Neuroscience 284 (2015) 180–191

AMILORIDE-SENSITIVE SODIUM CURRENTS IN FUNGIFORM TASTE CELLS OF RATS CHRONICALLY EXPOSED TO NICOTINE A. BIGIANI *

these, is well recognized its action on the taste system. As taste substance, nicotine elicits bitter sensations in humans (Pfaffmann, 1959), it is coded as bitter stimuli in monkeys (Scott et al., 1999), and it induces aversive responses in hamsters (Brining et al., 1991). Consistent with these findings, nicotine activates gustducin, a specific G protein involved in bitter transduction (Ming et al., 1998, 1999). Finally, taste response to nicotine in rats and mice are mediated, at least in part, by molecular pathways involving the TRPM5 channel, which is required for the peripheral transduction of bitter, sweet, and umami chemicals (Oliveira-Maia et al., 2009). Thus, nicotine seems to interact with the molecular machinery involved in bitter detection. Besides being a taste stimulus by itself, nicotine can also affect sensory reception for other tastants. In rats, nicotine modulates chorda tympani response to salty stimuli by interacting with both the amiloride-sensitive pathway involving ENaC (epithelial sodium channel), and the amiloride-insensitive pathway involving TRPV1t channel, (Lyall et al., 2007). In addition to its acute effects, nicotine is believed to induce alterations in taste system during chronic exposure, a condition commonly found in smokers. In regular smokers or people using smokeless tobacco, the ability to detect taste stimuli is impaired, as indicated by the increase in the threshold for recognition of sweet, salty, sour, and bitter stimuli, and by the reduction in the perceived intensity at supra-threshold concentrations (e.g., Mela, 1989; Pomerleau et al., 1991; Sato et al., 2002; Yamauchi et al., 2002). Of interest for the impact on human health is the finding that smoking has been recognized as one of the agents that may decrease the effectiveness of the gustatory system to detect salt in foodstuff (Contreras, 1978; Rosenthal et al., 2001; Hooper et al., 2002; Slama et al., 2002). Indeed, an increase in consumption of saltier foods in smokers has been documented (Tell et al., 1984; Woodward et al., 1994; Mediavilla Garcia et al., 2001). Rodents have also been shown to respond to smoking or chronic nicotine exposure with alterations of taste behavior similar to those described in humans (e.g., Grunberg, 1982; Wager-Srdar et al., 1984; Etscorn et al., 1986; Parker and Doucet, 1995). A paper by Tomassini et al. (2007) has provided evidences that chronic exposure to nicotine alters morphometric features of taste buds in rat fungiform papillae, suggesting that this drug can have profound effects on the peripheral taste organs. It is not clear, however, how nicotine affects taste cell physiology during long-term administration.

Dipartimento di Scienze Biomediche, Metaboliche e Neuroscienze, Sezione di Fisiologia e Neuroscienze, Universita` di Modena e Reggio Emilia, via G. Campi 287, 41125 Modena, Italy

Abstract—Many studies have demonstrated that chronic exposure to nicotine, one of the main components of tobacco smoke, has profound effects on the functionality of the mammalian taste system. However, the mechanisms underlying nicotine action are poorly understood. In particular no information is available on the chronic effect of nicotine on the functioning of taste cells, the peripheral detectors which transduce food chemicals into electrical signals to the brain. To address this issue, I studied the membrane properties of rat fungiform taste cells and evaluated the effect of long-term exposure to nicotine on the amiloride-sensitive sodium currents (ASSCs). These currents are mediated by the epithelial sodium channels (ENaC) thought to be important, at least in part, in the transduction of salty stimuli. Patch-clamp recording data indicated that ASSCs in taste cells from rats chronically treated with nicotine had a reduced amplitude compared to controls. The pharmacological and biophysical analysis of ASSCs revealed that amplitude reduction was not dependent on changes in amiloride sensitivity or channel ionic permeability, but likely derived from a decrease in the activity of ENaCs. Since these channels are considered to be sodium receptors in taste cells, my results suggest that chronic exposure to nicotine hampers the capability of these cells to respond to sodium ions. This might represent a possible cellular mechanism underlying the reduced taste sensitivity to salt typically found in smokers. Ó 2014 IBRO. Published by Elsevier Ltd. All rights reserved.

Key words: taste cell, patch-clamp, nicotine, amiloridesensitive sodium currents, ENaC, smoking.

INTRODUCTION Nicotine is a pharmacologically active substance found in tobacco smoke that is able to determine profound biological effects (reviewed in: Yildiz, 2004). Among

*Tel: +39-059-205-5349; fax: +39-059-205-5363. E-mail address: [email protected] Abbreviations: ASSCs, amiloride-sensitive sodium currents; EGTA, ethylene glycol tetraacetic acid; ENaC, epithelial sodium channels; GHK, Goldman–Hodgkin–Katz; HEPES, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid; nAChRs, nicotinic acetylcholine receptors. http://dx.doi.org/10.1016/j.neuroscience.2014.09.077 0306-4522/Ó 2014 IBRO. Published by Elsevier Ltd. All rights reserved. 180

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I have addressed the issue of the chronic effect of nicotine on taste cell functioning by studying the membrane properties of rat fungiform taste cells. Specifically, I have applied the patch-clamp recording technique to single taste cells to evaluate whether longterm exposure to nicotine affected the amiloridesensitive sodium currents (ASSCs). These currents are mediated by ENaC, which represents one of the best characterized components of the sodium transduction mechanism (reviewed in Lindemann, 1996; Stewart et al., 1997; DeSimone and Lyall, 2006). The key role of ENaC in sodium taste detection, at least in rodents, has been recently established by Chandrashekar et al. (2010).

