrestins (27), intriguing possibilities have been raised regarding interactions with the mitogen-activated protein kinase pathway. As is the case for most insect arrestins characterized to date, AgARR1 lacks a putative clathrin-binding domain (Fig. 1) thought to be important in aiding receptor endocytosis. Additionally, an interaction with the endocytic adaptor protein AP-2 also has been demonstrated for vertebrate nonvisual arrestins; this association was shown to require specific basic residues in the C terminus, some of which are present in AgARR1 (Fig. 1).
clonal Abs against full-length rAgARR1. Immunoblots (Fig. 2b) revealed that this serum recognizes an ⬇45-kDa protein in crude extracts of both female and male A. gambiae antennae and in heads devoid of olfactory tissue. The apparent size of both the antennal
firm that AgArr1 is present in olfactory tissues (i.e., antennae and maxillary palps) of A. gambiae, as well as to characterize its overall expression profile, RT-PCR analysis was carried out by using embryonic, larval, and pupal developmental life stages of the mosquito, as well as several adult tissues from both female and male A. gambiae (Fig. 2a). AgArr1 mRNA, absent from embryos, can be detected from early larval stages into mature adulthood. Examination of adult tissues shows that AgArr1 mRNA is expressed in adult female and male olfactory appendages. No AgArr1 cDNA was detected in the bodies of either sex (confirmed by Southern blotting, data not shown), whereas substantial AgArr1 template also was detected in heads devoid of olfactory tissues. In these studies, the constitutively expressed ribosomal protein s7 (16) was used as a control for the level of template cDNA. Both AgArr1 and s7 primer sets were designed to span an intron so that mRNA-derived products could be distinguished from contaminating genomic DNA products (Fig. 2a). This analysis (confirmed by sequencing, data not shown) verifies the specific expression of AgArr1 in the developing organism as well as in adult olfactory tissues and heads devoid of olfactory tissue. This result raised the prospect that AgArr1 encodes an arrestin with a broader spectrum of action than a gene with expression exclusively in olfactory tissues. AgArr1 expression was examined at the protein level with polyMerrill et al.
Fig. 2. AgArr1 molecular analysis in A. gambiae. (a) RT-PCR analysis. AgArr1 is expressed in early larval stages through mature adulthood in specific tissues. Antennal- and head-derived RNA demonstrates amplification of AgArr1 in both female and male adult A. gambiae whereas RNA from adult bodies does not. Parallel reactions were performed for the constitutive ribosomal protein s7 gene as a template control. A, antennae; H, heads; B, bodies; *, genomic DNA carried over during RNA preparation. (b) Immunoblot. Polyclonal sera detects AgARR1 protein from crude protein extracts of adult female and male A. gambiae olfactory tissues (antennae and palps) or heads but not bodies. Parallel blots probed with sera against actin (Sigma, 1:250) is a positive control for loading and transfer. PNAS 兩 February 5, 2002 兩 vol. 99 兩 no. 3 兩 1635
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Tissue and Developmental Specificity of AgArr1 Expression. To con-
Fig. 3. In vivo localization of AgArr1 in A. gambiae. Affinity purified polyclonal sera against rAgARR1 specifically labels antennal cell bodies (a and b). AgARR1 antisera also labels retinal photoreceptors (d); preimmune sera does not give comparable signal (c and e). c, cuticle; scb, subsensillar cell bodies; s, sensilla; and pr, photoreceptors. [Scale bars represent 25 m (a–c) or 10 m (d–f ).]
