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Proc Natl Acad Sci U S A. Sep 5, 2006; 103(36): 13526–13531.
Published online Aug 28, 2006. doi:  10.1073/pnas.0601107103
PMCID: PMC1569196

Olfactory responses in a gustatory organ of the malaria vector mosquito Anopheles gambiae


The proboscis is an important head appendage in insects that has primarily been thought to process gustatory information during food intake. Indeed, in Drosophila and other insects in which they have been identified, most gustatory receptors are expressed in proboscis neurons. Our previous characterization of the expression of AgOR7, a highly conserved odorant receptor (OR) of the Afrotropical malaria vector mosquito Anopheles gambiae in the labellum at the tip of the proboscis was suggestive of a potential olfactory function in this mosquito appendage. To test this hypothesis, we used electrophysiological recording and neuronal tracing, and carried out a molecular characterization of candidate OR expression in the labellum of A. gambiae. These studies have uncovered a set of labial olfactory responses to a small spectrum of human-related odorants, such as isovaleric acid, butylamine, and several ketones and oxocarboxylic acids. Molecular analyses indicated that at least 24 conventional OR genes are expressed throughout the proboscis. Furthermore, to more fully examine AgOR expression within this tissue, we characterized the AgOR profile within a single labial olfactory sensillum. This study provides compelling data to support the hypothesis that a cryptic set of olfactory neurons that respond to a small set of odorants are present in the mouth parts of hematophagous mosquitoes. This result is consistent with an important role for the labellum in the close-range discrimination of bloodmeal hosts that directly impacts the ability of A. gambiae to transmit malaria and other diseases.

Keywords: olfaction, proboscis, insect, olfactory receptor neuron

In the Afrotropical malaria vector mosquito Anopheles gambiae and in other insects, olfactory signal transduction is initiated by G protein-coupled receptors (GPCRs) on the dendrites of olfactory receptor neurons (ORNs), which have, thus far, been characterized in several insect species (14). In A. gambiae, 79 GPCR genes hypothesized to encode odorant receptors (AgORs) have been identified (5). Genes encoding candidate odorant receptors (ORs) are diverse, with one notable exception comprising AgOR7 and other members of a highly conserved nonconventional OR subfamily that is widely expressed throughout insect olfactory organs (2, 57). In addition to widespread expression in olfactory organs such as the antennae and maxillary palps of A. gambiae and Dengue virus vector mosquito Aedes aegypti, Ag/AaOR7 has recently been localized to ≈25 distinct type-2 (T2) sensilla on the proboscis (8, 9). In contrast to Ag/AaOR7, their Drosophila ortholog, DOr83b (1), is not expressed in the proboscis of the adult fruit fly. This result suggests that mosquitoes and perhaps other bloodfeeding insects may contain a set of olfactory inputs derived from the proboscis that is absent in other arthropods.

Recent studies have demonstrated that Drosophila Dor83b is not directly responsive to odorants but, rather, is a general component of the olfactory signal transduction machinery (1013). In A. gambiae, AgOR7 expression in the proboscis may, therefore, support similar olfactory capacity, leading to the prediction that the proboscis would be responsive to odorant stimuli and that other conventional odorant-activated AgORs would also be expressed in the proboscis. To test this hypothesis, we conducted electrophysiological experiments to characterize the olfactory responses from the proboscis of A. gambiae to a diverse panel of odorants that included several human sweat compounds (14, 15). Furthermore, we characterized AgOR gene expression in the proboscis and axonal projections to antennal lobes (ALs), a primary olfactory processing center in the insect brain (16). Taken together, the resulting data strongly support the view that the proboscis is an accessory olfactory organ in the malaria vector mosquito.


Characterization of Olfactory Responses from the Proboscis of Female An. gambiae.

