Abstract
Fat represents a calorically potent food source that yields approximately twice the amount of energy as carbohydrates or proteins per unit of mass. The highly palatable taste of FAs promotes food consumption, activates reward circuitry, and is thought to contribute to hedonic feeding underlying many metabolism-related disorders. Despite a role in the etiology of metabolic diseases, little is known about how dietary fats are detected by the gustatory system to promote feeding. Previously, we showed that a broad population of sugar-sensitive taste neurons expressing Gustatory Receptor 64f (Gr64f), is required for reflexive feeding responses to both FAs and sugars. Here, we report a genetic silencing screen to identify specific populations of taste neurons that mediate fatty acid taste. We find neurons identified by expression of Ionotropic Receptor 56d (IR56d) are necessary and sufficient for reflexive feeding response to FAs. Functional imaging reveals that IR56d-expressing neurons are responsive to short- and medium-chain FAs. Silencing IR56d neurons selectively abolishes fatty acid taste, while their activation is sufficient to drive feeding responses. Analysis of co-expression with Gr64f identifies two subpopulations of IR56d-expressing neurons. While physiological imaging reveals that both populations are responsive to FAs, IR56d/Gr64f neurons are activated by medium-chain FAs and are sufficient for reflexive feeding responses to FA. Moreover, flies can discriminate between sugar and FAs in an aversive taste memory assay, indicating that FA taste is a unique modality in Drosophila. Taken together, these findings localize fatty acid taste within the Drosophila gustatory center and provide an opportunity to investigate discrimination between different categories of appetitive tastants.
Authors Summary Fat represents a calorically potent food source that yields approximately twice the amount of energy as carbohydrates or proteins per unit of mass. Dietary lipids are comprised of both triacylglycerides and FAs, and growing evidence suggests that it is the free FAs that are detected by the gustatory system. The highly palatable taste of FAs promotes food consumption, activates reward centers in mammals, and is thought to contribute to hedonic feeding that underlies many metabolism-related disorders. Despite a role in the etiology of metabolic diseases, little is known about how dietary fats are detected by the gustatory system to promote feeding. We have identified a subset of sugar-sensing neurons in the fly that also respond to medium-chain FAs and are necessary and sufficient for behavioral responses to FAs. Further, we find that despite being sensed by shared neuron populations, flies can differentiate between the taste of sugar and FAs, fortifying the notion that FAs and sugar represent distinct taste modalities in flies.
Introduction
Fat represents a calorically potent food source that yields approximately twice the amount of energy as carbohydrates or proteins per unit of mass. In mammals, dietary lipids are detected by taste cells, mechanosensory and olfactory neurons, as well as by post-ingestive feedback [1–4]. Dietary lipids are comprised of triacylglycerides and FAs (FAs), and growing evidence suggests that it is the free FAs that are detected by the gustatory system [5–7]. Sensing of FAs promotes food consumption, activates reward circuitry, and is thought to contribute to hedonic feeding that underlies many metabolism-related disorders [8,9]. Despite a role in the etiology of metabolic diseases, little is known about how dietary fats are detected by the gustatory system to promote feeding.
In flies and mammals, tastants are sensed by dedicated gustatory receptors that localize to the taste cells or taste receptor neurons [10–12]. These cells are sensitive to different taste modalities such as sweet, bitter, salty, sour or umami, and project to higher order brain structures for processing [10,13,14]. While these taste modalities have been extensively studied, much less is known about how FAs are detected and how information about this sensory stimulus is processed. Taste neurons in Drosophila are housed in gustatory sensilla located on the tarsi (feet), proboscis (mouth) and wings. Each sensillum contains dendrites of up to four gustatory receptor neurons (GRNs), which are activated by different categories of tastants [16]. Two main classes of neurons include one group that senses sweet tastants and promotes feeding, and a second, non-overlapping group that senses bitter tastants and promotes food avoidance [17,18]. We have previously shown that Drosophila is attracted to medium-chain FAs [19]. Consumption of FAs relies on taste, rather than smell, as it is not impaired by surgical ablation of olfactory organs [19]. Additionally, FA consumption is abolished in pox-neuro mutants in which all external taste hairs are converted to mechanosensory bristles, indicating that the chemical signature rather than oily texture of FAs is associated with perception [19]. Silencing Gr64f neurons, which are required for sugar sensing [20,21], abolishes behavioral response to FAs, suggesting that shared populations of gustatory neurons detect FAs and sugars [19]. Unlike sugars, FAs sensing is dependent on functional Phospholipase C (PLC) indicating independent sensory pathways for FA and sugar perceptions [18]. However, any further characteristics of the physiological response or the specific neuronal identity of the neurons mediating FA response are unknown.