EXPERIMENTAL PROCEDURES Experiments were performed in compliance with the Italian law on animal care No. 116/1992, and in accordance with the European Community Council Directive (EEC/609/86). All efforts were made to reduce both animal suffering and the number of animals used. Animals Male Sprague–Dawley rats (70–90 days of age) were used in this study. Animals were housed two per cage on a 12-h light/dark cycle in climate-controlled conditions with ad libitum access to water and food. Nicotine treatment Two common modes to achieve chronic nicotine exposure in laboratory animals are drug administration via drinking water or daily subcutaneous injections (e.g., Liu et al., 2005). In this work, I administered nicotine to rats via drinking water. The oral administration route allowed a more continuous exposure to nicotine, resembling the one observed in habitual smokers. The oral administration procedure closely followed published protocols (e.g., Liu et al., 2005). Briefly, rats were given free access to ()nicotine bitartrate dehydrate (100 lg/mL, corresponding to about 200 lM of free nicotine) or to vehicle (100 lg/mL tartaric acid) in their drinking water (tap water) for at least 3 weeks before experimentation (Tomassini et al., 2007). At the concentration used, nicotine does not represent a taste stimulus for rats (Simons et al., 2006). Isolation of rat fungiform taste buds Electrophysiological experiments were performed on single taste cells in isolated taste buds. My procedure to isolate taste buds from fungiform papillae closely followed published protocols (e.g., Doolin and Gilbertson, 1996; Kossel et al., 1997). Taste buds were plated on the bottom of a chamber consisting of a standard glass slide onto which a silicon ring 1–2 mm thick and 15 mm ID was pressed. The glass slide was precoated with Cell-Tak (3 lg/cm2; Becton Dickinson, Milan, Italy) to improve adherence of isolated taste buds to the bottom of the chamber. The chamber was placed on the stage of an upright Olympus microscope (model BHWI), and taste buds were viewed with Nomarski optics. During the exper-

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iments, isolated taste buds were continuously perfused with Tyrode solution [containing (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 glucose, 10 sodium pyruvate, and 10 HEPES; pH 7.4] by means of a gravity-driven system. Patch-clamp recording Whole-cell patch-clamp recordings were made from cells in isolated taste buds as described previously (e.g., Doolin and Gilbertson, 1996; Kossel et al., 1997). Recording pipettes were made from soda lime glass capillaries (Baxter Scientific Products, McGaw Park, IL, USA) on a two-stage vertical puller (Model No. PP-830, Narishige, Tokyo, Japan). Typical pipette resistances were 2–4 MX when filled with a standard pipette solution containing (in mM) 120 KCl, 1 CaCl2, 2 MgCl2, 10 HEPES, 11 EGTA, and 2 ATP, pH 7.2 adjusted with KOH. All voltages have been corrected for liquid junction potential measured between pipette solution and Tyrode (bath) solution (Neher, 1992). Patched taste cells were identified on the basis of their voltage-gated ion currents as described previously (Bigiani and Cuoghi, 2007). Their ‘‘resting’’ membrane properties (zero-current potential, input resistance, and cell membrane capacitance) were analyzed as follows: The zero-current potential (V0) is the voltage at which no current is required in voltage-clamp, and it is assumed to be an estimation of the cell’s resting potential. It depends on the presence of ion channels constitutively open (‘‘resting’’ channels), but it is also affected by the value of seal resistance (Bigiani et al., 1996). To compare V0 between different cells, therefore, I have included in the analysis only cells with seal resistance larger than 5 GX (Bigiani et al., 1996), and discarded all the other ones. Input resistance (Rin) is defined as the ratio between the applied voltage and the current required to hold the cell at that voltage, and it is often used as an indirect estimation of the whole-cell membrane resistance (Bigiani et al., 1996). Rin was evaluated as slope resistance of the current–voltage (I–V) relationship at the holding potential of about 80 mV to avoid activation of voltage-gated currents that would alter its value. Cell membrane capacitance (Cm) is the capacitance of the whole-cell membrane, and it is directly related to the actual extension of the membrane lipid bilayer. Under whole-cell conditions, it was evaluated by integrating the capacitative current transient during application of a 10-mV voltage step from a holding potential of 80 mV (for details, see Bigiani and Roper, 1993). The presence of functional ASSCs in taste cells was monitored by studying the effect of bath-applied amiloride, a specific blocker of ENaC at micromolar concentrations, on the whole-cell current recorded at a given holding potential (e.g., Gilbertson et al., 1993; Doolin and Gilbertson, 1996; Lin et al., 1999; Bigiani and Cuoghi, 2007). Dose–response curves for the amiloride effect were obtained by measuring the change in stationary current (Iam) induced by adding increasing concentrations of blocker into the bath solution. The data were fitted to the equation:

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Iam =I50 ¼ ½am=f½am þ Ki g

which represents the single-binding isotherm (Hill coefficient = 1) for the blocking effect of amiloride, and where I50 is the amplitude of the response to 50 lM amiloride (maximal response), [am] is the variable amiloride concentration, and Ki is the apparent inhibition constant. Ion selectivity of ASSCs was evaluated by using the Goldman–Hodgkin–Katz (GHK) equation for membrane current (Hille, 2001). Briefly, current–voltage curves for the response to amiloride at different membrane potentials were fitted with the sum of two GHK functions representing independent cationic currents: a Na+ current and a K+ current flowing through opened ENaCs. From the reversal potential values of the responses to amiloride I evaluated the permeability ratio for Na+ and K+ (PNa/PK). Data analysis Analysis and plotting were performed using Prism 3.03 software (Graph Pad Software, San Diego, USA). Results are presented as mean ± standard error of the means (S.E.M.). Data comparisons were made with a Mann–Whitney test (MWT) or with X2 test analysis.

RESULTS Electrophysiological cell subtypes Mammalian taste cells are excitable cells expressing several types of voltage-gated ion currents (e.g., Be´he´ et al., 1990). Previously, we have identified two electrophysiological cell types in rat fungiform taste buds, ‘‘Type A’’ and ‘‘Type B’’ cells, endowed with large (>500 pA) voltage-gated sodium currents (an electrophysiological hallmark of ‘‘mature’’ taste cells: Mackay-Sim et al., 1996; Bigiani et al., 2002) and distinguishable on the basis of the relative amplitude of the voltage-gated potassium currents (Bigiani and Cuoghi, 2007). A third type of taste cells, which lack voltage-gated sodium currents and exhibit only voltage-gated potassium currents, was also found in fungiform taste buds from control rats. These cells, hereafter named as ‘‘Type C’’ cells, may include developing cells and/or glial-like cells (Bigiani, 2001; Medler et al., 2003; Romanov and Kolesnikov, 2006). Fig. 1A shows examples of voltage-gated ion currents recorded from these different subsets of taste cells. Pharmacological dissection with specific ion channel blockers allowed me to identify unambiguously the downward current deflections as voltage-gated sodium currents, and the upward current deflections as voltagegated potassium currents (data not shown). The relative occurrence of these cell types among patched cells (n = 224) in control rats is shown in Fig. 1B (CON). Morphological observations indicate that chronic nicotine treatment affects the size of taste buds by reducing the total number of their cells (Tomassini et al., 2007). Therefore, I asked whether the electrophysiological cell types could be identified also in animals chronically treated with nicotine, and whether they occurred among recorded cells in the same proportion found in