and head forms of AgARR1 conforms well to the 42.8-kDa predicted size of AgARR1. Little, if any, immunoreactivity was observed in extracts from male or female bodies that had been stripped of all appendages or in parallel blots by using appropriate preimmune serum controls (Fig. 2b and data not shown). Probing duplicate blots with polyclonal antisera against a C-terminal fragment of actin (Sigma), which recognizes a 42-kDa actin protein in A. gambiae, confirms that the lack of AgARR1 signal in bodies is not because of insufficient material (Fig. 2b). Immunolocalization of AgARR1 Protein. To fully address the question of AgArr1 tissue specificity, immunolocalization studies were carried out on cryosections prepared from heads and antennae of adult female A. gambiae. These studies directly establish the presence of AgARR1 protein in both olfactory and visual tissues (Fig. 3). Indeed, significant staining for AgARR1 protein appears below the cuticle in subsensillar cell bodies where primary olfactory neurons are expected to lie (Fig. 3 a and c). AgARR1 labeling appears mainly in cell bodies, presumably because of de novo protein synthesis before transport to dendrites. The inability to detect AgARR1 in dendrites is most likely a limitation of light microscopy on the highly cuticularized antennal sensilla of A. gambiae, and similar staining patterns also are observed for Drosophila arrestins (see below). Considerable labeling of AgARR1 protein also is observed in retinal tissue (Fig. 3e); this finding is consistent with the above RT-PCR and immunoblot results from head tissue and implicates AgArr1 in visual transduction. The lack of substantial immunoreactivity in control reactions by using the appropriate preimmune sera (Fig. 3 b and f ) demonstrates that the localizations seen with anti-ARR1 sera are not spurious, but legitimate signals. Arrestin Expression Patterns in D. melanogaster. The unexpected finding that AgArr1 is expressed in both visual and olfactory tissues of A. gambiae led us to speculate that dual sensory expression of arrestins may be a characteristic of Dipteran insects. To test this hypothesis, we conducted studies in the model system D. melanogaster to determine whether the three Drosophila arrestins would also display overlapping sensory expression patterns and to investigate the functional consequence of such a finding. RT-PCR analysis (Fig. 4a) demonstrates that DmArr1 and DmArr2, previously only shown to be visual in function (24, 28), are also detectable in RNA prepared from olfactory organs. Robust signals also are observed from head tissue, owing to their presence in the phototransduction system (Fig. 4a). As before, primers spanned an intron to distinguish cDNA-derived bands from genomic contamination by the size of the expected product. In this case, the larger bands observed in DmArr1 and DmArr2 reactions (asterisks, Fig. 4a) represent amplification of genomic DNA as confirmed by sequencing (data not shown). Lastly, the nonvisual DmKrz arrestin is 1636 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.022505499
Fig. 4. Arrestin expression patterns in D. melanogaster. (a) RT-PCR analysis. Considerable expression of DmArr1 and DmArr2 is noted in reactions performed with antennal RNA; both are also highly detected in RNA derived from heads, whereas little, if any, signal is seen in bodies. DmKrz mRNA is amplified from antennae, heads, and bodies. All reactions simultaneously amplified the ribosomal protein 49 gene as an internal template control. A, antennae; H, heads; B, bodies; *, genomic DNA carried over during RNA preparation. (b) Immunolocalization. Antisera against DmARR1 labels cells in maxillary palps (part 1) and antennae (part 5) of wild-type flies but not in mutant palps (part 2) or antennae (part 6) lacking DmArr1. Sera against DmARR2 also shows staining in wild-type maxillary palps (part 3) but not in palps from DmArr2 mutants (part 4).
expressed in both head and body tissues, consistent with its role in neurogenesis and fat body formation (17). Additionally, DmKrz cDNA products are observed in antennal reactions, indicating that it may also have an olfactory role (Fig. 4a). In all reactions, the ribosomal protein 49 gene was coamplified as an indicator of relative template levels. Immunolocalization of Drosophila visual arrestins by using Abs against either DmARR1 or DmARR2 (20) specifically detected these proteins in olfactory organs (Fig. 4b). In whole-mount staining, DmARR1 and DmARR2 are localized to the ventrolateral surface of maxillary palps (Fig. 4 b1 and b3), corresponding to the location of olfactory neurons (29). Reduced staining observed in palps from flies mutant for DmArr1 (Fig. 4b2) or DmArr2 (Fig. 4b4) demonstrates the specificity of these sera. Additionally, Abs against DmARR1 label a subset of cells in antennal cryosections (Fig. 4b5); antisera for DmARR2 did not yield antennal staining (data not shown). Again, Drosophila carrying a mutant allele of DmArr1 show little, if any, antennal immunoreactivity (Fig. 4b6). In a pattern similar to that observed for AgArr1 in mosquito antennae, the Drosophila signals are distributed in a manner consistent with cell bodies of olfactory neurons. Arrestin Function in Drosophila Olfaction. Arrestin mutants in Dro-
sophila are defective in olfactory physiology, as measured in electropalpogram (EPG) and electroantennogram (EAG) recordings, which detect the summed potentials of olfactory receptor neurons in the maxillary palp and antenna, respectively. The Drosophila arr12 mutant, a missense mutation resulting in ⬍1% of protein levels (20), shows a reduction of ⬇60% in EPG amplitude to the odor of butanol (BU) compared to a control strain (Fig. 5a). To determine whether the defect maps to the DmArr1 locus, we tested arr12 in heterozygous condition with a deficiency chromosome that uncovers the DmArr1 locus and found a similarly reduced amplitude. However, an arr12兾⫹ heterozygote also shows a strong Merrill et al.
