Introduction

The sense of taste enables animals to find nutritious food while avoiding potentially harmful substances in their environment. Most animals have evolved sophisticated systems to detect and steer clear of consuming levels of substances that are toxic. Salts such as NaCl are vital for a wide array of physiological functions. However, consuming excessive salt can contribute to various health issues in mammals, including hypertension, osteoporosis, gastrointestinal cancer, and autoimmune diseases ^1–6. Therefore, high concentrations of salt are rejected by most animals.

Multiple studies have delved into how Na⁺ is sensed in the Drosophila taste system, shedding light on the mechanisms behind the attraction to low salt and aversion to high salt ^7–14. The major taste organs in flies, are two bilaterally symmetrical labella, each of which is decorated with 31 gustatory hairs (sensilla). These sensilla are characterized based on size (small, S; intermediate, I; large, L). The I-type sensilla harbor either two gustatory receptor neurons (GRNs), while the S- and L-sensilla contain four. These GRNs fall into five classes (A-E) based on their response profiles. These include A GRNs (formerly sugar GRNs), which respond to attractive compounds such as low salt, sugars, glycerol, fatty acids and carboxylic acids, B GRNs (formerly bitter GRNs), which are activated by high Na⁺, bitter compounds, acids, polyamines, tryptophan, and L-canavanine, ‘C’ GRNs respond to water, ‘D’ GRNs detect high levels of cations such as Na⁺, K⁺ and Ca²⁺, and ‘E’ GRNs sense low Na⁺ levels ¹⁵.

Several of the 66 member ionotropic receptor (IR) family function in the sensation of low and high salt. These include IR76b and Ir25a, which are IR-coreceptors, and therefore have broad roles in sensing many taste stimuli including low and high salt (sodium) ^8,12,16, calcium ¹⁷, several carboxylic acids ^18–20, fatty acids ^16,21-23, amino acids ²⁴, and carbonation ²⁵. A subset of A GRNs, as well as glutamatergic E GRNs are responsible for sensing low salt^8,11,12,15, and this sensation depends on IR56b working together with the broadly tuned IR25a and IR76b ⁸. Conversely, detection of high salt depends on B GRNs and D GRNs, and IR7c, in conjunction with IR25a and IR76b ⁷. Additionally, two Pickpocket channels, PPK11, PPK19, and Sano have been associated with high salt aversion^26,27. ppk19 and ppk11, which are members of the pickpocket (ppk) gene family, are expressed in taste-sensing terminal organs and play a role in appetitive and aversive behavior in response to low and high salt concentrations, respectively ²⁷.

In addition to the labellum and taste hairs on other external structures, fruit flies are endowed with an internal organ in the proboscis, called the pharynx, which functions in the decision to keep feeding or reject a food ^28–32. The pharynx includes three separate taste organs that line the esophagous: the labral sense organ (LSO), the ventral cibarial sense organ (VCSO), and the dorsal cibarial sense organ (DCSO) ^28,30,31. Each of these organs include hairless sensilla that house GRNs. A pair of GRNs in the LSO express a member of the gustatory receptor family, Gr2a, and knockdown of Gr2a in these GRNs impairs the avoidance to slightly aversive levels of Na^{+ 14}. Pharyngeal GRNs also promote the aversion to bitter tastants, Cu²⁺, L-canavanine, and bacterial lipopolysaccharides ^33–36. Other pharyngeal GRNs are stimulated by sugars and contribute to sugar consumption ^28,32,37. Remarkably, two pharyngeal GRNs function in the rejection rather the acceptance of sucrose ³⁸.

In this work, we investigated whether IRs function in the pharynx for avoidance of high Na^+. We found that IR60b, along with co-receptors IR25a and IR76b are required in the pharynx for preventing high salt consumption. IR60b is exclusively expressed in a pair of pharyngeal GRNs in the LSO, and these neurons are specifically activated by salt but not by any tested bitter compounds. When we optogenetically activated these IR60b-positive GRNs, proboscis extension responses were inhibited, indicating that these GRNs promote aversive behavior. Moreover, introduction of rat TRPV1 into the IR60b neurons induces aversive towards capsaicin, implying that these IR60b-positive GRNs are essential for instinctive avoidance. To validate the findings, we used a two-way choice DrosoX assay to measure actual ingestion levels. We found that the three Ir mutants consumed high salt at levels similar to sucrose over an extended period, emphasizing the critical role of this single pair of pharyngeal GRNs in controlling harmful ingestion of high salt.