Taste neurons from the legs and proboscis predominantly project to the subeosophageal zone (SEZ), the primary taste center of the Drosophila central nervous system, but the downstream central brain circuitry contributing to taste processing is less well understood [22–25]. Determining how diverse tastants activate GRNs that convey information to the SEZ, and how this information is represented in higher order brain centers, is central to understanding the neural basis for taste processing and feeding behavior. Identifying the neural principles underlying FA taste processing requires identifying the precise populations of taste neurons responsive to FAs, and their innervation of the primary taste center. Recent studies in Drosophila have identified taste neurons that are responsive to diverse modalities including salt, sugar, amino acids, water, carbonation, bitter, polyamines, and electrophilic tastants [17,26–32], yet little is known about the populations underlying FA taste. Here, we show that the IR56d population of gustatory neurons, which partially overlaps with Gr64f neurons, is necessary and sufficient to produce a feeding response to FAs. The IR56d-Gr64f population of neurons is selectively responsive to medium-short-chain FAs. Our results reveal a defined channel that senses FAs to promote food consumption, providing a mechanism for differentiation between attractive tastants of different modalities.
Results
Hexanoic acid elicits calcium activity in Gr64f sensory neurons
We previously reported that silencing the Gr64f population of primary taste neurons abolishes behavioral response to both sugars and FAs [10]. To investigate the responsiveness of these neurons to FAs directly, the Ca2+ sensor GCaMP5 was expressed under control of Gr64f-GAL4 [33,34] (Fig. 1A). The Ca2+ response to proboscis application of water, sucrose, or the medium-chain fatty acid hexanoic acid (HxA) was monitored in vivo in the axonal projections of Gr64f neurons within the SEZ (Fig. 1B). Flies were provided either 10mM sucrose or 1% HxA, as these concentrations induce comparable levels of proboscis extension reflex behavior. Both 10mM sucrose and 1% HxA induced robust activity in the SEZ, while little response was observed to water alone (Fig. 1C-F). The temporal dynamics of Ca2+ activity differed between the two tastants, with HxA eliciting a broader response (Fig. 1E, F), yet both elicited comparable peak changes in fluorescence (Fig 1C). Therefore, both sugars and FAs activate Gr64f sweet-sensing GRNs, fortifying the notion that this neuronal class is generally responsive to attractive tastants.
To localize FA-sensitive neurons within the broad Gr64f population, we selectively silenced populations of taste neurons predicted to overlap with Gr64f neurons and examined the proboscis extension reflex (PER) in response to sucrose and HxA. The synaptobrevin cleavage peptide Tetanus Toxin-Light Chain (TNT) was expressed under the control of Gustatory Receptor or Ionotropic Receptor promoters known to overlap with Gr64f (Table S1) [20,30,35,36]. Of the 10 GAL4 lines tested, silencing with GAL4 drivers for Gr61a, IR56b, and Gr64f resulted in PER defects to sucrose and HxA, By contrast, silencing Gr64e neurons significantly reduced response to sucrose without affecting response to HxA, and silencing Gr5a, Gr43a, or IR56d neurons significantly reduced PER to HxA without affecting sucrose response.