We initially used the electrolabellogram (ELG), a transepithelial electrophysiological recording adapted from the well established electroantennogram (EAG) (17) technique, from the surface of the labellum of the proboscis to examine whether this appendage of female adult A. gambiae manifests peripheral olfactory responses to a diverse panel of odorant stimuli (Table 1, which is published as supporting information on the PNAS web site). In these analyses, the mosquito labellum displayed robust olfactory responses to several compounds (Fig. 1), which elicited either fast downward or, occasionally, upward voltage changes. Tested chemicals included butylamine, previously isolated from human effluents (18), butanol, and several short-chain aliphatic carboxylic acids, such as acetic, butylic, isovaleric, oxobutylic, and oxovaleric acids, each resulting in significant ELG responses in the labellum compared with solvent controls (t test, P < 0.01, Fig. 1 C and D). In these studies, several acidic stimuli elicited upward responses in ELG recordings at high (10−2) concentrations (Fig. 1 C and D), whereas control recordings from the shaft of A. gambiae proboscis (arrowhead in Fig. 1A) corresponding to a putative nonolfactory area of the proboscis showed no significant responses to acetic acid (filled arrowhead, Fig. 1 C and D) or other acidic odorants (data not shown). Importantly, as a control, olfactory responses were not observed when the Drosophila melanogaster labellum was tested for sensitivity toward the full spectrum of odorants used in this study (Fig. 1E). This specificity suggests that upward ELG responses elicited in these assays reflect bona fide processes that are due to the excitation of olfactory receptors on the mosquito labellum that are not present in the fly. Furthermore, small but significant downward ELG responses were also observed with ketones, such as acetylpyridine and acetylthiophene, as well as the Henkel 100, a complex odorant mixture (Fig. 1D, t test, P < 0.05).

Fig. 1.
ELG recordings from the epidermis of the proboscis labellum. (A) Scanning electron micrograph of the head and appendages of a female A. gambiae. The dotted box indicates the labellum at the distal tip of the proboscis. The arrowhead depicts the shaft ...

Olfactory responses from a representative T2 sensillum on the labellum of A. gambiae (boxed area Fig. 1A and arbitrarily denoted as S1 in Fig. 1B) were also examined by using single sensillum recordings (SSRs). This sensillum is stereotypically located on the medial portion of the right labellum between the third and fourth mechanosensory hairs (Fig. 1B), closely approximating the vicinity of the ELG recording site (Fig. 1B, arrowhead) that has been shown to house neurons expressing AgOR7 (8). To test for olfactory activity, individual S1 sensilla were examined for physiological responses to a broad range of odorant stimuli where robust responses to several ketone odorants, including acetothiophene, acetylpyridine, acetylthiazole, and acetylphenone, as well as significant responses to butylamine and other acidic compounds were observed (Fig. 2; and see Fig. 5, which is published as supporting information on the PNAS web site). Furthermore, an examination of response amplitudes from representative odorant-induced spike trains revealed the existence of at least two distinct neurons associated with each S1 sensillum. Of these, an identifiable neuron (denoted as a) characterized by spike amplitudes of ≈55 μV responded to acetophenone family compounds butylamine and oxovaleric, oxobutylic, and acetic acids (Fig. 2B Inset). A distinctive b neuron with a larger spike amplitude selectively responded to acetophenone family compounds (Fig. 2B). In contrast, S1 sensillum neurons showed no response to isovaleric acid, which evokes strong ELG responses (Fig. 1) as well as 1-octen-3-ol, 2-methyl phenol, 4-methyl phenol, indole, geranyl acetone, and ammonia, which have been identified as human sweat compounds and shown to elicit significant responses from A. gambiae antennal preparations (14, 15, 1921), (Fig. 1D).

Fig. 2.
SSRs from S1 sensillum on the labellum. (A) Representative 10-s extracellular recording traces from S1 sensillum in the absence of odorant stimulus (spontaneous response rate; mean values ± SE are provided in the adjacent histogram) and in response ...

Central Projections of Proboscis Neurons to the Brain.

Axonal projections to the ALs and subesophageal ganglion (SOG) of the insect CNS convey information associated with distinct chemosensory modalities (22). Indeed, an examination of these diagnostic deutocerebrum axonal targets provides a highly reliable method that has been used to distinguish olfactory from gustatory sensory neurons (23, 24). To determine the projection patterns of proboscis ORNs, anterograde dye-tracing studies from the mosquito proboscis to the CNS were carried out. As expected, these experiments revealed extensive neuronal arborization to the SOG (double arrowhead in Fig. 3 C and D), consistent with a major role for the mosquito proboscis in gustatory processing (25). Importantly, axonal projections (arrowhead in Fig. 3 C and D) derived from proboscis neurons and targeting distinct regions of the ventroposterior domain (posteromedial region of the AL in a ventral section in Fig. 3D) of the ALs were also observed (Fig. 3 C and D, arrow) further validating the presence of a cryptic set of ORNs in this appendage. Although the Drosophila nc82 monoclonal antibody (the kind gift of R. F. Stocker, University of Fribourg, Fribourg, Switzerland) did not adequately outline AL neuropil to define distinct mosquito glomeruli, we were able to partially distinguish the AL glomerular structure as a result of background fluorescence with excitation at 458 nm. In this manner, we were able to identify a region corresponding to at least two (denoted as L1 and L2, respectively, Fig. 3G) posteromedial glomeruli in both ALs that clearly and reproducibly received afferents from proboscis ORNs (arrowheads, Fig. 3 EJ). Of these, the L1 glomerulus appears to be predominantly labeled.