Results and discussion

Ir60b functions in the repulsion to high salt

To identify potential salt sensors in Drosophila melanogaster, we conducted binary food choice assays using 30 Ir mutants (Figures 1A and S1A). Through screens in which we gave the flies a choice between 2 mM sucrose alone or 2 mM sucrose plus a low, attractive level of salt (50 mM NaCl), we confirmed that Ir76b ¹², Ir25a, and Ir56b ⁸, are essential for detecting low salt (Figure S1A). Moreover, consistent with Dweck et al. ⁸, using tip recordings to assay tastant-induced action potentials, we found that loss of Ir56b nearly eliminated spikes in response to low salt (Figure S1B). Using a Ir56b-GAL4 to drive UAS-mCD8::GFP, we also confirmed that the reporter was restricted to a subset of A GRNs, which were marked tdTomato (Figures S1D— S1F). We generated a UAS-Ir56b transgene which restored normal frequencies of action potentials in Ir56b-expressing GRNs (Figure S1B). Moreover, ectopic expression of UAS-Ir56b in GRNs that typically have minimal responses to low salt, caused a large increase in salt-induced action potentials (Figure S1C).

Figure 1.

In our behavioral screen for Ir mutants required for avoiding high salt (300 mM NaCl), we found that in addition to Ir7c, Ir25a, and Ir76b as previously described ⁷, Ir60b was also required (Figure 1A). The Ir60b mutant, Ir60b³, was generated by removing 768 base pairs, which spanned from 44 base pairs upstream of the predicted transcription start site to encompass the coding region for the initial 241 residues of the 577-amino acid protein (Figure S2A-C). Additionally, we verified the impairment in high salt avoidance using Ir60b¹, a gene previously investigated by Joseph et al. (Figure S2D) ³⁸. We conducted dose-response behavioral assays using Ir60b mutants, as well as Ir25a, Ir76b and Ir7c and found that all four mutants exhibited significant deficiencies in avoiding salt concentrations ranging from 200 mM to 500 mM (Figure 1B). Nevertheless, all of the mutants exhibited a strong aversion to extremely high salt concentrations, reaching 1000 mM, a level twice as concentrated as that found in the ocean. This extreme condition could potentially trigger the activation of additional pain or alarm neurons, serving as a protective mechanism to prevent potential tissue and organ damage.

Activation of Ir60b neurons inhibits motivation to feed

To investigate whether activation of Ir60b neurons induces aversive behavior, we used both chemogenetic and optogenetic approaches. Capsaicin, a ligand for the mammalian TRPV1 channel, does not normally elicit responses in flies (Figure 1C) ³⁹. Therefore, we expressed UAS-trpV1 under the control of the Ir60b-GAL4, and presented the flies with a choice between a 2 mM sucrose and a 2 mM sucrose containing 100 mM capsaicin. We found that the transgenic flies actively avoided capsaicin (Figure 1C), whereas expression of TRPV1 in A (sweet) GRNs (Gr64f-GAL4 and UAS-trpV1) induced a preference for capsaicin (Figure 1C). These findings support the idea that the activation of Ir60b neurons leads to gustatory avoidance.

To further test the proposal that Ir60b-positive GRNs elicit aversive behavior, we expressed CsChrimson, a light-activated cation channel⁴⁰ in Ir60b neurons. As controls we drove UAS-CsChrimson either in A GRNs (Gr5a-GAL4) or B GRNs (Gr66a-GAL4). Upon stimulation with red lights and sucrose, nearly all of the control flies (UAS-CsChrimson only) or flies expressing UAS-CsChrimson in A GRNs extended their proboscis (Figure 2B). In contrast, the PER was notably diminished in flies expressing UAS-CsChrimson in B GRNs (Gr66a-GAL4) or in Ir60b neurons (Gr66a-GAL4; 56.7±4.2% and Ir60b-GAL4; 55.0±5.0%, respectively; Figure 1D). These results provide compelling evidence supporting the notion that Ir60b-positive GRNs induce behavioral aversion.

Figure 2.