amplitude reduction, suggesting that the arr12 mutation is at least partially dominant. We therefore introduced a duplication covering the arr12 locus and showed that it partially rescued the phenotype, providing evidence that the EPG defect does in fact map to the DmArr1 locus. As further evidence that the EPG defect maps to DmArr1, we tested another allele, arr11, a 5-kb insertion in the second intron (20), and found that this mutation, which causes a 90% reduction in protein levels, shows a 40% reduction in amplitude, either in homozygous or heterozygous condition with the deficiency chromosome (Fig. 5b). The reduced EPG phenotype is not restricted to BU; the response to ethyl acetate (EA) is also affected as shown for arr12 and arr12兾Df in Fig. 5c. Mutations in the other visual arrestin gene, DmArr2, showed no detectable alterations in EPG recordings. We then asked whether arrestin function also is required for correct olfactory physiology in the antenna. Although we did not detect EAG abnormalities when either DmArr1 or DmArr2 alleles were tested singly, double mutants showed significant reductions in EAG amplitude. Specifically, arr12;arr23 exhibited a reduced amplitude of response when tested either with BU (Fig. 5d) or EA (Fig. 5e). The arr23 allele is considered one of the strongest because ⬍1% of DmARR2 is detected in arr23 mutants (20). As a test of whether this effect maps to the DmArr2 locus, we also tested arr21, an independently isolated missense allele of DmArr2 that results in ⬇80% of wild-type protein levels (20) and found that arr12; arr21兾arr23 mutants also were defective in response to both odors (not shown). These results suggest that DmArr1 and DmArr2 are both required for normal olfactory physiology. Discussion We have isolated AgArr1, an arrestin from the malaria vector mosquito, A. gambiae and have directly observed its expression in regions of the antenna where olfactory neurons are presumed to reside. These findings support a potential role of AgArr1 in the Merrill et al.
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Fig. 5. Olfactory electrophysiology in Drosophila arrestin mutants. (a) arr12 reduces EPG amplitude in response to 0.5-s pulses of BU, diluted 2 ⫻ 10⫺3 in paraffin oil, measured as described in ref. 19. Df refers to Df(2L)I131 nub b pr. The deficiency uncovers the DmArr1 locus at 36E3 (20). Dp refers to Dp(2; Y)H3 covering the DmArr1 locus (22). ⫹兾⫾ Refers to Canton S, which was used because the parental chromosome for the DmArr1 mutants was not available. For each genotype, n ⫽ 5 flies. Flies were tested 10 times each; mean amplitude per fly was computed, and a mean for each genotype was calculated. Significance was calculated by ANOVA. Error bars indicate SEM (b) EPG amplitudes of arr11 tested with 2 ⫻ 10⫺3 BU, n ⫽ 5 flies. (c) EPG amplitudes of arr12 tested with a 1 ⫻ 10⫺2 dilution of EA, n ⫽ 5 flies. (d) EAG amplitude of arr12; arr23 tested with a 2 ⫻ 10⫺3 dilution of BU, n ⫽ 5 flies. (e) EAG amplitude of arr12; arr23 tested with a 2 ⫻ 10⫺2 dilution of EA, n ⫽ 6.
regulation of olfactory signaling in A. gambiae. Although undetectable in embryonic stages, AgArr1 is expressed early in larval development through mature adulthood, suggesting that this gene may function in sensory or other pathways throughout the mosquito’s life cycle. A surprising discovery during the course of these studies was the detection of AgArr1 in retinal tissues, a finding that suggests AgArr1 could be a more broadly distributed arrestin, which participates in the desensitization of several signaling pathways. This notion led us to question whether expression across multiple sensory systems is typical among insect arrestins or is unique to AgArr1. To address this, we examined several arrestins in a well-characterized insect system, D. melanogaster. Our hypothesis that visual arrestins also may be involved in olfactory responses is supported by the expression of two visual arrestins DmArr1 and DmArr2, as well as the nonvisual DmKrz arrestin, in the antennae. Importantly, a similar overlap between olfactory and visual systems was previously suggested as phospholipase C (norpA) and a phosphatidylinositol transfer protein (rdgB) of D. melanogaster are required for normal performance by both sensory systems (19, 21). Although both insects demonstrate dual sensory expression of arrestins, only Drosophila offers convenient genetic tools to investigate functional significance. We have shown that Drosophila arrestin mutants exhibit a decrease in the amplitudes of electrophysiological responses to olfactory stimuli. Similar amplitude deficits can also be seen in arrestin mutant visual system responses (20). Although the well-established role of arrestins in receptor desensitization (30) might suggest that arrestin mutations would be expected to lead to some kinetic-signaling impairment, such an effect was not detected in these studies. To examine this effect, more sensitive means will be required including single-unit electrophysiology (31) or patch clamp recordings (32). It will be of interest to use these methods to examine the olfactory physiology in arrestin mutants to determine the molecular mechanism(s) that underlie these results, as several explanations seem plausible. Faulty desensitization may elevate basal receptor activity such that the EPG兾EAG amplitude elicited from this higher baseline in mutants would appear lower than the amplitude elicited from flies with wild-type receptor activity levels. Another possibility concerns the role of arrestins in GPCR internalization and resensitization (4). If receptor recycling is compromised by arrestin mutations, then reduced levels of available resensitized receptors would lower the amplitude of response to new stimuli. Recent studies also have found a role for arrestins