We chose to further investigate the role of IR56d neurons in FA sensing, since IRs have been found to be involved in detection of non-sweet appetitive tastants, including salt and amino acids [28,37,38]. IR56d neurons have previously been reported to project to two distinct regions in the SEZ. One region overlaps with sweet-sensing projections, and the other is indicated to originate in the taste pegs of the proboscis [36]. Outside of sensing carbonation, little is known about ligands that activate the taste pegs or the role they play in the Drosophila gustatory system. The sweet-sensing projections of IR56d resembled the region of Gr64f projections that was activated by HxA. Consistent with previous reports, expression of GFP in IR56d neurons (IR56d-GAL4>cd8::GFP) revealed two populations of projections: one innervating the anterior SEZ and previously defined as emanating from labellar bristles, and a second population, emanating from the taste pegs, innervating the posterior SEZ (Fig 2A-C) [36]. To further analyze the role of IR56d neurons, we silenced them using TNT and tested PER upon application of tastants to the proboscis for consistency with live imaging experiments. To control for genetic background and any potential non-specific effects of TNT, we compared PER in flies with silenced IR56d neurons (IR56d-GAL4>UAS-TNT) to flies expressing an inactive variant of TNT (IR56d-GAL4>imp-TNT) [35]. Expression of imp-TNT in IR56d neurons did not affect PER in response to sucrose or HxA, while expression of TNT selectively inhibited HxA response (Fig 2D). Therefore, IR56d neurons are required for behavioral response to HxA, but dispensable for response to sucrose.
Broad activation of sweet-sensing neurons expressing the trehalose receptor Gr5a induces feeding response in the absence of tastants [17,39,40]. To determine whether activation of IR56d neurons is sufficient to induce PER, we selectively expressed the thermo-sensitive cation channel transient receptor potential A1 (TRPA1) in IR56d neurons, or Gr5a neurons as a positive control, and measured heat-induced PER [41,42]. TRPA1 expression induces neuronal activity at temperatures above 28°C, but has little effect on neuronal activity in flies at 22°C, allowing for thermogenetic modulation of neuronal activity [41,42]. In agreement with previous findings, broad activation of sweet-sensing neurons induced robust PER (Fig 2F) [27,43,44]. Similarly, selective activation of IR56d neurons robustly induced PER as compared to control flies harboring UAS-TRPA1 or IR56d-GAL4 alone (Fig 2F). Therefore, activation of IR56d neurons alone is sufficient to induce PER.
IR56d neurons project to both taste peg and sweet-sensing regions of the SEZ, and each region can be distinguished anatomically (Fig 2A-C; [36]). To determine whether sugars and FAs can differentially activate the two regions, we expressed GCaMP5 in IR56d neurons (IR56d-GAL4>GCaMP5) and measured tastant-evoked activity separately within the areas of anterior and posterior projections. The anterior region, which overlaps with Gr64f neurons, responded to both HxA and sucrose with similar magnitude (Fig 3A-B). The projections from the taste pegs, however, showed a strong response to HxA, but did not respond to sucrose (Fig 3C-D). Therefore, while IR56d neurons that project posteriorly are responsive to both sucrose and HxA, those originating in taste pegs are selectively responsive to HxA, revealing two distinct functional classes of IR56d taste neurons.
Flies exhibit feeding responses upon presentation of several FA classes [19]. It is possible that the IR56d neurons are broadly responsive to different classes of FAs. Alternatively, different classes of FAs may be sensed by distinct, or partially overlapping, populations of sensory neurons. To test the responsiveness of IR56d neurons to different classes of FAs, we first compared behavioral responses of IR56d-silenced flies to short-chain pentanoic acid (5-carbon), medium-chain octanoic acid (8 carbon), and long chain oleic acid (18-carbon). As compared to control IR56d>imp-TNT flies, IR56d-silenced flies exhibited weaker responses to octanoic acid, but retained robust PER to pentanoic acid (Fig 4A), suggesting that PER to short chain FAs is not dependent on IR56d neurons. Oleic acid did not elicit strong PER in either. We directly examined activity induced by different FAs in IR56d neurons with in vivo Ca2+ imaging in IR56d-GAL4>GCaMP5 flies. Octanoic acid robustly activated posterior IR56d projections, while both pentanoic and octanoic acid activated anterior IR56d projections (Fig 4B,C). Oleic acid, which did not induce PER, also did not activate IR56d projections in either posterior or anterior regions (Fig 4B,C). Together, these findings show that IR56d neurons respond to short and medium chain FAs, and further, that sub-populations of IR56d neurons that are separately represented in anterior and posterior locations in the SEZ have distinct FA response profiles.