Ir60b is not required in the labellum to sense high salt

To investigate the physiological responses of labellar sensilla to high salt (300 mM), we conducted tip recordings on each of the 31 sensilla (Figure 2A). Five sensilla, including three S-type (S4, S6, and S8) and two L-type (L4 and L6), exhibited the strongest responses to high salt (Figure S3A). These responses were largely dependent on the broadly tuned IR25a and IR76b, as well as Ir7c (Figures 2B, 2C and S3B) as reported⁷. Interestingly, the Ir60b³ deletion mutant did not affect the neuronal responses to high salt in external sensory organs (Figures 2B and 2C). We inactivated individual GRNs by expressing the inwardly rectifying K⁺ channel (UAS-Kir2.1) ⁴¹ in A GRNs (Gr64f-GAL4) ⁴², B GRNs (Gr66a-GAL4) ⁴³, C GRNs (ppk28-GAL4) ⁴⁴, and D GRNs (ppk23-GAL4) ¹⁷, and confirmed that the aversive behavior and neuronal responses to high salt primarily relied on B and D GRNs (Figures 2D and 2E) as described ¹¹.

To examine the gustatory repulsion to high salt that is mediated through the labellum, we conducted proboscis extension response (PER) assays by stimulating the labellum. Starved control and Ir mutant flies extend their proboscis when the labellum is lightly touched by a 100 mM sucrose probe (Figure 2F). Upon a second sucrose offering, the various fly lines exhibited slightly and similarly diminished responses (Figure 2H). When we added 300 mM salt to the sucrose, it significantly reduced the PER in the control group (Figures 2G and 2I; first offering 40.9 ± 4.0%; second offering 41.5 ± 3.7%). Both the Ir25a² and Ir76b¹mutants also exhibited suppressed PERs, but the suppression was not as great as in the control (Figures 2G and 2I). In contrast, high salt reduced the PER by the Ir60b³ mutant to a similar extent as the control (Figures 2G and 2I; first offering 41.6 ± 6.5%; second offering 47.7 ± 6.7%). This indicates that the labellum of the Ir60b³ detects 300 mM salt normally, even though the mutant is impaired in avoiding high salt in a two-way choice assay (Figure 1A).

High salt sensor in the pharynx

The observations that Ir60b is required for the normal aversion to high salt, but does not appear to function in labellar hairs raises the possibility that Ir60b is required in the pharynx for salt repulsion. Ir60b is expressed in the pharynx where it plays a role in limiting sucrose consumption ³⁸. Gr2a is also expressed in the proboscis and contributes to the repulsion to moderate salt levels (150 mM) ¹⁴. However, the Gr2a^GAL4 mutant displays a normal response to high salt (450 mM) ¹⁴. In our two-way choice assay, which focuses on 300 mM NaCl, we found that salt repulsion displayed by the Gr2a^GAL4 mutant was also indistinguishable from the control (Figure S4).

To investigate a role for the pharynx in high salt (300 mM) repulsion, we conducted tests on Poxn mutants (Poxn^70–28/Poxn^ΔM22-B5), in which the external chemosensory sensilla have been converted to mechanosensory sensilla ⁴⁵. As a result, Poxn mutants possess only intact internal gustatory organs. We found that the aversive behavior to high salt was reduced in the Poxn mutants relative to the control (Figure 2J). However, the diminished avoidance was significantly different from Poxn^70–28/Poxn^ΔM22-B5;Ir60b³ mutants, even though Poxn^70–28/Poxn^ΔM22-B5;Ir60b³ mutants were not significantly different from Ir60b³ in avoidance to high salt (Figure 2J). Furthermore, the Poxn^70–28/Poxn^ΔM22-B5;Ir60b³ double mutant exhibited avoidance of high salt at a similar level to Ir60b³. This suggests that the internal sensor (IR60b-positive GRNs) can override the activation of the labellum in terms of aversive ingestion of high salt. Subsequently, we hypothesized that IR60b might not act as a primary gustatory sensor but rather as a regulator that allows for continued ingestion.