Fig. 3.
Central projection patterns of proboscis neurons to the brain. (A and B) Schematic sagittal (A) and ventral (B) views of the mosquito brain. Each subregion indicates neuropil in the mosquito brain. Pr, protocerebrum; OL, optic lobe. (C) Twelve-micrometer ...

Expression and Localization of the ORs in the Proboscis and Single Sensillum.

Diverse olfactory responses should correlate with the expression of several AgOR family members. To examine this relationship, we dissected whole proboscises, including shafts from male and female heads, and performed a series of nonquantitative RT-PCR analyses (Fig. 6 and Supporting Methods, which are published as supporting information on the PNAS web site) using a set of primers specific to each candidate AgOR (5). In a total of eight experiments (four for each gender), 16 AgORs were reproducibly identified in cDNAs prepared from both male and female proboscises (shaded rows, Table 2, which is published as supporting information on the PNAS web site), and an additional 9 AgORs (shaded rows with asterisk, Table 2), were reproducibly amplified exclusively from female tissue.

In an attempt to further dissect AgOR expression within the proboscis, we used a single sensillum RT-PCR approach to identify AgORs within an individual S1 sensillum on the labellum of the proboscis (see Supporting Methods). Antisense RNA amplification (26) was used to generate sufficient material for cDNA synthesis from individual S1 sensilla after SSR analyses, and AgOR7 expression was used as an assessment of ORN cDNA integrity. In this manner, only cDNA samples positive for AgOR7 were subsequently screened for the presence of other AgORs identified from the aforementioned whole-proboscis RT-PCR studies (Table 2). Of 10 AgOR7-positive S1 sensillum preparations, one OR in particular (AgOR6) was consistently amplified in 6 preparations (arrow in Table 2). In addition, AgOR53 was detected in 3 preparations, AgOR12 and 18 were each detected twice, and seven other AgORs were detected only once (Table 2). Taken together, these data strongly suggest that AgOR6 is expressed in ORNs associated with S1 sensillum on the proboscis of A. gambiae. Double-labeling studies using AgOR6 in situ hybridization coupled with AgOR7 immunostaining were also carried out to more precisely localize AgOR6 transcripts in the labellum of A. gambiae. Here, AgOR6 mRNA was detected in multiple cells throughout the medial portion of the labellum along with AgOR7 protein (Fig. 4). These studies also confirm that AgOR6 mRNA is consistently coexpressed with AgOR7 protein in a subset of labellum ORNs (Fig. 4C, arrowheads), whereas the remainder of ORNs presumably expresses other conventional AgORs. AgOR6 transcripts were also localized to ORNs in the adult antennae (data not shown).

Fig. 4.
Colabeling of AgOR6 and AgOR7 by in situ hybridization and immunostaining on a sagittal section of the labellum. (A) In situ hybridization with antisense AgOR6 probes (red), where AgOR6-positive cells are evident. (B) AgOR7-positive cells detected with ...


Olfactory Responses in the Gustatory Organ of the Mosquito.

This report extends previous studies (8) that localized AgOR7 to the antennae, maxillary palpi, and, unexpectedly, the labellum of the proboscis of A. gambiae, which has, until now, been thought to be an exclusively gustatory sensory organ. Here, ELG analyses revealed significant labial olfactory responses to odorants such as several ketone odorants, butylamine, which has been identified in human skin emanations (18), and other short-chain carboxlyic acids (e.g., acetic, isovaleric, lactic, oxobutylic, and oxovaleric acids). These acidic odorants frequently elicited upward ELG responses at high (10−2) and low (10−4) concentrations (Fig. 1 C and D). To verify whether these ELG responses were the result of nonphysiological artifacts as reported in other insects (27), an identical recording electrode was used to assess olfactory responses from nonchemosensory proboscis regions, such as the shaft (Fig. 1A, arrowhead), where no activity was observed (Fig. 1 C and D, arrowhead). Furthermore, parallel recordings were made from the labellum of Drosophila, where no OR expression has been observed (6) and where, once again, we observed no significant olfactory responses (Fig. 1E). It has been reported that acetic acid elicits downward responses in EAG recordings in A. gambiae (15), whereas upward EAG responses at high doses have been reported in both houseflies (24) and the sable fly, Stomoxys calcitrans (28). However, at lower doses (<0.1-mg application to a filter paper), acetic acid and other acidic compounds (propionic, butylic, and valeric acids) have elicited downward responses in the housefly (27). Moreover, in A. gambiae, the ELG responses to acidic compounds showed a fast rising phase at 10−2 dilutions compared with identical stimuli recorded from nonchemosensory areas in the proboscis shaft (Fig. 1D, arrows and arrowheads, respectively).