Since silencing of either IR56d or Gr64f neurons abolishes PER to hexanoic and octanoic acid, one possibility is that neurons co-expressing both IR56d and Gr64f are required for medium-chain FA response (Fig 5A). To validate co-expression of IR56d and Gr64f, we used the LexA system to label Gr64f neurons (Gr64f-LexA>LexOp-CD8:GFP) and the GAL4 system to label IR56d neurons (IR56d-GAL4>UAS-RFP) (Fig 5A). Examining SEZ projections revealed co-localization within the posterior SEZ, with no co-localization detected in the anterior SEZ, suggesting the posterior IR56d neurons co-express Gr64f and IR56d. To determine whether the IR56d+/Gr64f+ co-expressing neurons are required for FA taste, we used intersectional strategies to selectively silenced anterior IR56d neurons of the taste pegs. Specifically, we repressed TNT expression in IR56d+/Gr64f+ using Gr64f-LexA to drive expression of the GAL80 repressor (IR56d-GAL4>UAS-TNT; Gr64f-LexA>LexAop-GAL80) (Fig 5B). Selectively silencing IR56d neurons that do not overlap with Gr64f did not affect PER to HxA, or as expected, to sugar, and no difference was observed between flies expressing imp-TNT and TNT, suggesting the taste peg neurons are dispensable for the reflexive feeding response to FAs (Fig 5B). Flies lacking Gr64f-LexA, but still harboring a copy of LexAop-GAL80 (IR56D-GAL4-TNT; LexAop-GAL80/+), showed reduced PER as expected (Fig S1).
Although both sugars and FAs induce feeding behavior, it is unclear whether flies can qualitatively differentiate between these two classes of appetitive tastants. We have developed an assay in which an appetitive tastant is applied to the tarsi, paired with bitter quinine application to the proboscis, and the suppression of PER in subsequent response to the appetitive tastant is measured [40,45]. To determine whether flies can differentiate between sugars and FAs, we applied sucrose (conditioned stimulus) to the tarsi followed immediately with quinine application (unconditioned stimulus) to the proboscis. Following three training trials, memory was tested by application of sucrose (control) or HxA to the tarsi, in the absence of quinine, and measuring PER (Fig 6B). We also performed reciprocal experiments in which flies were trained with HxA and tested for PER to sucrose or HxA.
As previously reported, pairing sugar with quinine significantly reduced PER over the course of three training trials and this memory persisted in the absence of quinine ([46,47]; Fig 6B). Further, there were no differences in PER to HxA between flies repeatedly provided sucrose in the absence of quinine and flies trained with sucrose-quinine pairing, indicating that the aversive taste memory formed to sucrose is not generalized to HxA. Conversely, the pairing of HxA and quinine resulted in a robust PER reduction that was not generalized to sucrose (Fig. 6C). Quantification of the percentage reduction in PER reveals that the ‘matched’ groups, where quinine is paired with an attractive tastant suppressed PER by 79-94%, while there was little PER suppression when the tested tastant was unmatched from the one previously paired with quinine (Fig 6D). This reciprocal discrimination between sucrose and HxA is different from the unilateral discrimination between two sugars reported previously [45] and indicates that flies can discriminate between sucrose and HxA based on their identity. Thus, sugars and FAs act as independent taste modalities in flies.