control animals. I then applied the patch-clamp recording technique to taste cells obtained from rats chronically treated with nicotine. I was able to identify unambiguously 119 cells from these animals and to assign them to one of the above categories (Fig. 1B, NIC). By comparing the occurrence of the three cell types among patched cells in control and nicotine-treated animals, I could conclude that nicotine did not change the relative proportion of functional types of taste cells when administered chronically (X2 test, P = 0.712). ‘‘Resting’’ membrane properties Next I asked whether chronic nicotine treatment affected the ‘‘resting’’ membrane properties (V0, Rin, Cm) of the three cell types identified in rat fungiform taste buds. These properties are by definition non voltage dependent and represent the reference set-point for the electrophysiological behavior of a given cell. To this aim, I performed patch-clamp experiments on taste cells isolated from control animals and nicotine-treated littermates. Data are collected in Fig. 2 and show that no significant differences were found for Type B and Type C cells. On the contrary, membrane properties of Type A cells changed significantly between the two experimental groups. Specifically, in cells from treated animals V0 became more negative with a concomitant increase in Rin. These changes could be interpreted as though depolarizing ion channels were shut off by nicotine treatment. Type A cells also had a larger surface membrane area in treated taste buds, as indicated by the Cm increase. As a whole, these results clearly indicated that nicotine affected a specific cell subset in fungiform taste buds, namely Type A cells. ASSCs To further characterize the effect of nicotine on the membrane properties of taste cells, I then analyzed the ASSCs. These currents are known to occur in rat fungiform taste cells (e.g., Doolin and Gilbertson, 1996; Kossel et al., 1997; Lin et al., 1999; Bigiani and Cuoghi, 2007) and are mediated by ENaCs, which are constitutively open and, therefore, affect both V0 and Rin values. Thus, I asked whether the observed modification of the ‘‘resting’’ membrane properties induced by nicotine in fungiform taste cells could be ascribed to an action on ASSCs. When cells are bathed by physiological saline, open ENaCs mediate an inward, stationary sodium current that can be recorded with the patch-clamp technique. In these conditions, amiloride can be used as a pharmacological probe to monitor ENaC-mediated current in taste cell membrane by evaluating its blocking effect. I used 1 lM amiloride to assure specificity of the effect. It is well known that amiloride, at larger concentrations, blocks not only ENaCs but also other ion channels and exchangers (Lindemann, 1996). Fig. 3A shows the effect of 1 lM amiloride application on the whole-cell current recorded from control taste cells at a membrane potential of about 80 mV. Amiloride caused a large reduction in the stationary inward current, hereafter referred as to ‘‘response to amiloride’’,

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Fig. 1. Electrophysiological cell types in rat fungiform taste buds. (A) Whole-cell, voltage-clamp recordings from taste cells held at about 80 mV and step depolarized (10-mV increments). On the basis of the occurrence and amplitude of voltage-gated sodium currents (downward deflections in the current records) and potassium currents (upward deflections in the current records), three main types could be identified. The large amplitude of sodium currents in ‘‘Type A’’ and ‘‘Type B’’ cells indicate that these are mature taste cells. Type C cells may represent immature or glia-like taste cells. Note the presence of a conspicuous holding current (arrow) in Type A cell. Dotted line: zero current. (B) Occurrence of taste cell types in control (CON) and nicotine-treated (NIC) rats. Statistical analysis of the relative proportion of identified cells for each subset (Cell Type) indicated no significant difference between control rats and nicotine-treated littermates. X2 test, P = 0.712.

only in Type A cells. Type B and Type C were unresponsive or slightly sensitive to amiloride (Fig. 3B; see also Bigiani and Cuoghi, 2007). I therefore evaluated the response to amiloride after chronic exposure to nicotine in Type A cells. I found that the amplitude of the response to 1 lM amiloride was markedly reduced in taste cells from nicotine-treated animals as compared to those obtained from control littermates (Fig. 4A). An interesting result was also that long-term administration of nicotine produced an increase in the relative proportion of cells lacking detectable amiloride response (see the position of the median in the box and whiskers plot on the right in Fig. 4B). I evaluated for each experimental group (control rats and nicotine-treated littermates) the percentage of ‘‘unresponsive cells’’, that is, cells with a response to 1 lM amiloride smaller than 10 pA. I found that in controls 22% of cells were unresponsive (19 out of 87 cells), whereas in treated animals 37% of cells did not produce sizeable amiloride response (24 out of 65 cells). This finding suggested that long-term administration of nicotine also reduced the number of taste cells endowed with functional ENaCs. For the sake of completeness, I also performed some recordings with Type B (n = 10) and Type C cells (n = 18) from nicotine-treated animals, without finding any significant changes in their negligible responses to amiloride documented in control animals (data not shown). A diminished amplitude of the response to amiloride in Type A cells could be due to a nicotine-induced reduction

in ENaC sensitivity to amiloride. This was evaluated by analyzing dose–response relationships, which were obtained by applying different drug concentrations to cells during whole-cell recording (Fig. 5A, B). By fitting the dose–response curves with a standard singlebinding isotherm equation (Fig. 5C), I found that the inhibition constant (Ki) for amiloride in taste cells of control rats was similar to the one evaluated in cells of nicotine-treated littermates (0.109 lM and 0.112 lM, respectively). Therefore, a reduced amplitude of the response to 1 lM amiloride in taste cells of treated animals was not caused by a reduced sensitivity of ENaCs to amiloride. Another possibility to explain the reduced response to amiloride was a nicotine-induced change in the ion selectivity of ENaC, which is permeable to Na+ but also to K+ (Lindemann, 1996). A reduction in the permeability ratio (PNa/PK) would cause a decrease in response amplitude at any membrane potential. I evaluated ENaC ion selectivity by studying the voltage dependence of the response to 1 lM amiloride (Fig. 6A). I found that the current–voltage relationship was not linear, as predicted by the GHK equation for a generic membrane current (Hille, 2001). Accordingly, data points could be nicely fitted to GHK function (Fig. 6B) and yielded a similar PNa/PK in taste cells from control rats and from nicotinetreated littermates (4.1 and 4.7, respectively). Note that according to these values, the amplitude of amiloride response at 84 mV should be even a little bit larger in