Taken together, these results strongly suggest that upward ELG responses to acidic compounds as well as the downward responses (depolarizations) associated with acetylpyridine, acetylthiophene, butylamine, and other compounds in the labellum are bona fide electrophysiological responses from ORNs. Indeed, all of the acidic stimuli identified in this study have been shown to be present in human sweat/skin emanations and, in many cases, have been directly linked to behavioral responses in A. gambiae (15, 2932). The same is true for ammonia, which works synergistically with other odorants to attract A. gambiae (19).

A more detailed characterization of olfactory function in one representative (S1) labellum sensillum is provided when SSR studies are used, where vigorous spike trains to ketones and acidic compounds (Figs. 2 and 6) were observed. In keeping with our ELG data, acetothiophene, acetylpyridine, and butylamine as well as oxocarboxylic and acetic acids elicited robust and characteristic olfactory responses from identifiable S1 neurons. However, in contrast to a significant ELG response, isovaleric acid failed to evoke significant SSR recordings, whereas strong responses were observed for acetylthiazole and acetophenone, two odorants that did not evoke significant ELG deflections. It is not surprising that individual SSR profiles would diverge from more broadly tuned ELG responses that are subject to several variables, including placement of the recording electrode as well as sensilla and ORN population density. Indeed, equivalent dilutions of one odorant, (α-pinene) in Drosophila similarly does not provoke EAG responses (33) but strongly activates at least one specific (ab7a) antennal ORN in SSR studies (34). In general, we note that the large-amplitude S1 sensillum ORN appears to be more specialized as its responses were restricted to acetophenone and other ketone stimuli, whereas the small-amplitude S1 ORN displays more generalized responses to both acidic and ketone odorants as well as robust sensitivity to butylamine.

Overall, the compounds that elicit strong responses from labial sensilla represent odorants of lower volatility relative to those known to evoke responses from the antennae of A. gambiae (14). Indeed, the vapor pressures of these odorants comprise a narrow range [lactic acid 0.08 torr (1 torr = 133 Pa); isovaleric acid 0.36 torr; acetylpyridine 0.37 torr; acetophenone 0.4 torr] relative to odorants that evoke antennal responses, such as ammonia and indole, with vapor pressures of 6,650 and 10.4 torr, respectively. This finding suggests that A. gambiae, and perhaps other hematophagous insects, may use their labella to detect low volatile kairomones, which may be important for orientation behaviors at close proximity. The importance of close-range olfactory cues is not without biological precedent. Indeed, part of the penultimate stage for oviposition site selection occurs at extremely close range in many species of lepidopteran insects, where, presumably, chemosensory information is processed (35). This suggests that these animals obtain critical chemical information at close proximity but without direct contact. Several mosquito species show enhanced attractive orientation flights toward human skin compounds as temperature is increased (36), indicating that human kairomones are likely to be evaporated and detected by the insects. This result is consistent with the hypothesis that a small set of sensory neurons expressing AgOR7 in the proboscis of A. gambiae, and perhaps other mosquitoes, may be important for determining olfactory profiles in close proximity to a host, where they provide critical olfactory information to the mosquito as part of the penultimate steps in alighting, probing, and bloodfeeding behaviors.

Central Projections of Proboscis Neurons to the Antennal Lobes of the Brain.