in mitogen-associated protein kinase signaling pathways by means of SH3 domain-containing proteins (27); if such a transduction cascade is somehow required to maintain olfactory receptor expression or function at the membrane, then loss of arrestins would indirectly affect receptor responsiveness by the loss of this feedback regulation. Although arrestin mutations have been shown to cause neuronal degeneration in the visual system (20, 33, 34), scanning electron microscopy and histological analysis have not revealed degeneration at a gross level in olfactory tissues (data not shown). We note that rdgB mutations of Drosophila affect the physiology of both the visual and olfactory systems, but degeneration is seen only in the visual system (21, 35). The simplest interpretation of the data remains that both DmArr1 and DmArr2 are required for normal olfactory physiology. We have found defects in the physiology of the maxillary palp in mutants of DmArr1, but not DmArr2. Because both genes are expressed in the maxillary palp it is plausible that our inability to detect a functional deficit for DmArr2 in this organ reflects limitations in the sensitivity of our assay, the nature of the available DmArr2 alleles, or a form of redundancy between DmArr1 and DmArr2. Genetic redundancy appears to occur in the antenna, where EAG defects are observed in double, but not single mutants. This redundancy could arise from the ability of either arrestin to act upon the same odor receptor. Alternatively, each arrestin may associate with a different subset of odor receptors, with mutations
of a single arrestin affecting the signaling of only some receptors. A recent study (31) of coding in the antenna has revealed at least three types of neurons that respond to ethyl acetate with distinct spectrums, each presumably expressing a different receptor. Perhaps mutations of a single arrestin affects the signaling of a subset of neurons, and our assay is only sensitive enough to detect the combined loss of neurons associated with DmArr1 and DmArr2. In this context, we note that our inability to detect DmArr2 immunohistochemically in the antenna could reflect either a low level of expression in individual neurons or a small number of neurons in which it is expressed. This study describes visual arrestins that also show expression and function in nonvisual (i.e., olfactory) tissues in Dipteran insects. In addition to previous reports that certain elements involved in activation are common across multiple sensory systems (19, 21), these findings support the idea that components of desensitization are also shared between visual and olfactory pathways. One of the more interesting results in this study is the demonstration that although these elements are shared between pathways, they may be functioning in different manners in each system; olfactory phenotypes display primarily an amplitude deficit whereas the effects on the visual system appear to have both kinetic and amplitude components (20). It will be intriguing to determine the way in which these proteins are functioning in each sensory modality. Moreover, this study demonstrates that arrestins are required for invertebrate olfactory signaling. There is considerable evidence for nonvisual arrestins acting in vertebrate olfactory signal transduction pathways. Abs against -arrestin-2 have been used to label rat olfactory epithelium and incubating -arrestin-2 Abs with olfactory cilia preparations results in a loss of odorant-induced desensitization (and increases in cAMP levels), supporting the idea that vertebrate nonvisual arrestins control olfactory desensitization (36).
Finally, this study identifies a dual sensory modality arrestin from an insect disease vector of significant medical importance; furthermore, this study uses Drosophila as a model system to demonstrate that arrestins previously thought to act solely in phototransduction are not only expressed in other sensory systems but also are required for proper olfactory function. As the arrestin from A. gambiae shows considerable homology and comparable expression patterns to the Drosophila genes, it is likely that AgArr1 may play a similar role in mosquito physiology. In light of the broad distribution and likelihood of involvement in a range of odorant responsiveness, arrestin genes form a particularly attractive target for interference of the olfactory system. In view of the critical role for olfaction in mosquito host choice and vectorial capacity (10), it is tempting to suggest that experimental approaches designed to alter olfactory genes such as AgArr1 could lead to a reduction in the ability of mosquitoes to effectively transmit malaria, West Nile encephalitis, and other globally important medical and veterinary diseases.
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We thank the entire faculty and staff at the European Molecular Biology Laboratory (Heidelberg, Germany) where this work was initiated as well as A. Nicole Fox and other members of the Zwiebel laboratory for helpful discussions. We thank Peter Clyne and Audrey Hing for carrying out supplementary experiments and for discussion. We thank Dr. Claudio Pikielny (University of Medicine and Dentistry of New Jersey–Robert Wood Johnson Medical School, Piscataway, NJ) for comments on this manuscript as well as assistance with subtractive hybridization and Dr. Aileen McAinsh for editorial assistance. This investigation received financial support from the United Nations Development Program兾World Bank兾 World Health Organization Special Program for Research and Training in Tropical Diseases (Grant 980697 to L.J.Z.) and grants from the National Institutes of Health兾National Institute of Deafness and Other Communication Disorders (DC02174–16 to J.R.C. and DC04692–01 to L.J.Z.).
Merrill et al.