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DISCUSSION Chronic exposure to nicotine affects the anatomical properties of taste buds in rat fungiform papillae (Tomassini et al., 2007), raising the possibility that also their functional properties could undergo some changes. Here I showed that indeed in the long run nicotine has an effect on the membrane electrophysiological properties of a specific subset of taste cells in rat fungiform taste buds. Electrophysiological cell subtypes in fungiform taste buds

Fig. 2. ‘‘Resting’’ membrane properties of the electrophysiological cell types identified in rat fungiform taste buds. Zero-current potential (V0), input resistance (Rin) and cell membrane capacitance (Cm) were recorded from Type A, Type B, and Type C cells in both control rats (CON) and in nicotine-treated littermates (NIC). Histograms represent mean values ± SEM. Comparison of the values for each parameter in the two experimental groups revealed that there were significant changes only in Type A cells. MWT, ⁄P = 0.0274, ⁄⁄P = 0.0092, ⁄⁄⁄ P = 0.0067. Number of observations (n) in the two experimental groups (CON/NIC) as follows: Type A cells: V0, n = 56/33; Rin, n = 51/29; Cm, n = 37/37. Type B cells: V0, n = 16/11; Rin, n = 17/ 12; Cm, n = 8/11. Type C cells: V0, n = 33/15; Rin, n = 39/20; Cm, n = 22/17.

nicotine-treated taste cells than in control ones. This result clearly indicated that a reduced amplitude of the response to 1 lM amiloride in taste cells of treated animals was not caused by a change (a reduction) in the permeability ratio PNa/PK of ENaC.

In fungiform papillae of normal rats, three main types of taste cells can be identified on the basis of the occurrence of voltage-gated Na+ (INa) and K+ currents (IK), and on their relative amplitude (see also Bigiani and Cuoghi, 2007): Type A cells (‘‘mature’’ taste cells with large INa and small IK), Type B cells (‘‘mature’’ taste cells with large INa and large IK), and Type C cells (‘‘immature and/or glia-like’’ cells, lacking INa). These cell subtypes correspond nicely with the electrophysiologically identified cell subpopulations described in mouse taste buds (Romanov and Kolesnikov, 2006). In my preparation, their relative occurrence among patched cells (n = 224) was: 58% Type A, 13% Type B, 29% Type C. Of course, this cell typing does not include all the possible cells found in taste buds: given the continuous turnover that characterizes these sensory organs (e.g., Farbman, 1980), it would not be surprising to find cells with intermediate properties, and indeed this was the case also in my patch-clamp recording experiments. However, in this study I focused my attention only on these three welldefined categories since they displayed clear-cut membrane properties. Morphological observations indicate that long-term nicotine treatment reduces the size of fungiform taste buds by reducing the total number of their cells (Tomassini et al., 2007). My data indicate that the functional types of taste cells found in control animals still occur in the same proportion in fungiform taste buds of nicotine-treated animals: 56% Type A, 18% Type B, 26% Type C (n = 119 identified cells). Therefore, it is reasonable to conclude that nicotine exposure reduces the entire taste population without changing the pattern of relative proportion of functional cell types. Long-term effect of nicotine on sodium receptors (ENaCs) Chronic administration of nicotine induced a significant reduction in the ASSC in Type A cells. This current is mediated by the ENaCs, which are constitutively open and produce an inward, depolarizing current when taste cells are bathed in sodium-rich saline. The amplitude of the current blocked by amiloride (response to amiloride) was on average about twice larger in control cells than in treated ones. The analysis of the response to amiloride in terms of its drug sensitivity (Ki) and its ion selectivity (PNa/PK) suggested that the basic biophysical and pharmacological features of ENaCs did not change

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Fig. 3. Response to amiloride in the electrophysiological cell types identified in rat fungiform taste buds. (A) Sample responses recorded by holding the cell membrane at about 80 mV. Amiloride (1 lM) was bath applied (Am, boxes under records). Note that amiloride induced a conspicuous reduction in the stationary inward current (Is, dashed line) only in Type A cell. Type B and Type C cells in this example were unresponsive to amiloride. IAm, amplitude of the response to amiloride. (B) Comparison of the amplitude of responses to 1 lM amiloride measured in the three cell types. Histograms represent mean values ± SEM (n = 87/10/14).

Fig. 4. Effect of long-term exposure to nicotine on the response to amiloride in Type A cells. (A) Sample response to amiloride recorded from a Type A cell of a control rat (CON) and from a cell of a nicotine-treated animal (NIC). Cells were held at about 80 mV and amiloride (1 lM) was bath applied (Am, boxes over records). Note that amiloride induced a larger reduction in the stationary inward currents (dashed line) in control cells than in cells from treated animals. (B) Distribution of the amplitude of the response to amiloride in the two experimental groups (CON, 87 cells from control animals; NIC, 65 cells from nicotine-treated littermates) represented in the form of box and whiskers plots. Boxes show the middle half of the data (the 25th and 75th percentiles) and the horizontal line marks the median, whereas the ‘‘whiskers’’ extending from the top and the bottom of the boxes show the main body of the data. Asterisks: Statistical analysis indicated that the two distributions were significantly different (MWT, P < 0.001).

in nicotine-treated animals. In other words, differences in the amplitude of the response to amiloride were not due to changes in the sensitivity to amiloride or to a change in ion selectivity of sodium channels in taste cells from nicotine-

treated animals. Since changes in ENaC molecule, such as subunit composition or covalent modifications, are associated with changes in amiloride sensitivity and ion selectivity (e.g., Ismailov et al., 1995; McNicholas and

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Fig. 5. Sensitivity to amiloride of Type A cells. (A) Responses to increasing concentrations of amiloride recorded from a cell of a control rat. The cell was held at about 80 mV and amiloride was bath applied (boxes over records). IS, stationary inward current; I50, amplitude of response to 50 lM amiloride. (B) Responses to increasing concentrations of amiloride recorded from a cell of a nicotine-treated rat. The cell was held at about 80 mV and amiloride was bath applied (boxes over records). (C) Dose–response curves for the effect of amiloride on Type A cells from control (CON) and nicotine-treated (NIC) rats. Amplitude of the responses (IAm) was normalized to the amplitude of the one obtained with 50 lM amiloride (I50). Points represent mean ± SEM (bars) of 5 and 6 cells, respectively. Sigmoidal curves are the best fit of the data for a single-binding isotherm, which yields an apparent inhibition constant (Ki) of 0.109 lM in controls and 0.112 lM in nicotine-treated animals.