Quantification of reduced high salt ingestion in Ir60b mutants

To assess the ingestion of different food types, we employed the binary food choice assay, a qualitative method that utilizes blue, red, or purple dye colors in the abdomen ⁴⁶. However, for a more precise quantification of food ingestion, we recently developed the DrosoX system (Figure S5A) ⁴⁷. This system allowed us to directly measure the actual amount of food ingested over a period of 6 hours. In these experiments, we present flies with two capillaries: one containing 100 mM sucrose and the other containing 100 mM sucrose mixed with 300 mM NaCl. Control flies exhibited a preference for sucrose-only food, consuming it approximately four times more than the sucrose mixed with salt (Figure 3A). In contrast, the Ir25a, Ir60b, and Ir76b mutants displayed similar total ingestion levels (Figure 3A) and ingestion volume per hour (Figure 3B; ingestion index) for both tastants. In a prior study, it was observed that Ir60b mutant flies consumed high salt at a comparable rate to the control group when the total feeding time was recorded ³⁸. However, the DrosoX system now enables us to precisely quantify the ingestion volume. Additionally, we concurrently assessed two distinct tastants and compared their respective consumption levels. Consequently, our approach for evaluating avoidance behavior differs significantly.

Figure 3.

To further investigate the requirement for these genes, we performed genetic rescue experiments. We introduced their respective wild-type cDNAs under the control of their cognate GAL4 drivers, which resulted in a conversion from salt-insensitive behavior to the salt-sensitive behavior observed in wild-type flies (Figures 3C-3H). In addition, the defects in the Ir25a² and Ir76b¹ mutants were fully rescued by expressing the wild-type Ir25a and Ir76b transgenes, respectively, in the pharynx using the Ir60b-GAL4 (Figure 3I-3L). This suggests that both IR25a and IR76b act as coreceptors in the IR60b-expressing GRNs. Furthermore, we investigated whether the expression of UAS-Ir60b driven by Ir25a-GAL4 or Ir76b-GAL4 could rescue the defects observed in Ir60b³. Remarkably, despite the broad expression of IR60b using these GAL4 drivers, the Ir60b salt ingestion defect was eliminated (Figures 3M and 3N). Thus, it appears that simultaneous activation of GRNs that elicit attractive and aversive salt responses lead to repulsion. This suggests that activation of GRNs that induce salt aversion may suppress GRNs that function in salt attraction. If so, this would be reminiscent of bitter GRNs that suppress sugar GRNs through a GABAergic mechanism ⁴⁸.

Next, we addressed whether IR60b is specifically required for regulating the ingestion high salt. To investigate this, we assessed the consumption of caffeine, strychnine, and coumarin in Ir60b³ flies. We found that Ir60b³ displayed similar consumption patterns to the wild-type control flies for these bitter compounds (Figures 3O, 3P, S5B—S5E). This is in contrast to the impairments exhibited by the Gr66a^ex83mutant (Figures 3O, 3P, S5B—S5E), which is widely required for sensing many bitter chemicals. This indicates that IR60b is involved in regulating the avoidance of high salt ingestion rather than general avoidance responses to toxic compounds. Nevertheless, the role of IR60b in suppressing feeding is not limited to high salt, since IR60b also functions in the pharynx in inhibiting the consumption of sucrose ³⁸.

A single neuron in the LSO depends on Ir25a, IR60b and Ir76b for responding to both high salt and sucrose

In addition to IR60b, two other broadly required IRs (IR25a and IR76b) also function in repulsion to high salt. Moreover, we found that we could rescue the Ir25a, Ir60b or Ir76b DrosoX phenotypes using the same Ir60b-GAL4 to drive expression of the cognate wild-type transgenes in the corresponding mutant backgrounds. These finding imply that all three Irs are coexpressed in the pharynx. Therefore, we examined the relative expression patterns of the Ir60b-GAL4 reporter with Ir25a and Ir76b in the pharynx. We observed that the Ir76b-QF reporter was expressed in two cells within the labral sensory organ (LSO), one of which colocalized with Ir60b-GAL4 expression (Figure 4A). Additionally, the expression pattern of Ir25a-GAL4 perfectly overlapped with that of Ir76b-QF in the LSO (Figure 4B). Thus, we suggest that Ir25a, Ir60b and Ir76b function in the same GRN in the LSO to limit consumption of high salt. We attempted to induce salt activation in the I-type sensilla by ectopically expressing Ir60b, similar to what was observed with Ir56b ⁸; however, this did not generate a salt receptor (Figures S6A). Thus, the IR25a/IR60a/IR76b channel may require an additional subunit.

Figure 4.