Anterograde dye fillings from proboscis neurons revealed extensive arborization to a distinct set of at least two glomeruli in A. gambiae ALs (Fig. 3), It is, therefore, possible that, in A. gambiae, these posteromedial glomeruli represent a proboscis-specific projection area. If this region encompasses the entire proboscis projection zone of the A. gambiae AL, where the responses of 24 proboscis AgORs are directed, then it is reasonable to suggest that each of these glomeruli is targeted by multiple AgORs expressing ORNs, implying that a previously uncharacterized mechanism may underlie the encoding of olfactory information from the labellum, where the majority of ORNs express multiple AgORs that target common AL glomeruli. Alternatively, individual labellum ORNs that express distinct AgORs converge onto a restricted number of common AL glomeruli. In any case, the presence of labial projection axons that target the AL provides compelling support for the presence of a cryptic set of ORNs on what is certainly a predominantly gustatory appendage.

Expression and Function of ORs in the Labellum.

Previous studies in Heliothis virescens using whole-appendage expression surveys have revealed the presence of candidate OR transcripts in the proboscis (37, 38). This result is reminiscent of physiological studies in another lepidopteran, where the labial pit organ (LPO) and its projections of Manduca sexta have defined accessory olfactory pathways that are exclusively responsive to CO2 (39), where they play a key role in finding host plants at a distance (40, 41). Interestingly, the LPO is not responsive to other volatile odorants, suggesting that this structure may be functionally more closely related to the maxillary palps in mosquitoes, which are the site of CO2 sensitivity in these insects (42). In this report, we provide a demonstration that fully functional AgORs are expressed in chemosensory sensilla located on the labial portion of the proboscis, which is a predominantly gustatory appendage in A. gambiae. Importantly, OR expression in the proboscis has, until now, not been described in any other dipteran insects, consistent with electrophysiological studies presented here, demonstrating a lack of olfactory sensitivity in the D. melanogaster labellum (Fig. 1E). These findings imply that there may be important functional and organizational differences between the chemosensory processes of A. gambiae and D. melanogaster that may reflect significantly different life-cycle characteristics, including feeding habits, oviposition demands, and other elements. In this light, it is especially tempting to focus on the anautogenous requirement for vertebrate blood that is characteristic of A. gambiae and other hematophagous mosquitoes as a critical distinction between the life cycles of these dipterans.

In A. gambiae, the presence of 24 conventional AgORs has been detected from whole-proboscis RT-PCR screens (Table 2). Among the AgORs identified in the S1 sensillum, AgOR6 is observed in the majority (6 of 10) of these assays, indicating that it is highly expressed in one or more ORN associated with individual S1 sensilla and, moreover, may reasonably be expected to be tuned to one or more of the odorants that evoke the strongest responses in these assays. Analyses of SSR spike train amplitudes (Fig. 2) suggest that two ORNs are likely to be located in the S1 sensillum of the proboscis labellum. Furthermore, our in situ hybridization and RT-PCR data are consistent with the view that AgOR6 is expressed in a subset of AgOR7-positive labellum ORNs, and AgOR6 is one of several AgORs that facilitate olfactory responses in this appendage.

These data argue against our earlier hypothesis that AgOR7 may also function in a gustatory role on the labellum of A. gambiae (43) and, instead, provide compelling evidence for the presence of cryptic ORNs on this mosquito chemosensory appendage. Although these data do not formally rule out a role for AgOR7 in gustation, we favor the hypothesis that AgOR7 is a true homolog to the nonconventional Drosophila Dor83b protein and that, accordingly, its expression defines the majority of ORNs in this system. The strong olfactory responses recorded in this study from the labellum of A. gambiae may convey information that is critical to the later-stage events in bloodfeeding, host preference, and other behaviors of this mosquito and, therefore, may have profound effects on its vectorial capacity.


Insect Preparations.

A. gambiae sensu stricto (G3 strain) were reared as described (3). Nonbloodfed 3- to 4-day-old female mosquitoes were used for electrophysiological recordings and neuroanatomical studies. Before the electrophysiological experiments described below, adult mosquitoes were cooled at 4°C and restrained in a pipette tip, holding head and appendages in place.

Odorant Stimulation.

Odorants (>98% purity) were obtained from Sigma or Aldrich. Henkel 100, which contains 100 different volatile chemicals (44), was supplied by Henkel (Düsseldorf, Germany). Other odorants were chosen on the basis of behavioral and electrophysiological responses to A. gambiae, as shown in previous studies (Table 1). A humidified and purified, continuous air stream (4 ml/s) was delivered to mosquitoes through a glass pipette by using a stimulus controller (Syntech, Hilversum, The Netherlands). Twenty-five milliliters of diluted odorants (10−2 and 10−4, vol/vol) were applied to a filter disk (VWR, West Chester, PA) that was inserted into a Pasteur pipette. Each odorant was delivered in a 0.5-s, air pulse through the Pasteur pipette to the glass pipette.