Discussion
Here, we identify a small population of neurons within the taste system that is responsive to FAs. Other taste modalities, such as sweet and bitter, have been well studied in flies and mammals, but much less is known about FA taste. We previously identified a requirement for Gr64f-GAL4 neurons in the taste of FA [19]. The findings presented here further localize the reflexive feeding response to taste of hexanoic and octanoic acid, both medium-chain FAs, to a subpopulation of IR56d neurons. ‘Sweet-sensing’ neurons in Drosophila have been broadly classified as ones responding to sugars and other sweet tastants such as glycerol, subsets of which may also be involved in sensing other appetitive tastants such as amino acids [17,28,48]. Our findings indicate that IR56d neurons represent channels for FA detection, and likely allow flies to discriminate between different attractive tastants.
Localization of fatty acid-sensitive neurons
We identify a subpopulation of sweet sensing neurons that are responsive to FAs. We have previously shown that genetic silencing or ablation of most sweet-sensing neurons using Gr64f-GAL4 abolished FA taste, suggesting shared neurons detect sugar and FA taste [19]. In flies, subpopulations of Gr64f neurons are selectively sensitive to different tastants including a Gr64e population that is sensitive to glycerol [48] and Gr5a subset that is sensitive to trehalose [17]. To localize the Gr64f neurons responsible for FA taste we conducted a targeted screen and silenced neurons that are believed to overlap with Gr64f neurons, which led us to study the IR56d population of neurons. ISilencing Ir56d neurons appears to selectively disrupt HxA taste without affecting response to sucrose, supporting the notion that independent channels within the Gr64f population mediate response to sugar and FAs. It is possible that FA-sensitive neurons are broadly tuned to FAs, or selectively respond to distinct classes of FAs. Our Ca2+ imaging experiments indicate that IR56d neurons are responsive to medium-chain HxA (six carbons (C6), saturated) and octanoic acid (C8, saturated) in both anterior and posterior regions, and to short-chain pentanoic acid (C5, saturated), but only in the anterior projections. We do not find IR56d neurons responsive to long chain, unsaturated oleic acid (C18, mono-unsaturated). These findings are supported by behavioral data revealing that flies exhibit robust PER in response to pentanoic acid, HxA, and octanoic acid, but not oleic acid. Therefore, it seems likely that flies are selectively responsive to short/medium chain FAs, but less responsive to long-chain and/or unsaturated FAs. It is possible that a population of taste neurons independent from IR56d are activated by pentanoic acid because it elicited a robust behavioral response that was independent of Ir56d neurons. Further, IR56d neurons may be activated by long-chain FAs that were not tested, and these could modulate feeding response and induce PER. Nevertheless, the findings presented here reveal specificity for short/medium chain FAs within a defined population of taste neurons, and further specificity for medium chain FAs in a subset of those neurons.
Discrimination between attractive substances
The robust PER response induced by two different medium-chain FAs, hexanoic and octanoic acid, suggests they may be part of the fruit flies diet. Typical dietary fats, including many plant based oils, such as coconut oil, are rich in longer chain FAs including palmitic acid, oleic acid and linoleic acid [49]. However medium chain FAs are present in fermenting fruits such as guava and pollen, and it is possible this provides a substrate for Drosophila in the wild [50,51]. While less is known about the food substrates of D. melanogaster, the closely related D. sechelia consume Morinda fruit, which is rich in FAs [52,53]. Further, we have previously shown that a diet of HxA alone is sufficient for survival [19]. Therefore, it is possible that FA attraction evolved to promote consumption of calorically rich fruits consumed by Drosophila.