To determine whether this GRN in the LSO is activated by high salt, we examined Ca²⁺ responses in the LSO using UAS-GCaMP6f, expressed under the control of each GAL4 driver. In the wild-type LSO, we identified a single cell that responded to 300 mM NaCl (Figure 4C). These data show that the single GRN in the LSO that expresses all three reporters responds to high salt. Moreover, this neuron responded robustly to 300 mM to 1000 mM Na⁺ but not to a low level of Na⁺ (50 mM; Figure 4E). We then examined the Ca²⁺ responses in the Ir25a², Ir60b³, and Ir76b¹ mutants, and found that each of them failed to respond to NaCl (Figures 4D and 4E). Additionally, we rescued the deficits in the GCaMP6f responses exhibited by each mutant by expressing a wild-type transgene under control of the corresponding GAL4 driver (Figure 4F). We also tested other Cl^- salts (CaCl₂, MgCl₂, and KCl) to determine if Cl^- rather an Na⁺ induced responses in the IR60b neuron. However, none of these other salts affected these neurons at 50 mM, 300 mM, and 500 mM concentrations tested (Figure 4G). In contrast, NaBr induced GCaMP6f responses (Figure 4H). Thus, the Ir60b neuron is responsive to Na⁺ and not Cl^-. Due to the effects of NaBr on the Ir60b neuron, we used the DrosoX assay to determine whether 300 mM NaBr suppressed ingestion of sucrose. We found that the impact of NaBr on sucrose ingestion was similar to that with NaCl (Figures S6B and S6C). We also found that bitter compounds such as quinine, caffeine, strychnine, lobeline, denatonium, and coumarin could not activate Ir60b neurons at the concentrations of 5 mM and 50 mM tested (Figure 4I).

It has been shown previously that Ir60b is required in single neuron in the pharynx for suppressing sucrose feeding, and this neuron responds to sucrose. ³⁸ Therefore, we tested whether the same neuron in the LSO that responds to salt also responds to sucrose. Using GCaMP6f, we found that the Ir60b neuron was activated by sucrose in the LSO of control flies, but not in the Ir25a, Ir60b and Ir76b mutants (Figure 4J). Thus, we conclude that the same LSO neuron depends on the presence of the same three receptors (IR25a, IR60b, and IR76b) for suppressing feeding in response to high salt or to sucrose. Mcdowell et al. demonstrated the presence of IR7c in labellar GRNs sensitive to high salt. They also unveiled the collaborative role of IR7c with IR25a and IR76b in perceiving and responding to high salt concentrations. Consequently, we investigated whether IR7c played a potential role in Ir60b-positive pharyngeal GRNs. However, our experiments did not reveal any physiological defects in Ir7c mutant flies (Figure S7A). Furthermore, our findings indicated that Ir7c is not expressed within the Ir60b-positive GRNs (Figure S7B—D).

Although prior research had identified the involvement of IR7c, IR25a, and IR76b in the labellar GRNs for high salt detection, our study introduces a perspective by highlighting the combination of IR25a, IR60b, and IR76b as internal molecular sensors responsible for detecting and ingesting high salt.

Materials and methods

Key resources table

Generation of Ir60b³ and UAS-Ir60b lines

The Ir60b³ mutant were generated by ends-out homologous recombination ⁵¹. For generating the construct to injections, approximately two 3-kb genomic fragments were amplified by PCR, and subcloned the DNAs into the pw35 vector of NotI and BamHI sites. Assuming the ‘‘A’’ of the ‘‘ATG’’ starts codon as “+1”, the deleted region was −44 to +724. The construct was injected into w¹¹¹⁸ embryos by Best Gene Inc. We outcrossed the mutant with w¹¹¹⁸ for 6 generations.

To generate the UAS-Ir60b transgenic strain, we employed mRNA to perform reverse transcription polymerase chain reaction (RT-PCR) on the full-length Ir60b cDNA, which was subsequently subcloned into the pUAST vector. The insertion took place between the EcoRI and NotI sites designated for UAS-Ir60b. The primer set used for amplification is as follows: 5’-GAGAATTCAACTCGAAAATGAGGCGG-3’ and 5’-ATGCGGCCGCAATGCTAATTTTG-3’. The integrity of the cloned cDNA was verified through DNA sequencing. Subsequently, the transformation vector harboring the respective constructs was introduced into w¹¹¹⁸ embryos via injection (KDRC).