Canessa, 1997), the electrophysiological data suggest that nicotine treatment unlikely affected the molecular structure or the biochemical status of ENaC. Biophysical analysis of cloned ENaC channels expressed in heterologous system has revealed that changes in amiloride sensitivity, in ion selectivity, and single-channel conductance are strictly inter-related: variation of one of these parameters is often associated with changes in the other two (Ismailov et al., 1995;

McNicholas and Canessa, 1997). Given that amiloride sensitivity and ion selectivity did not change after nicotine treatment, it is conceivable that also single-channel conductance of ENaC did not change in my preparation. Therefore, data could be interpreted on the basis of a reduction in the ‘‘activity’’ of functional ENaCs in the membrane of taste cells following nicotine exposure. The activity of ion channels is defined as the number of functional channels (N) times the single-channel open probability

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Fig. 6. Current–voltage relationships for the response to amiloride in Type A cells. (A) Sample records of the response to 1 lM amiloride (Am, boxes over records) elicited at different membrane potentials in a control taste cell (CON) and in a cell from a nicotine-treated rat (NIC). (B) Amplitude of the response to amiloride (IAm) recorded at a given membrane potential was plotted against membrane potential (Vm). The I–V plots tended to be nonlinear, exhibiting the curvature predicted by the Goldman–Hodgkin–Katz (GHK) function. Points represent mean ± SEM (n = 8– 11) and were fitted with the sum of two GHK functions representing independent Na+ and K+ currents flowing through opened amiloride-sensitive sodium channels (ASSCs). Note that the reversal potentials (arrows) were very close, indicating a similar permeability ratio (PNa/PK) for ASSCs in taste cells from both experimental groups.

(P0) (Ling et al., 1992). Of course, activity decrease could be due to a reduced number of N in the cell membrane or to a decrease in P0. In any case, a reduced activity of functional sodium channels would produce a smaller electrical signal during application of the blocker amiloride (reduced response). A reduction in the activity of ENaCs in Type A cells from treated animals could explain the observed increase in Rin (less channels open in taste cells

membrane) and the more negative value of V0 (depolarizing channels less active). Regarding the mechanism by which nicotine would produce a reduction in ENaC activity in taste cell membrane, my data do not allow to make any conclusion at this moment. An interesting possibility is that nicotine could affect ENaC trafficking in taste cells. ENaC is subjected to a continuous turnover in other cell

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types and the density of functional channels on the cell membrane is the result of the equilibrium between channel exocytosis and endocytosis (for a review, see: Rotin et al., 2001; Snyder, 2005). A decrease in the exocytosis or an increase in endocytosis would have an impact on the number of functional channels occurring at the cell membrane. In human nasal epithelial cells maintained in cultures, bath-applied nicotine has been reported to induce endocytosis of amiloride-sensitive sodium channels with consequent reduction in the overall transport of Na+ across the cell membrane and reduction in the capacitance of cell monolayers (Klimek et al., 2000). If the same mechanism occurs in taste cells, then one should expect a reduction in Cm after nicotine exposure with a concomitant reduction in ASSC. However, Cm underwent a significant increase (about 25%) in Type A cells during nicotine treatment. This is consistent with an increase in the cell surface area, that is, Type A cells had a more expanded membrane lipid bilayer. A possibility is that also in taste cells nicotine induces an initial endocytosis of ENaCs, which would be associated with a reduction in Cm. However, in the long run (chronic conditions) taste cells could try to compensate for the lack of functional channels in the cell membrane by accelerating the exocytosis of newly formed ENaCs. Likely, this could result in the insertion of premature, not yet functional channels. Thus, an increased Cm observed in taste cells from nicotine-treated animals might indicate an increased exocytosis of non-functional amiloride-sensitive sodium channels. At physiological pH, nicotine occurs mainly as cation (Nielsen and Rassing, 2002) and it is expected to affect taste cells from the extracellular space. Recently, expression of nicotinic acetylcholine receptors (nAChRs) has been demonstrated in rat fungiform taste buds by Oliveira-Maia et al. (2009). Interestingly, these authors suggest that ‘‘the effects of cigarette smoking on food consumption and taste preferences could result, among other factors, from peripheral taste modulation due to chronic exposure to nicotine, acting on nAChRs in taste buds’’. Thus, it is tempting to speculate that the chronic effect of nicotine on the ASSCs might be produced by long-term stimulation of taste nAChRs by nicotine. Molecular, genetic, immunocytochemical, and functional studies on mice have shown that taste buds comprise distinct functional subsets of cells (reviewed in: Chaudhari and Roper, 2010). Among them, the so-called Type II cells express membrane receptors and downstream signaling molecules involved in detecting sweet, bitter, and umami stimuli (e.g., Clapp et al., 2006; DeFazio et al., 2006). In addition, they exhibit voltagegated sodium and potassium currents, but lack voltagegated calcium currents (Medler et al., 2003; Vandenbeuch et al., 2008). Interestingly, mouse taste cells believed to be sodium sensors form an ENaCexpressing cell population separate from taste cells endowed with sweet, bitter or umami markers (Chandrashekar et al., 2010). By patching onto isolated Type II cells identified on the basis of the expression of GFP under the TRPM5 promoter, Vandenbeuch et al.