Our aversive memory assay indicates flies can discriminate between these attractive tastants. Sugar induces broad activation of Gr64f neurons, while the activation induced by HxA appears more restricted, and therefore it is possible that differences in activation allow for differentiation [54]. Alternatively, we find that HxA also activates IR56d neurons that emanate from the taste pegs and do not co-express Gr64f, so it is possible that the activation of these neurons by HxA allows discrimination. Finally, considering the different biochemical pathways involved in sugar and FA sensing [19], their identity can be coded by unique temporal and spatial dynamics of sensory neuron activation [16,55,56]. Differences in activation are suggested to provide a mechanism for olfactory discrimination within defined neural populations, and it is possible that similar mechanisms are utilized for attractive tastants [53]. In Drosophila, attractive tastants have been found to induce a wide range of excitatory responses ranging from acute to prolonged firing [29,57], providing a potential mechanism for discrimination. While the sensillar response to FAs has not been reported, the differences in Ca2+ response to sugar or HxA presentation within the SEZ suggest differences in temporal activation.
We find that flies can discriminate between sugar and FAs, but it is not known whether they can discriminate qualitatively between different classes of FAs. A previous study examining discrimination between different sugars found that flies are unable to discriminate based on quality, but could discriminate based on perceived palatability [45]. Here, we find that pentanoic acid elicits a robust PER response that is independent of IR56d neurons. The findings that distinct populations of neurons respond to FAs from different classes raises the possibility that flies may discriminate based on pattern of neuronal activation.
Role of Phospholipase C in fatty acid taste
We have previously reported evidence that Phospholipase C (PLC) signaling in sweet-sensing Gr64f neurons is required for FA taste [19]. Flies with mutation or knockdown of the PLC-ß ortholog no receptor potential A (norpA) do not respond to HxA or octanoic acid but respond normally to sugars, indicating that distinct intercellular signaling mechanism underlie response to FAs and sugars [19,58]. Our findings that the IR56d-Gr64f co-expressing neurons respond to both sugars and FAs provide a system to directly test the role of norpA in neuronal response within a defined neuronal population. Further, our findings show that PER response to pentanoic acid does not require IR56d neurons, suggesting distinct populations of taste neurons regulate response to short-chain (pentanoic acid) and medium-chain FAs (HxA and octanoic acid). Testing the response of norpA deficient flies to short-chain FAs will inform whether PLC is more generally required for FA taste, or is only specific to medium-chain FAs detected by IR56d neurons.
Higher order neurons involved in fatty acid perception
While much is known about taste coding within the SEZ, less is known about the higher order circuits governing taste. Sweet sensing neurons project to the antennal mechanosensory and motor center (AMMC) and downstream PAM dopaminergic neurons that are activated by sugar [39,59]. In addition, a separate populations of dopamine neurons, the PPL1 cluster, is required for olfactory appetitive memory and taste aversive conditioning [60–62]. A central question will be whether these higher order neurons downstream of the SEZ are also sensitive to FAs, or selectively tuned to sugar. To date, higher order neurons responsive to FA taste have not been identified. It is possible that sugar and FA taste signal through shared higher order dopaminergic neurons or, alternatively, each taste modality may activate distinct populations of higher order neurons that convey valence to the mushroom bodies, the memory and sensory integration center in Drosophila [63–65].
Conclusions
Taken together, this study provides insight into the coding of FAs within the fly gustatory system. Our results reveal a population of sweet-sensing neurons that are tuned for medium-chain FAs, but not short- or long-chain FAs. Further, the finding that flies are capable of discriminating between FAs and sugars suggests coding differences, either at the level of spatial or temporal neuronal activation, that provides a mechanism to distinguish between tastants of the same valence. Understanding how FAs are coded within the fly brain provides a model for understanding taste in more complex systems and will offer insight into generalizable mechanisms for taste discrimination.