A restrained mosquito in a pipette tip was positioned on modeling wax (Hygenic, Akron, OH). Antennae, proboscises, and maxillary palpi were carefully attached to Scotch double-stick tape (3M, St. Paul, MN) on a coverglass mounted on modeling wax. A sharp glass recording electrode with an 0.84-mm i.d. with 1–2 MΩ resistance (World Precision Instruments, Sarasota, FL) was prepared by using a horizontal electrode puller (Model P-97; Sutter Instruments, Novato, CA) and filled with 0.1 M KCl. This electrode was placed in contact with the epithelium of the labellum, together with a similarly prepared reference electrode placed on the thorax, as modified from EAG procedures described in ref. 45. Data were imported to a 4-channel IDAC-UAB analyzed with EAG2000 software (Syntech, Hilversum, The Netherlands) on a personal computer.


Intact mosquito proboscises were hand-dissected and placed onto a coverglass by using Scotch double-stick tape (3M). Electrode gel (Spectra 360; Parker Laboratories, Fairfield, NJ) was applied to the amputated part of the proboscis. Glass recording and reference electrodes (1 mm i.d. with 5–7 MΩ resistance) were prepared (Model P-97; Sutter Instruments) and filled with insect saline (46). The reference electrode was inserted into the electrode gel, and the recording electrode was inserted into the lumen of a sensillum on the labellum until action potentials were achieved (47). The preparation was viewed at a 1,200× magnification by using a BX-40 microscope with a 50× LMPlanFl objective lens (Olympus, Melville, NY). Signals were imported via a 4-channel IDAC-USB and analyzed with AutoSpike software (Syntech) on a personal computer. Offline analysis of each individual 10-s recording was carried out by using AutoSpike software (47) such that small a and large b spike neurons were clearly distinguishable according to their spike amplitude. Olfactory responses were quantified by subtracting the number of action potentials 1 s before odor stimulation from the number of spikes 1 s after the onset of odor stimulation from individual preparations.

Anterograde Dye Filling.

Anterograde labeling using fluorescence dyes was performed as described (48), with slight modifications. The labellum was severed, and the remaining proboscis shaft was immediately immersed in a glass electrode filled with 1% neurobiotin (Vector Laboratories, Burlingame, CA) in PBS. Animals were kept in the humid chamber for 5–7 h, after which mosquito heads were fixed in 4% paraformaldehyde solution at 4°C overnight. Brains were dissected and washed in PBS for 2–4 h in the dark, followed by dehydration with a 0–100% ethanol series, followed by incubation in 100% propylene oxide (Sigma, St. Louis, MO) for 5 min before a descending ethanol-to-PBS rehydration procedure. The brains were then incubated with a 1:50 dilution of streptavidin–Alexa Fluor 546 conjugate (Molecular Probes, Carlsbad, CA) at 4°C overnight. DNA counterstaining of cells in the mosquito brain was carried out with a 1:1,000 dilution of TOTO3 (Molecular Probes) in PBST for 20 min. Whole mounts of the brain were washed in PBS for 1–2 h and mounted on a glass slide with Vectashield (Vector Laboratories). For plastic sectioning, brain preparations were dehydrated in a 25–100% ethanol series, followed by 100% acetone, and subsequently embedded in Spurr's epoxy resin (49) before 12-μm sections were prepared on a sliding microtome (HM340E; Microm, Waldorf, Germany). Whole mounts as well as plastic sectioned preparations of the stained mosquito brain were observed by using an LSM 510 confocal microscope (Zeiss, Thornwood, NY) under which optical sections were scanned at 0.5- to 2-μm intervals to capture fluorescence images from back-filling experiments.

In Situ Hybridization and Immunolabeling.

Paraffin-embedded preparations were sectioned at 10- to 12-μm thickness by using a sliding microtome (HM340E; Microm), subsequently dewaxed with Citri-Solv (Fisher BioSciences, Rockville, MD), and rehydrated in an ethanol series to PBS. In situ hybridization and probe preparation were carried out as described (6, 50), with digoxigenin-labeled RNA probes comprising ≈800 bp of AgOR6 coding sequence. Signals were visualized by alkaline phosphatase (AP) coupled to anti-DIG antibodies (Roche, Indianapolis, IN) at 1:1,000 dilution. AP signals were detected by using Fast Red tablets (Roche) according to the manufacturer's instructions. Anti-AgOR7 immunostaining was carried out as described (8). Images were captured with confocal microscopy as described above.