(2008) found that these cells did not exhibit amiloride-sensitive currents. Unexpectedly, they recorded these currents from ‘‘unlabelled’’ cells lacking voltage-gated sodium currents (an electrophysiological hallmark of Type I cells, thought to be glia-like cells: Medler et al., 2003). Thus, they concluded that likely Type I cells were the sodium sensors in mouse fungiform taste buds. Type A cells described here possess large voltagegated sodium currents and are expected to be chemosensitive in function given the expression of amiloride-sensitive sodium channels (e.g., Chandrashekar et al., 2010). According to mouse data, however, ENaC occurs in Type I cells, which lack voltage-gated sodium channels (Vandenbeuch et al., 2008). It is known that rats and mice show several molecular and physiological differences in taste reception. For example, acid-sensing mechanisms rely upon different ion channels (Lin et al., 2002; Ugawa et al., 2003; Richter et al., 2004; Huang et al., 2006). Also, there are significant differences as to the proportion of taste cells immune-reactive for specific transduction and synaptic markers (Ma et al., 2007). Finally, sodium detection threshold is much lower in Sprague–Dawley rats (Lu et al., 2009) than in C57BL/6J mice (Eylam and Spector, 2003). Thus, it is conceivable that the discrepancy observed for the functional subtypes expressing amiloride-sensitive sodium channels in rats and mice could reflect difference between species, as also suggested previously (e.g., Vandenbeuch et al., 2008). In any case, the present data indicate that chronic treatment with nicotine produces a reduction in the activity of amiloride-sensitive sodium channels. It is tempting to speculate that rat taste cells likely become less sensitive to sodium ions after nicotine exposure. A reduced activity of functional sodium receptors (ENaCs) would produce a smaller electrical signal when taste cells are stimulated with an increase in external sodium concentration. Therefore, a reduced activity of ENaCs would be reflected in taste cells being less sensitive to sodium variations in the saliva. Indeed, behavioral experiments have shown that chronic administration of nicotine alters salt taste preference and affects saline consumption in rats (Wager-Srdar et al., 1984). The present findings suggest that the long-term effects of nicotine on taste cell functioning is highly specific, targeting a particular cell subset (Type A cells), at least with regard to ASSCs. It would be interesting to test whether long-term administration of nicotine can exert any effects on voltage-gated membrane currents expressed by Type A, but also by Type B cells and by Type C cells. In nociceptors of rat trigeminal ganglia, acute application of nicotine inhibits voltage-gated Na+ currents (Liu et al., 2004). Also of interest it will be the evaluation of the chronic effect of nicotine on the L-type voltage-gated Ca2+ current, which occurs specifically only in Type B cells (Bigiani and Cuoghi, 2007), which resemble Type III cells of mouse taste buds (Medler et al., 2003; Clapp et al., 2006; DeFazio et al., 2006). Further studies are required to address all these issues.

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Mechanism of salt taste deterioration in smokers My results on rats indicate that chronic administration of nicotine inhibits the activity of ENaC, which represents the ‘‘sodium receptor’’, for it allows an influx of sodium ions into taste cells when the saliva sodium concentration increases (Lindemann, 1996; Stewart et al., 1997; DeSimone and Lyall, 2006). Therefore, nicotine disrupts the first molecular step in sodium detection. The net result of this action would be a reduced capability of taste cells to detect and to discriminate sodium ions, which might impair the sensory analysis performed by taste system on salty food. An obvious question now is whether this mechanism has general validity. Specifically, is it conceivable that a similar mechanism could explain the reduced taste sensitivity to sodium ions (salty taste) in smokers (Contreras, 1978; Rosenthal et al., 2001; Hooper et al., 2002; Slama et al., 2002)? There are indications that ENaC may be involved in sodium taste detection in humans. First, amiloride effectively reduces intensity of perceived saltiness when applied to the tongue mucosa (Schiffman et al., 1983; Tennissen, 1992; Tennissen and McCutcheon, 1996). Second, measurements of the human lingual surface potential indicate that a functional amiloride-sensitive pathway contributes to salt-evoked lingual potential in the anterior tongue (where fungiform papillae are located), although a great variability exists among individuals (Feldman et al., 2003, 2009). Third, the occurrence of ENaC a, b and c subunits in human fungiform papillae has been confirmed by polymerase chain reaction (PCR) analysis (Rossier et al., 2004). Finally, recent studies using a combination of psychophysical approach and functional expression of ENaC in Xenopus oocytes support the notion that ENaC participates in salt taste perception in humans (Sta¨hler et al., 2008). On the basis of these evidences, we may infer that long-term exposure to nicotine could lead to an inhibition of amiloride-sensitive sodium channels in fungiform taste cells of those individuals expressing ENaC. Since both amiloride-sensitive and amiloride-insensitive components have been described for sodium taste in humans (Schiffman et al., 1983; Tennissen, 1992; Ossebaard and Smith, 1995; Smith and Ossebaard, 1995; Tennissen and McCutcheon, 1996; Anand and Zuniga, 1997; Ossebaard et al., 1997; Feldman et al., 2003), the effect of nicotine would depend on the relative importance of the amiloride-sensitive pathway compared to the amiloride-insensitive one in any given individual. In this regard, it is important to underscore that there is a large heterogeneity among individuals as to the amiloride sensitivity of salt taste (Feldman et al., 2003). In any case, the effect on ENaC may explain, at least in part, why salt taste perception is blunted in smokers, who therefore tend to ingest saltier food to compensate a sensory deficit.

CONCLUSION Sodium channels sensitive to amiloride are thought to be salt detectors in taste cells. Chronic exposure to nicotine reduces the activity of these channels, as indicated by the marked decrease in the ASSCs recorded from single

taste cells. The effect of nicotine on these currents represent a first demonstration that in the long run nicotine may impair taste reception by acting on the membrane properties of taste cells, the sensory receptors for food chemicals. Acknowledgment—Research described in this article was supported by Philip Morris USA Inc. and by Philip Morris International.