Binary food choice assay

We conducted binary food choice assays following the methods outlined in a previous study⁴⁶. Initially, a group of 40–50 flies (3-6 days old) were subjected to an 18-hour starvation period on a 1% agarose substrate. Two mixtures were prepared, each containing a specific dye, and they were distributed in a zigzag pattern. The first mixture consisted of 1% agarose, the indicated concentration of saponin, and 5 mM sucrose with red dye (sulforhodamine B, 0.1 mg/ml). The second mixture contained 1% agarose, 1 mM sucrose, and blue dye (Brilliant Blue FCF, 0.125 mg/ml). The prepared 72-well microtiter dish was then used to transfer the flies, which was placed in a dark and humid chamber. After feeding, the flies were sacrificed by freezing them at −20°C. Subsequently, their abdomen colors were examined under a microscope to identify the presence of red, blue, or purple dye, allowing us to segregate them accordingly. The counts were taken for the number of flies with blue (N_B), red (N_R), and purple (N_P) abdomens. The preference index (P.I) was calculated using the following equation: (N_B - N_R)/(N_R + N_B + N_P) or (N_R - N_B)/(N_R + N_B + N_P), depending on the specific dye/tastant combinations. A P.I. of −1.0 or 1.0 indicated a complete preference for either 5 mM sucrose with saponin or 1 mM sucrose alone, respectively. A P.I. of 0.0 indicated no preference between the two food alternatives.

Chemical reagent

Sucrose (CAS No. 57-50-1), tricholine citrate (TCC) (CAS No. 546-63-4), sulforhodamine B (CAS No. 3520-42-1), capsaicin (CAS No. 404-86-4), caffeine (CAS No. 58-08-2), CaCl₂ dihydrate (CAS No. 10035-04-8), KCl (CAS No. 7447-40-7), quinine (CAS No. 6119-47-7), strychnine (CAS No. 1421-86-9), lobeline (CAS No. 134-63-4), denatonium (CAS No. 6234-33-6), and coumarin (CAS No. 91-64-5) were purchased from Sigma-Aldrich (USA). Brilliant blue FCF (CAS No. 3844-45-9) was purchased from Wako Pure Chemical Industry (Japan). Paraformaldehyde (CAS No. 30525-89-4) was purchased from Electron Microscopy Sciences (USA). NaCl (CAS No. 7647-14-5) was purchased from LPS solution (Korea). NaBr (CAS No. 7647-15-6) was purchased from DUKSAN (Korea). Goat serum, New Zealand Origin was purchased from Gibco (USA).

Droso-X assay

We conducted Droso-X assays following the methods outlined in a previous study⁴⁷. The amount of ingestion was measured using a Droso-X system (Scitech Korea, Korea) located in a controlled incubator (25°C, 60% humidity). To quantify the ingestion, a mixture comprising 100 mM sucrose and the specified concentration of chemicals was injected into a glass tube (Cat. No. 53432-706; VWR International, USA) using a syringe (KOVAX-SYRINGE 1 ml 26G; KOREA VACCINE, Korea) and needle (Cat No. 90025; Hamilton, Switzerland). Each cuvette contained flies (3-6 days old) and was physically isolated to prevent them from consuming the solution prior to the experiment. The experiment was conducted for a duration of 6 hours, specifically from 9 am to 3 pm. The DROSO X&XD software (Scitech Korea, Korea) was utilized by the Droso-X system to record the amount of solution consumed. The ingestion amount at time X (X h) was calculated as the difference between the initial solution amount (0 h) and the solution amount at time X.

The ingestion index (I.I) was calculated in each time point using the following equation: (Ingestion volume_DrosoX - Ingestion volume_DrosoXD)/(Ingestion volume_DrosoX + Ingestion volume_DrosoXD) or (Ingestion volume_DrosoXD – Ingestion volume_DrosoX)/(Ingestion volume_DrosoXD + Ingestion volume_DrosoX), depending on the specific tastant combinations. A I.I. of 0.0 indicated no preference based on their ingestion between the two food alternatives.