Methods
Drosophila maintenance and fly stocks
Flies were grown and maintained on standard food (New Horizon Jazz Mix, Fisher Scientific). Flies were maintained in incubators (Powers Scientific; Dros52) at 25°C on a 12:12 LD cycle, with humidity set at 55-65%. The background control line used in this study is the w1118 fly strain unless otherwise noted. The following fly straits were ordered from The Bloomington Stock Center, UAS-imp-TNT (28840), UAS-TNT (28838), UAS-TRPA1 (26263); UAS-CaMPARI; (58762); UAS-GFP (32186); UAS-GCaMP5 (42037); +;; Gr43a-GAL4 (57637), Gr5a-GAL4 (57591), Gr61a-GAL4 (57658), Gr64a-GAL4 (57662), Gr64c-GAL4 (57663), Gr64d-GAL4 (57665), Gr64e-GAL4 (57666), Gr64f-GAL4 (57668), IR56b-GAL4 (60706), IR56d-GAL4 (60708), LexAop-Gal80 (32213). Gr64f-LexA was kind gift from H. Tanimoto and previously described in [20]. Seven to nine day old mated female flies were used for all experiments in this study, except when noted.
Proboscis Extension Reflex
For the initial screen using TNT, and specific testing of tarsal response, PER was measured by applying tastant to the tarsi, as previously described [19]. For all other PER experiments, including validation of Ir56d phenotypes, tastant was applied to the proboscis to match behavioral results with functional imaging. Flies were anesthetized on CO2, and mounted in a pipette tip so that their head and proboscis, but not tarsi, were exposed. After a 30-60m acclimation period, flies were presented with water and allowed to drink freely until water satiated. Flies that did not stop responding to water within 5 minutes were discarded. A small Kimwipe (Kimberley Clark) wick saturated with tastant was manually applied to the tip of the proboscis for 1-2s and proboscis extension reflex was monitored. Only full extensions were counted as a positive response. Each tastant was presented three times, with 10 seconds between each trial. Between tastant trials, the proboscis was washed with water and flies were allowed to drink. PER response was calculated as a percentage of proboscis extensions to total number of tastant stimulations. For experiments examining the effects of TRPA1 activation on PER, flies were mounted on a microscope slide using nail polish as described previously [19]. Flies were then placed on a heat plate heated to 34 °C and video of activity was recorded for 1 minute. The number of flies for each genotype showing PER within the 1 minute trial period was counted and the percentage of flies showing PER calculated.
In vivo functional imaging
Flies were anaesthetized on ice and placed in a small chamber with the head and proboscis accessible. A small hole was cut in tin foil and fixed to the stage leaving a window of cuticle exposed, then sealed using dental glue (Tetric Flow – Ivoclar Vivadent). The proboscis was extended and a small amount of dental glue was used to secure it in place, ensuring the same position throughout the experiment.
The cuticle and connective tissue was dissected to expose the SEZ, which was bathed in artificial hemolymph (140 mM NaCl, 2mM KCl, 4.5 mM MgCl2, 1.5mM CaCl2, and 5mM HEPES-NaOH with pH = 7.0). Mounted flies were placed under a confocal microscope (Nikon A1) and imaged using a 25x water-dipping objective lens. The pinhole was opened to allow a thicker optical section to be monitored. Recordings were taken at 4Hz with 256 resolution. Tastants were delivered to the proboscis for 1-2s with a KimWipe wick operated by microcontroller (brand, type). For analysis, regions of interest (ROI) were drawn manually, taking care to capture the same area between control and experimental. Baseline fluorescence was recorded over 5 frames, 10s prior to tastant application. ΔF/F was calculated for each frame as (fluorescence – baseline fluorescence)/baseline fluorescence * 100. Average fluorescence traces were created by taking the average and standard error of ΔF/F for all samples per frame.
Immunohistochemistry
Fly brains were dissected in ice-cold PBS and fixed in 4% formaldehyde, PBS,0.5% Triton-X 100 for 30 minutes. Brains were rinsed 3X with PBS, Triton-X for 10 minutes and incubated overnight at 4°C in NC82 (Iowa Hybridoma Bank [66]). The brains were rinsed again in PBS-TritonX, 3X for 10 minutes and placed in secondary antibodies (Donkey anti-Mouse 555; Life Technologies) for 90 minutes at room temperature. The brains were mounted in Vectashield (VectorLabs) and imaged on confocal microscope. Brains were imaged in 2um sections and are presented as the Z-stack projection through either the entire brain, or anterior and posterior regions of IR56d projections in the SEZ.