REFERENCES Anand KK, Zuniga JR (1997) Effect of amiloride on suprathreshold NaCl, LiCl, and KCl salt taste in humans. Physiol Behav 62:925–929. Be´he´ P, DeSimone JA, Avenet P, Lindemann B (1990) Membrane currents in taste cells of the rat fungiform papilla – evidence for two types of Ca currents and inhibition of K currents by saccharin. J Gen Physiol 96:1061–1084. Bigiani A (2001) Mouse taste cells with glialike membrane properties. J Neurophysiol 85:1552–1560. Bigiani A, Cuoghi V (2007) Localization of amiloride-sensitive sodium current and voltage-gated calcium currents in rat fungiform taste cells. J Neurophysiol 98:2483–2487. Bigiani A, Roper SD (1993) Identification of electrophysiologically distinct cell subpopulations in Necturus taste buds. J Gen Physiol 102:143–170. Bigiani A, Kim D-J, Roper SD (1996) Membrane properties and cell ultrastructure of taste receptor cells in Necturus lingual slices. J Neurophysiol 75:1944–1956. Bigiani A, Cristiani R, Fieni F, Ghiaroni V, Bagnoli P, Pietra P (2002) Postnatal development of membrane excitability in taste cells of the mouse vallate papilla. J Neurosci 22:493–504. Brining SK, Belecky TL, Smith DV (1991) Taste reactivity in the hamster. Physiol Behav 49:1265–1272. Chandrashekar J, Kuhn C, Oka Y, Yarmolinsky DA, Hummler E, Ryba NJP, Zuker CS (2010) The cells and peripheral representation of sodium taste in mice. Nature 464:297–301. Chaudhari N, Roper SD (2010) The cell biology of taste. J Cell Biol 190:285–296. Clapp TR, Medler KF, Damak S, Margolskee RF, Kinnamon SC (2006) Mouse taste cells with G protein-coupled taste receptors lack voltage-gated calcium channels and SNAP-25. BMC Biol 4:7. http://dx.doi.org/10.1186/1741-7007-4-7. Contreras RJ (1978) Salt taste and disease. Am J Clin Nutr 31:1088–1097. DeFazio RA, Dvoryanchikov G, Maruyama Y, Kim JW, Pereira E, Roper SD, Chaudhari N (2006) Separate populations of receptor cells and presynaptic cells in mouse taste buds. J Neurosci 26:3971–3980. DeSimone JA, Lyall V (2006) Taste receptors in the gastrointestinal tract. III. Salty and sour taste: sensing of sodium and protons by the tongue. Am J Physiol Gastrointest Liver Physiol 291:G1005–G1010. Doolin RE, Gilbertson TA (1996) Distribution and characterization of functional amiloride-sensitive sodium channels in rat tongue. J Gen Physiol 107:545–554. Eylam S, Spector AC (2003) Oral amiloride treatment decreases taste sensitivity to sodium salts in C57BL/6J and DBA/2J mice. Chem Senses 28:447–458. Etscorn F, Moore GA, Hagen LS, Caton TM, Sanders DL (1986) Saccharin aversions in hamsters as a result of nicotine injections. Pharmacol Biochem Behav 24:567–570. Farbman AI (1980) Renewal of taste bud cells in rat circumvallate papillae. Cell Tissue Kinet 13:349–357. Feldman GM, Mogyoro´si A, Heck GL, DeSimone JA, Santos CR, Clary RA, Lyall V (2003) Salt-evoked lingual surface potential in humans. J Neurophysiol 90:2060–2064.

190

A. Bigiani / Neuroscience 284 (2015) 180–191

Feldman GM, Heck GL, Smith NL (2009) Human salt taste and the lingual surface potential correlate. Chem Senses 34:373–382. Gilbertson TA, Roper SD, Kinnamon SC (1993) Proton currents through amiloride-sensitive Na+ channels in isolated hamster taste cells: enhancement by vasopressin and cAMP. Neuron 10:931–942. Grunberg NE (1982) The effects of nicotine and cigarette smoking on food consumption and taste preferences. Addict Behav 7:317–331. Hille B (2001) Ion channels of excitable membranes. Sunderland, : Sinauer. Hooper L, Bartlett C, Davey Smith G, Ebrahim S (2002) Systematic review of long term effects of advice to reduce dietary salt in adults. BMJ 325:1–9. Huang AL, Chen X, Hoon MA, Chandrashekar J, Guo W, Tra¨nkner D, Ryba NJP, Zuker CS (2006) The cells and logic for mammalian sour taste detection. Nature 442:934–938. Ismailov II, Berdiev BK, Benos DJ (1995) Biochemical status of renal epithelial Na+ channels determines apparent channel conductance, ion selectivity, and amiloride sensitivity. Biophys J 69:1789–1800. Klimek T, Glanz H, Ru¨ckes-Nilges C, Van Driessche W, Weber WM (2000) Nicotine-induced endocytosis of amiloride-sensitive sodium channels in human nasal epithelium. Acta Otolaryngol 120:286–290. Kossel AK, McPheeters M, Lin W, Kinnamon SC (1997) Development of membrane properties in taste cells of fungiform papillae: functional evidence for early presence of amiloride-sensitive sodium channels. J Neurosci 17:9634–9641. Lin W, Finger TE, Rossier BC, Kinnamon SC (1999) Epithelial Na+ channel subunits in rat taste cells: localization and regulation by aldosterone. J Comp Neurol 405:406–420. Lin W, Ogura T, Kinnamob SC (2002) Acid-activated currents in rat vallate taste receptor cells. J Neurophysiol 88:133–141. Lindemann B (1996) Taste reception. Physiol Rev 76:719–766. Ling BN, Kokko KE, Eaton DC (1992) Inhibition of apical Na+ channels in rabbit cortical collecting tubules by basolateral prostaglandin E2 is modulated by protein kinase C. J Clin Invest 90:1328–1334. Liu L, Zhu W, Zhang Z-S, Yang T, Grant A, Oxford G, Simon SA (2004) Nicotine inhibits voltage-dependent sodium channels and sensitizes vanilloid receptors. J Neurophysiol 91:1482–1491. Liu J-J, Mohila CA, Gong Y, Govindarajan N, Onn S-P (2005) Chronic nicotine exposure during adolescence differentially influences calcium-binding proteins in rat anterior cingulated cortex. Eur J Neurosci 22:2462–2474. Lu B, Yan J, Yang X (2009) Effects of sodium depletion on detection thresholds for salty taste in rats. Physiol Behav 97:463–469. Lyall V, Phan TH, Mummalaneni S, Mansouri M, Heck GL, Kobal G, Desimone JA (2007) Effect of nicotine on chorda tympani responses to salty and sour stimuli. J Neurophysiol 98:1662–1674. Ma H, Yang R, Thomas SM, Kinnamon JC (2007) Qualitative and quantitative differences between taste buds of the rat and mouse. BMC Neurosci 8:5. Mackay-Sim A, Delay RJ, Roper SD, Kinnamon SC (1996) Development of voltage-dependent currents in taste receptor cells. J Comp Neurol 365:278–288. McNicholas CM, Canessa CM (1997) Diversity of channels generated by different combinations of epithelial sodium channel subunits. J Gen Physiol 109:681–692. Mediavilla Garcia JD, Fernandez Torres C, Aliaga Martinez L, Leon Ruiz L, Sabio Sanchez M, Jimenez Alonso J (2001) Clinical characteristics of patients with essential hypertension regarding salt intake. Rev Clin Esp 201:627–631. Medler KF, Margolskee RF, Kinnamon SC (2003) Electrophysiological characterization of voltage-gated currents in defined taste cell types of mice. J Neurosci 23:2608–2617. Mela DJ (1989) Gustatory function and dietary habits in users and nonusers of smokeless tobacco. Am J Clin Nutr 49:482–489.