Electrophysiology

Electrophysiology, specifically the tip recording assay, was conducted following the previously described method⁵². The tip recordings were carried out based on Tanimura’s nomenclature. The average frequencies of action potentials (spikes/s) evoked in response are presented, with only spikes occurring between 50 and 550 ms included in the count. For the tip recordings, we followed the established protocol using the specified concentration of saponin dissolved in distilled water with 30 mM tricholine citrate (TCC) for the assay. These electrolytes, 1 mM KCl or 30 mM TCC, served as the recording medium. To begin the recordings, we immobilized flies (3-6 days old) by exposing them to ice. A reference glass electrode filled with Ringer’s solution was inserted through the back thorax and passed into the proboscis. The sensilla on the labial palp were stimulated with a compound dissolved in the buffer solution of the recording pipette, which had a tip diameter of 10-20 μm. The recording electrode was connected to a pre-amplifier (Taste PROBE, Syntech, Germany), which amplified the signals by a factor of 10 using a signal connection interface box (Syntech) and a 100-3,000 Hz band-pass filter. The recorded action potentials were acquired at a sampling rate of 12 kHz and analyzed using Autospike 3.1 software (Syntech). Subsequently, recordings were performed on the indicated sensilla on the labial palp.

Proboscis extension response assay

The proboscis extension response experiment was conducted with some modifications previously described by a previous study⁵³. A group of 20-25 flies (3-6 days old) was deprived of food for 18-20 hours in vials containing wet Kimwipe paper with tap water. After briefly anesthetizing the flies on ice, they were carefully trapped inside a pipette tip with a volume of 20-200 µl. To expose their heads, the edge of the pipette tip was gently cut using a paper cutter blade. The protruded head and proboscis were used to deliver stimuli during the experiment. To stimulate the flies’ tarsi, the head/proboscis and forelegs were extended outside the pipette tip without causing any harm. To eliminate any potential biases due to thirst, water was initially provided to the flies. For both the positive control and initial stimulation, a 2% sucrose solution was used. The tastant stimuli, consisting of either 2% sucrose or 300 mM NaCl, were presented using Kimwipe paper as the medium. To conduct these experiments, we selected flies that responded to sucrose. Flies that did not exhibit a reaction to the sucrose during the initial exposure were excluded from the experiment. The same conditions as the initial exposures were maintained for the second exposure. Each test round involved the use of more than 10 flies.

Immunohistochemistry

We performed immunohistochemistry as previously described⁵⁴ with slight modifications. The labella of flies (6-8 days old) were dissected and fixed in a solution containing 4% paraformaldehyde (Electron Microscopy Sciences, Cat No 15710) and 0.2% Triton X-100 for 15 minutes at room temperature. After that, the tissues were washed three times with PBST (1x PBS and 0.2% Triton X-100) and then bisected using a razor blade. Subsequently, the tissues were incubated in blocking buffer (0.5% goat serum in 1x PBST) for 30 minutes at room temperature. To detect the target protein, primary antibodies (mouse anti-GFP; Molecular Probes, Cat No A11120; diluted 1:1,000) were added to fresh blocking buffer and left to incubate with the labellum samples overnight at 4 °C. Following this, the tissues were washed three times with PBST and incubated with the secondary antibody (goat anti-mouse Alexa Fluor 488, diluted 1:200) for 4 hours at 4 °C. Afterwards, the tissues were washed three times with PBST and placed in 1.25x PDA mounting buffer (containing 37.5% glycerol, 187.5 mM NaCl, and 62.5 mM Tris pH 8.8). Finally, the samples were visualized using a Leica Stellaris 5 confocal microscope.