Aversive Taste Memory
PER induction was performed in one-week old mated females as described previously [5, 16]. One-week-old mated females were collected and placed on fresh food for 24 h, then starved for 48 h in food-vials on wet Kimwipe paper. Flies were later anaesthetized on CO2 pad and glued using nail polish (Cat#72180, Electron Microscopy Science) by their thorax and wing base on a microscopy slide and left to recover in a box with wet Kimwipe paper for 3-6 h prior to experiments. For experiments, the slide was mounted vertically under the dissecting microscope (Olympus SZX12) and PER was observed. Flies were satiated with water before and during the experiment. Flies that did not initially satiate within 5 minutes were excluded from conditioning. We used a 1ml syringe (Tuberculine, Becton Dickinson & Comp) for tastant presentation. We used purified water, 10 mM and 1000 mM sucrose, 0.4% hexanoic acid and 10 mM quinine hydrochloride solutions. The protocol was adapted from [40].Here, for pretest, each fly was given 10 mM sucrose or 0.4% HxA on their tarsi three times with 10 sec inter-trial interval and the number of full proboscis extensions was recorded. During training, same stimulation as before was followed by a droplet of 10 mM quinine was placed on extended proboscis and flies were allowed to drink it for up to 2 sec or until they retracted their proboscis. After each session, tarsi and proboscis were washed with water and flies were allowed to drink to satiation. After training, flies were tested with either that same substance without quinine or with the untrained substance (matched or opposite trained groups). Another group of flies was measured as described above but quinine was never presented (naïve groups). At the end of each experiment, flies were given 1000mM sucrose to check for retained ability of PER and non-responders were excluded [11].
Reagents
Sucrose was purchased from Fisher Scientific (FS S5-500). All other tastants were purchased from Sigma Aldrich. Sucrose, hexanoic acid (SA 153745), octanoic acid (SA C2875), pentanoic Acid (SA 240370) and quinine hydrochloride (SA 145904) were diluted in H20. Oleic Acid (SA O1008) was diluted in DMSO 1% DMSO (Sigma).
Statistical Analysis
All statistical tests were performed in R. For normally distributed data, Student’s t-test or ANOVA with Tukey’s post-hoc comparison was performed. For data that did not fit a normal distribution, nonparametric tests were used: Wilcoxon Rank-Sum or Kruskal-Wallis with Dunn’s post-hoc. Fisher’s Exact Test was used for binary categorical data. For all tests with multiple comparisons, Bonferroni p-value adjustment was performed.
Supplemental Data
Table S1. A neuronal silencing screen for taste neurons sensitive to HxA
(A) Data for PER in response to 100mM sucrose in flies with silenced populations of gustatory neurons by driving expression of TNT with the indicated. The ‘normalized’ column represents the PER value for experimental fly divided by the result for w1118. (B) Data for PER in response to 1% HxA. p-values computed by Kruskal-Wallis followed by Dunn’s post-hoc with Bonferroni p-value adjustment, with comparison to w1118 control flies. *p<0.05, **p<0.01, ***p<0.001.
Fig S1. Flies continue to suppress PER in response to FAs in absence of Gr64f to drive LexAop-Gal80
(A) PER for flies expressing either imp-TNT or TNT in IR56d neurons, (n = 31 for both groups), Without Gr64f-LexA to drive the LexAop-GAL80, as in Fig 5c, TNT is expressed in all IR56d neurons and PER to HxA is suppressed. Wilcoxon Rank Sum Test ***p<0.001
Acknowledgements
We are grateful to Michael Gordon (University of British Columbia), Seth Tomchik (Scripps, Florida), Anupama Dahanukar (University of California, Riverside) and Hiromu Tanimoto (Tohoku University, Japan) for technical support. This work was supported by NIH Grants 1R01 NS085252 to ACK and NSF IOS-1426265 to PM and ACK.