Ming D, Ruiz-Avila L, Margolskee RF (1998) Characterization and solubilization of bitter-responsive receptors that couple to gustducin. Proc Natl Acad Sci USA 95:8933–8938. Ming D, Ninomiya Y, Margolskee RF (1999) Blocking taste receptor activation of gustducin inhibits gustatory responses to bitter compounds. Proc Natl Acad Sci USA 96:9903–9908. Neher E (1992) Correction for liquid junction potentials in patch clamp experiments. Methods Enzymol 207:123–131. Nielsen HM, Rassing MR (2002) Nicotine permeability across the buccal TR146 cell culture model and porcine buccal mucosa in vitro: effect of pH and concentration. Eur J Pharm Sci 16:151–157. Oliveira-Maia AJ, Stapleton-Kotloski JR, Lyall V, Phan TH, Mummalaneni S, Melone P, Desimone JA, Nicolelis MA, Simon SA (2009) Nicotine activates TRPM5-dependent and independent taste pathways. Proc Natl Acad Sci USA 106:1596–1601. Ossebaard CA, Smith DV (1995) Effect of amiloride on the taste of NaCl, Na-gluconate and KCl in humans: implications for Na+ receptor mechanisms. Chem Senses 20:37–46. Ossebaard CA, Polet IA, Smith DV (1997) Amiloride effects on taste quality: comparison of single and multiple response category procedures. Chem Senses 22:267–275. Parker LA, Doucet K (1995) The effects of nicotine and nicotine withdrawal on taste reactivity. Pharmacol Biochem Behav 52:125–129. Pfaffmann C (1959) The sense of taste. In: Field J, Magoun HW, Hall VE, editors. Handbook of physiology: neurophysiology, Vol. 1. Washington, DC: American Physiological Society. p. 507–533. Pomerleau C, Garcia AW, Drewnowski A, Pomerleau OF (1991) Sweet taste preference in women smokers: comparison with nonsmokers and effects of menstrual phase and nicotine abstinence. Pharmacol Biochem Behav 40:995–999. Richter TA, Dvoryanchikov GA, Roper SD, Chaudhari N (2004) Acidsensing ion channel-2 is not necessary for sour taste in mice. J Neurosci 24:4088–4091. Romanov RA, Kolesnikov SS (2006) Electrophysiologically identified subpopulations of taste bud cells. Neurosci Lett 395:249–254. Rosenthal T, Shamiss A, Holtzman E (2001) Dietary electrolytes and hypertension in the elderly. Int Urol Nephrol 33:575–582. Rossier O, Cao J, Huque T, Spielman AI, Feldman RS, Medrano JF, Brand JG, le Coutre J (2004) Analysis of a human fungiform papillae cDNA library and identification of taste-related genes. Chem Senses 29:13–23. Rotin D, Kanelis V, Schild L (2001) Trafficking and cell surface stability of ENaC. Am J Physiol Renal Physiol 281:F391–F399. Sato K, Endo S, Tomita H (2002) Sensitivity of three loci on the tongue and soft palate to four basic tastes in smokers and nonsmokers. Acta Otolaryngol Suppl 546:74–82. Schiffman SS, Lockhead E, Maes FW (1983) Amiloride reduces the taste intensity of Na+ and Li+ salts and sweeteners. Proc Natl Acad Sci USA 80:6136–6140. Scott TR, Giza BK, Yan J (1999) Gustatory neural coding in the cortex of the alert cynomolgus macaque: the quality of bitterness. J Neurophysiol 81:60–71. Simons CT, Boucher Y, Carstens MI, Carstens E (2006) Nicotine suppression of gustatory responses of neurons in the nucleus of the solitary tract. J Neurophysiol 96:1877–1886. Slama M, Susic D, Frohlich ED (2002) Prevention of hypertension. Curr Opin Cardiol 17:531–536. Smith DV, Ossebaard CA (1995) Amiloride suppression of the taste intensity of sodium chloride: evidence from direct magnitude scaling. Physiol Behav 57:773–777. Snyder PM (2005) Regulation of epithelial Na+ channel trafficking. Endocrinology 146:5079–5086. Sta¨hler F, Riedel K, Demgensky S, Neumann K, Dunkel A, Ta¨ubert A, Raab B, Behrens M, Raguse J-D, Hofmann T, Meyerhof W (2008) A role of the epithelial sodium channel in human salt taste transduction? Chem Percept 1:78–90. Stewart RE, DeSimone JA, Hill DL (1997) New perspectives in a gustatory physiology: transduction, development, and plasticity. Am J Physiol 272:C1–C26.

A. Bigiani / Neuroscience 284 (2015) 180–191 Tell GS, Klepp KI, Vellar OD, McAlister A (1984) Preventing the onset of cigarette smoking in Norwegian adolescents: the Oslo youth study. Prev Med 13:256–275. Tennissen AM (1992) Amiloride reduces intensity responses of human fungiform papillae. Physiol Behav 51:1061–1068. Tennissen AM, McCutcheon NB (1996) Anterior tongue stimulation with amiloride suppresses NaCl saltiness, but not citric acid sourness in humans. Chem Senses 21:113–120. Tomassini S, Cuoghi V, Catalani E, Casini G, Bigiani A (2007) Longterm effects of nicotine on rat fungiform taste buds. Neuroscience 147:803–810. Ugawa S, Yamamoto T, Ueda T, Ishida Y, Inagaki A, Nishigaki M, Shimada S (2003) Amiloride-insensitive currents of the acidsensing ion channel-2a (ASIC2a)/ASIC2b heteromeric sour-taste receptor channel. J Neurosci 23:3616–3622.

191

Vandenbeuch A, Clapp TR, Kinnamon SC (2008) Amiloride-sensitive channels in type I fungiform taste cells in mouse. BMC Neurosci 9:1. Wager-Srdar SA, Levine AS, Morley JE, Hoidal JR, Niewoehner DE (1984) Effects of cigarette smoke and nicotine on feeding and energy. Physiol Behav 32:389–395. Woodward M, Bolton-Smith C, Tunstall-Pedoe H (1994) Deficient health knowledge, diet, and other lifestyles in smokers: is a multifactorial approach required? Prev Med 23:354–361. Yamauchi Y, Endo S, Yoshimura I (2002) A new whole-mouth gustatory test procedure. II. Effects of aging, gender and smoking. Acta Otolaryngol Suppl 546:49–59. Yildiz D (2004) Nicotine, its metabolism and an overview of its biological effects. Toxicon 43:619–632.

(Accepted 30 September 2014) (Available online 8 October 2014)