Taste sensation for five basic tastes (sweet, bitter, umami, sour, and salty) is initiated by the interaction between taste substances and taste receptor cells in the mouth. Taste receptor cells express G protein-coupled or ionotropic taste receptors on their apical membranes, which are exposed to a harsh environment, such as drastic changes in osmotic pressure or pH. In contrast, their basolateral membranes are present in stable physiological conditions. Hence, the maintenance of their physiological environment is essential to investigate the signal transduction mechanisms of taste. A previous study used in situ whole-cell patch clamping in which taste substances were applied only to the apical membranes. This study indicated that mouse taste receptor cells generated depolarizing receptor potentials with action potentials in response to NaCl and HCl (Ohtubo et al., 2001), which activate ionotropic taste receptors (Zhang et al., 2019; Heck et al., 1984). Although taste receptor cells generate action currents that are mediated by the sweet, bitter, and umami G protein-coupled taste receptors (Furue and Yoshii, 1997; Yoshida et al., 2006; Noguchi et al., 2003), the receptor potential changes were unknown because the cell-attached or extracellular loose-seal recordings were used to prevent the washout of cytosolic components. Furthermore, as the activation of G protein-coupled taste receptors induces intracellular Ca²⁺ concentration changes (Dando and Roper, 2009; Huang and Roper, 2010; Roebber et al., 2019), Ca²⁺ imaging was chosen for investigating the taste signal transduction. Therefore, despite the existence of several studies on taste responses at taste receptor cells, the receptor potentials mediated by G protein-coupled taste receptors remain unclear.

A single taste bud contains 10–100 taste bud cells (TBCs). Elongated TBCs are classified into three cell types (type I, II, and III) based on their morphology and functions (Murray, 1973; Chaudhari and Roper, 2010; Roper, 2013). Of the average 42 TBCs of a single fungiform taste bud of a mouse (Ohtubo and Yoshii, 2011), type I cells are the most abundant and are believed to function as glia-like supporting cells (Lawton et al., 2000; Bartel et al., 2006). Type II cells account for approximately 25% of a taste bud (Ohtubo and Yoshii, 2011) and function as taste receptor cells by expressing different G protein-coupled receptors for sweet, umami, and bitter taste substances (Nelson et al., 2002; Nelson et al., 2001; Chandrashekar et al., 2000; Adler et al., 2000). Type III cells comprise approximately 5% of a taste bud (Ohtubo and Yoshii, 2011) and are sensitive to acid (Huang et al., 2008; Kataoka et al., 2008). Hence, approximately 30% of TBCs are taste receptor cells (type II and type III cells). As individual taste receptor cells encode individual taste qualities (Adler et al., 2000; Nelson et al., 2001), it is difficult to obtain stable recordings in response to all five tastes from a given taste receptor cell in a single taste bud. It is likely that these essential features of TBCs contribute to the difficulty in obtaining electrophysiological recordings from taste receptor cells in response to taste substances that activate G protein-coupled taste receptors.

In the present study, we performed an “in situ perforated whole-cell patch clamping” to TBCs in the peeled lingual epithelium to elucidate the receptor potential changes mediated by G protein-coupled taste receptors (Figure 1A). Using this system, it was possible to apply the taste substances only to the apical membranes of TBCs (Furue and Yoshii, 1997; Ohtubo et al., 2001) and to keep the intracellular small molecules, such as second messengers, inside the TBCs. Our results indicate that taste receptor cells generate oscillating depolarization with action potentials in response to sweet, bitter, umami, and salty taste substances. The advantages of oscillating depolarization for taste signal transduction are discussed.

All experimental protocols were conducted in compliance with the Guiding Principles for the Care and Use of Animals in the Field of Physiological Sciences approved by the Council of the Physiological Society of Japan, the Animal Institutional Review Board of Kyushu Institute of Technology, and the guidelines of the U.S. National Institutes of Health. The protocols were approved by the president of Kyushu Institute of Technology.

Of the 88 TBCs, 21 responded to one taste substance among the five basic taste substances investigated (Table 1). When the irrigating solution on the apical membrane was changed from deionized water to a sweet (i.e., saccharin-containing) solution, the membrane potential of the TBCs was depolarized with or without action potentials and then repolarized to the resting level despite the presence of saccharin (Figure 1B). This change was repeatedly observed during sweet stimulation of the apical membrane. Such oscillatory changes in membrane potentials are defined as “oscillation” in this study, although the magnitude of the oscillation varied among different tastants.