Supplementary Material

Supporting Information:


We thank Drs. J. Carlson and E. Hallem (Yale University, New Haven, CT), Dr. H. W Honegger, Mr. R. J. Pitts, and other colleagues in the Zwiebel laboratory for their comments on this manuscript; Drs. N. Strausfeld and W. Gronenberg for advice about graphics; Dr. R. F. Stocker for nc82 antiserum; Drs. B. Appel, C. Carter, T. Fitzwater, A. Goldman, H. C. Park, and H. Yan for help with in situ hybridizations; Drs. G. Pitts and R. Baugh for advice on single sensillum PCR; and Ms. P. Russell and Z. Li for mosquito rearing. This work was supported by National Institutes of Health Grants A1056402 and DC04692 (to L.J.Z.).


Anopheles gambiae odorant receptor
antennal lobe
odorant receptor
olfactory receptor neuron
subesophageal ganglion
single sensillum recording


Conflict of interest statement: No conflicts declared.

This paper was submitted directly (Track II) to the PNAS office.


1. Vosshall L. B., Amrein H., Morozov P. S., Rzhetsky A., Axel R. Cell. 1999;96:725–736. [PubMed]
2. Krieger J., Raming K., Dewer Y. M., Bette S., Conzelmann S., Breer H. Eur. J. Neurosci. 2002;16:619–628. [PubMed]
3. Fox A. N., Pitts R. J., Robertson H. M., Carlson J. R., Zwiebel L. J. Proc. Natl. Acad. Sci. USA. 2001;98:14693–14697. [PMC free article] [PubMed]
4. Clyne P. J., Warr C. G., Freeman M. R., Lessing D., Kim J., Carlson J. R. Neuron. 1999;22:327–338. [PubMed]
5. Hill C. A., Fox A. N., Pitts R. J., Kent L. B., Tan P. L., Chrystal M. A., Cravchik A., Collins F. H., Robertson H. M., Zwiebel L. J. Science. 2002;298:176–178. [PubMed]
6. Vosshall L. B., Wong A. M., Axel R. Cell. 2000;102:147–159. [PubMed]
7. Krieger J., Klink O., Mohl C., Raming K., Breer H. J. Comp. Physiol. A. 2003;189:519–526. [PubMed]
8. Pitts R. J., Fox A. N., Zwiebel L. J. Proc. Natl. Acad. Sci. USA. 2004;101:5058–5063. [PMC free article] [PubMed]
9. Melo A. C., Rutzler M., Pitts R. J., Zwiebel L. J. Chem. Senses. 2004;29:403–410. [PubMed]
10. Neuhaus E. M., Gisselmann G., Zhang W., Dooley R., Stortkuhl K., Hatt H. Nat. Neurosci. 2005;8:15–17. [PubMed]
11. Nakagawa T., Sakurai T., Nishioka T., Touhara K. Science. 2005;307:1638–1642. [PubMed]
12. Larsson M. C., Domingos A. I., Jones W. D., Chiappe M. E., Amrein H., Vosshall L. B. Neuron. 2004;43:703–714. [PubMed]
13. Elmore T., Ignell R., Carlson J. R., Smith D. P. J. Neurosci. 2003;23:9906–9912. [PubMed]
14. Meijerink J., Braks M. A., Van Loon J. J. J. Insect Physiol. 2001;47:455–464. [PubMed]
15. Cork A., Park K. C. Med. Vet. Entomol. 1996;10:269–726. [PubMed]
16. Stocker R. F. Cell Tissue Res. 1994;275:3–26. [PubMed]
17. Alcorta E. J. Neurophysiol. 1991;65:702–714. [PubMed]
18. Ellin R. I., Farrand R. L., Oberst F. W., Crouse C. L., Billups N. B., Koon W. S., Musselman N. P., Sidell F. R. J. Chromatogr. 1974;100:137–152. [PubMed]
19. Smallegange R. C., Qiu Y. T., van Loon J. J., Takken W. Chem. Senses. 2005;30:145–152. [PubMed]
20. Healy T. P., Copland M. J. Med. Vet. Entomol. 2000;14:195–200. [PubMed]