Ex vivo calcium imaging

Ex vivo Ca²⁺ imaging was performed as previously described⁵⁵ with slight modifications. Ex-vivo calcium imaging was conducted using a low melting agarose method. For the experimental process, flies (6-8 days old) expressing UAS-GCaMP6f driven by Ir25a-GAL4, Ir60b-GAL4, and Ir76b-GAL4 were used (incubation conditions: humidity: 50-60%, temperature: 25°C, Light/Dark: 12/12 hours). A 0.5% low melting agarose solution was prepared and applied to a confocal dish (Cat No. 102350, SPL LIFE SCIENCE, Korea). A mild swallow deep well was prepared for sample fixation. Subsequently, the heads of the flies were carefully decapitated using sharp razor blades, followed by excising a small portion of the labellum in the extended proboscis region to facilitate tastant access to pharyngeal organs. The prepared tissue sample was then carefully fixed in an inverted position in the pre-prepared well. Videos were recorded with Axio Observer 3 (Carl Zeiss) Adult hemolymph (AHL) composites 108 mM NaCl, 5 mM KCl, 8.2 mM MgCl₂, 2 mM CaCl₂, 4 mM NaHCO₃, 1 mM NaH₂PO₄, and 5 mM HEPES pH 7.5. A pre-stimulus solution, AHL was used, followed by the stimulus solution after 60 seconds, enabling direct access of the stimulant with AHL to the pharyngeal neurons. GCaMP6f fluorescence was observed using a fluorescence microscope with a 20x objective, specifically focusing on the relevant area of the pharynx. Videos were recorded at a speed of two frames per second. Neuronal fluorescent activity changes were recorded for 5 minutes following stimulus application. Fiji/ImageJ software (https://fiji.sc) was used to measure fluorescence intensities. A region of interest (ROI) was drawn around the cell bodies, and the Time-Series Analyzer Plugin, developed by Balaji, J. (https://imagej.nih.gov/ij/plugins/time-series.html), was utilized to measure the average intensity for ROIs during each frame. The average pre-stimulation value before chemical stimulation was calculated. ΔF/F (%) was determined using the formula (F_max-F₀)/F₀ × 100%, where F0 represents the baseline value of GCaMP6f averaged for 10 frames immediately before stimulus application, and Fmax is the maximum fluorescence value observed after stimulus delivery.

Statistical analysis

The error bars on the graph indicate the standard error of the means (SEMs), while the dots represent the number of trials conducted for the experiment. To compare multiple datasets, we utilized single-factor ANOVA coupled with Scheffe’s analysis as a post hoc test. Pairwise comparisons were conducted using Student’s t-test. Statistical significance is denoted by asterisks (*p < 0.05, **p < 0.01). We performed all statistical analyses using Origin Pro 8 for Windows (ver. 8.0932; Origin Lab Corporation, USA).

Acknowledgements

This work was supported by grants to Y.L. from the National Research Foundation of Korea (NRF) funded by the Korea government (MIST) (NRF-2021R1A2C1007628) and Biomaterials Specialized Graduate Program through the Korea Environmental Industry and Technology Institute (KEITI) funded by the Ministry of Environment (MOE) and grants to C.M. from the National Institute on Deafness and other Communication Disorders (NIDCD), R01-DC007864 and R01-DC016278. S.D., B.S., and D.N were supported by the Global Scholarship Program for Foreign Graduate Students at Kookmin University in Korea.

Additional information

Funding

Author contributions

J.S. and S.D. performed most of the genetic studies and physiology. J.S. and Y.L. conceived and designed the experiments. B.S. confirmed optogenetics and ectopic experiments. D.N. worked on low salt sensors. Y.K. generated most new reagents and conducted screening for high salt sensor in the initial stage. A.G. worked on the initial calcium imaging. C.M. and Y.L. supervised the project and wrote the manuscript.

Additional files

Supplementary files

• Supplementary Figures S1-S5

Data availability

Source data for all figures contained in the manuscript and SI have been deposited in ’fiigshare’ (https://doi.org/10.6084/m9.figshare.23939394).

Supplementary files

• Supplemental figure 1 code for ‘Requirements for three Irs for preferred low salt-containing food’ presented in Figure 1.

• Supplemental figure 2 code for ‘Gene structure of Ir60b locus, generation of Ir60b³ and behavioral defect of Ir60b¹ in high salt avoidance in Figure 1.

• Supplemental figure 3 code for ‘Electrophysiological responses with high salt’ presented in Figure 2.

• Supplemental figure 4 code for ‘The preference of Gr2a mutant with high salt’ presented in Figure 2.

• Supplemental figure 5 code for ‘DrosoX system and measurement of food intake using strychnine and coumarin’ presented in Figure 3.

• Supplemental figure 6 code for ‘Measurement of food intake of 300 mM NaBr and ectopic experiment of IR60b’ presented in Figure 4.

• Supplemental figure 7 code for ‘Quantification of calcium response in Ir7c mutants and control groups within the Ir60b-positive GRNs, alongside expression profiling of Ir7c and Ir60b’ presented in Figure 4.