Saccharin (10 mM) induced oscillating depolarizations with action potentials (Figure 1B). The rising phase of the respective waves comprised no or 1–2 action potentials. The firing frequency increased qualitatively with increasing concentrations of saccharin. Further, the peak magnitude of oscillations was increased from approximately −30 to −10 mV with the increasing concentration (Figure 1B). These oscillating changes in response to saccharin were obtained from 2 of 75 TBCs examined. Similar oscillating receptor potential changes were observed in response to the bitter taste substances (Figure 1C, 10 mM denatonium, 1 of 67 TBCs) and umami taste substance (Figure 1D, 500 mM MSG, 2 of 55 TBCs). Because G protein-coupled taste receptors for sweet, bitter, and umami taste substances are expressed in type II cells (Adler et al., 2000; Chandrashekar et al., 2000; Nelson et al., 2001; Nelson et al., 2002), our results suggest that the type II taste receptor cells generate the oscillating receptor potentials with action potentials.

Interestingly, TBCs responding to 1 M NaCl exhibited two types of receptor potentials, oscillation and depolarization (Figures 2A,B; Table 1). The magnitude and frequency of oscillation were increased with increasing NaCl concentrations (Figure 2A). Oscillating changes responsive to 1 M NaCl were obtained from 2 of 37 TBCs examined. In contrast to the oscillating depolarization, 9 of the 37 TBCs showed membrane depolarization with a phasic component and a subsequent tonic component (Figure 2B). The membrane potential of TBCs started to depolarize when the taste substance reached the apical membrane, and it continuously depolarized during the presence of the stimuli (and did not repolarize while the stimuli were present); repolarization to the resting membrane potential was observed when the tastant was washed out. In this study, “depolarization” refers to the membrane potentials maintaining depolarization in the presence of the taste stimuli. The magnitude of depolarization with the tonic component in TBCs responding to 1 M NaCl was increased to Δ10.5 ± 5.7 mV (n = 9) from the resting membrane potential of approximately −60 mV.

A sour taste substance (HCl) induced depolarization with action potentials (Figure 2C, 5 of 27 TBCs). The magnitude of depolarization in TBCs responding to 10 mM HCl was Δ19.4 ± 9.5 mV (n = 4). Although both NaCl and HCl taste stimuli generate action potentials at the rising phase of depolarization, the after-hyperpolarization was observed in the HCl-responding TBCs, but not in the NaCl-responding TBCs (arrowheads in Figures 2B,C). Previous studies reported that the action potentials in type III cells displayed after-hyperpolarization in response to current-injection–induced membrane depolarization, whereas those in type II cells did not (Kimura et al., 2014; Ohtubo, 2021). Therefore, the cell types may differ between NaCl- and HCl-responding TBCs.

As the kinetics differ between type II and type III cells (Kimura et al., 2014; Iwamoto et al., 2020; Takeuchi et al., 2021), we compared the activation kinetics of the outward rectifying currents that were recorded before the application of the taste substance, to identify the cell types of HCl-responding TBCs. Type II cells mainly expressed Cs⁺-insensitive and slowly activated outwardly rectifying currents that passed through voltage-gated hemichannels and calcium homeostasis modulator 1 and 3 (CALHM1/3) channels and generated tail currents during repolarization. In contrast, Type III cells expressed Cs⁺-sensitive and rapidly activated K⁺ currents but did not generate tail currents. Type I cells expressed relatively small Cs⁺-insensitive outwardly rectifying, small tailed, and relatively small voltage-gated Na⁺ currents. The cell types of the HCl-responding TBCs were identified based on the electrophysiological features of the voltage-gated currents.

The activation kinetics of voltage-activated outward currents at +50 mV obtained from TBCs responding to HCl was 0.63 ± 0.07 ms (n = 5), which was significantly faster than that obtained from oscillating TBCs (4.80 ± 0.89 ms, n = 7, p < 0.0001, Figure 2D) and NaCl-depolarizing TBCs (4.13 ± 1.56 ms, n = 9, p = 0.0002). The activation kinetics of TBCs responding to HCl was similar to that obtained from immunohistochemically identified type III cells (0.86 ± 0.41 ms, n = 6) (Takeuchi et al., 2021). These data suggest that HCl-responding TBCs are type III cells, whereas NaCl-depolarizing TBCs are not type III cells. As type II cells generate relatively larger voltage-gated Na⁺ and outwardly rectifying currents (Figure 2D) than type I cells (Kimura et al., 2014; Takeuchi et al., 2021), the NaCl-depolarizing TBCs are likely to be type II cells.