21. Dekker T., Steib B., Carde R. T., Geier M. Med. Vet. Entomol. 2002;16:91–98. [PubMed]
22. Hildebrand J. G., Shepherd G. M. Annu. Rev. Neurosci. 1997;20:595–631. [PubMed]
23. Wang Z., Singhvi A., Kong P., Scott K. Cell. 2004;117:981–991. [PubMed]
24. Thorne N., Chromey C., Bray S., Amrein H. Curr. Biol. 2004;14:1065–1079. [PubMed]
25. Shanbhag S. R., Park S. K., Pikielny C. W., Steinbrecht R. A. Cell Tissue Res. 2001;304:423–437. [PubMed]
26. Baugh L. R., Hill A. A., Brown E. L., Hunter C. P. Nucleic Acids Res. 2001;29:E29. [PMC free article] [PubMed]
27. Kelling F. J. PhD thesis. Germany: University of Groningen; 2001. pp. 95–104.
28. Warnes M. L., Finlayson L. H. Physiol. Entomol. 1986;11:469–473.
29. Meijerink J., Braks M. A., Braak A. A., Adam W., Dekker T., Posthumus M. A., Beek T. A., Van Loon J. J. A. J. Chem. Ecol. 2000;26:1367–1382.
30. Knols B. G. J., van Loon J. J. A., Cork A., Robinson R. D., Adam W., Meijerink J., de Jong R., Takken W. Bull. Entomol. Res. 1997;87:151–159.
31. Enserink M. Science. 2002;298:90–92. [PubMed]
32. Engstrom Y., Kadalayil L., Sun S. C., Samakovlis C., Hultmark D., Faye I. J. Mol. Biol. 1993;232:327–333. [PubMed]
33. Park K. C., Ochieng S. A., Zhu J., Baker T. C. Chem. Senses. 2002;27:343–352. [PubMed]
34. de Bruyne M., Foster K., Carlson J. R. Neuron. 2001;30:537–552. [PubMed]
35. Mechaber W. L., Capaldo C. T., Hildebrand J. G. J. Insect Sci. 2002 www.insectscience.org/2.5.
36. Schreck C. E., Kline D. L., Carlson D. A. J. Am. Mosquito Control Assoc. 1990;6:406–410. [PubMed]
37. Krieger J., Grosse-Wilde E., Gohl T., Dewer Y. M., Raming K., Breer H. Proc. Natl. Acad. Sci. USA. 2004;101:11845–11850. [PMC free article] [PubMed]
38. Krieger J., Klink O., Mohl C., Raming K., Breer H. J. Comp. Physiol. A. 2003;189:519–526. [PubMed]
39. Kent K. S., Harrow I. D., Quartararo P., Hildebrand J. G. Cell Tissue Res. 1986;245:237–245. [PubMed]
40. Thom C., Guerenstein P. G., Mechaber W. L., Hildebrand J. G. J. Chem. Ecol. 2004;30:1285–1288. [PubMed]
41. Guerenstein P. G., Yepez E. A., Van Haren J., Williams D. G., Hildebrand J. G. Naturwissenschaften. 2004;91:329–333. [PubMed]
42. Grant A. J., Wigton B. E., Aghajanian J. G., O’Connell R. J. J. Comp. Physiol. A. 1995:389–396. [PubMed]
43. Pitts R. J., Fox A. N., Zwiebel L. J. Proc. Natl. Acad. Sci. USA. 2004;101:5058–5063. [PMC free article] [PubMed]
44. Wetzel C. H., Oles M., Wellerdieck C., Kuczkowiak M., Gisselmann G., Hatt H. J. Neurosci. 1999;19:7426–7433. [PubMed]
45. Ayer R. K., Jr., Carlson J. J. Neurobiol. 1992;23:965–982. [PubMed]
46. O’Shea M., Adams M. E. Science. 1981;213:567–569. [PubMed]
47. de Bruyne M., Clyne P. J., Carlson J. R. J. Neurosci. 1999;19:4520–4532. [PubMed]
48. Ehmer B., Gronenberg W. Cell Tissue Res. 1997;290:153–165. [PubMed]
49. Spurr A. R. J. Ultrastruct. Res. 1969;26:31–43. [PubMed]
50. Goldman A. L., Van der Goes van Naters W., Lessing D., Warr C. G., Carlson J. R. Neuron. 2005;45:661–666. [PubMed]

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