The frequency analysis showed that the peak of the oscillations was 0.67–1.43 Hz (n = 3) in response to 30 mM saccharin (peak value = 1.43 Hz), 10 mM denatonium (0.95 Hz), and 1 M NaCl (0.67 Hz; Figure 2E). This oscillation at approximately 1 Hz frequency is suitable for the effective recovery from the inactivation of the voltage-gated Na⁺ channels because the slow time constant of the recovery was approximately 0.8 s at −70 mV in type II taste receptor cells of mice (Ohtubo, 2021). Therefore, type II taste receptor cells can continuously generate action potentials in response to taste substances (Figures 1, 2).

The magnitude of oscillation was Δ39–53 mV during the changes (n = 3) in response to 30 mM saccharin (upper value, −12 mV; lower value, −65 mV; Δ53 mV), 10 mM denatonium (−23 mV; −69 mV; Δ46 mV), and 1 M NaCl (−22 mV; −61 mV; Δ39 mV). Additionally, when applied to the basolateral membrane, tetrodotoxin (TTX) inhibited the generation of action potentials but did not affect the oscillations (Figure 2F). The magnitude of oscillation was similar in the presence and absence of TTX (Figure 2F), suggesting that TTX-sensitive Na⁺ channels are not involved in the generation of oscillating depolarization.

In the present study, we demonstrated that taste receptor cells generate oscillating receptor potentials with action potentials in response to sweet, bitter, umami, and salty taste substances using “in situ perforated whole-cell patch clamping.” A previous study using a similar method and mouse soft palate taste buds showed that the restricted application of 10 mM quinine hydrochloride (a bitter substance) to the apical membrane elicited oscillatory receptor potentials with action potentials on their upstrokes (Noguchi, 2003). Although our sample size was only seven cells that elicited oscillating receptor potentials, our results reconfirmed that a bitter substance can induce oscillation and newly demonstrated that sweet, umami, and salty taste substances also induce oscillating receptor potentials. Additionally, we showed that the frequency and magnitude of oscillations increased with increasing taste concentrations. These oscillating membrane potential changes that occur in a concentration-dependent manner may affect the secreting patterns of neurotransmitters, such as ATP, because ATP is a transmitter that conducts signal transduction between taste receptor cells and taste nerves (Finger et al., 2005).

Taste signal conduction from type II cells to taste nerves occurs via both action potential-dependent and action potential-independent ATP secretion through CALHM1/3 channels (Huang and Roper, 2010; Murata et al., 2010). CALHM1/3 channels are nonselective, voltage- and/or Ca²⁺-sensitive large pore-forming channels (Taruno et al., 2013; Ma et al., 2018) that are activated by the membrane depolarization of > −30 mV (Iwamoto et al., 2020). Previous studies have shown that these channels can open without deactivation during a long depolarization period under voltage-clamp conditions (Kimura et al., 2014; Takeuchi et al., 2021). Therefore, the depolarizing oscillation of > −30 mV may itself activate CALHM1/3 channels, which could be possible in case of action potential-independent ATP secretion. Additionally, the prolonged opening of CALHM1/3 channels may disrupt cellular homeostasis because ions and small molecules, such as ATP, move through CALHM1/3 channels according to electrochemical gradients. Thus, oscillating receptor potentials are suitable for the maintenance of physiological homeostasis of type II taste receptor cells by limiting the opening of ATP-permeable channels.

Because of the slow recovery from the inactivation of voltage-gated Na⁺ channels in mouse TBCs, TBCs hardly generate high-frequency firings in response to membrane depolarization induced by current injection (Ohtubo, 2021). Meanwhile, TBCs continuously generate action currents (action potentials) in response to sweet, bitter, and umami taste substances during taste stimuli (Furue and Yoshii, 1997; Noguchi et al., 2003; Yoshida et al., 2006). This discrepancy can be explained by considering membrane potential oscillations. During these oscillations, recovery from the inactivation of voltage-gated Na⁺ channels accelerated by the repolarization of membrane potentials enables repetitive firing during taste stimuli in the taste receptor cells (Figures 1, 2). This repetitive firing may contribute to the action potential-dependent ATP secretion.

Analysis of the activation kinetics of outward rectifying currents suggested that HCl-responding cells were type III cells, which is in agreement with the results of the previous studies that type III cells sense sour taste substances (Huang et al., 2008; Kataoka et al., 2008). Additionally, our data suggested that NaCl-depolarizing TBCs were type II cells because type II cells generated relatively larger voltage-gated Na⁺ and outwardly rectifying currents than type I cells (Kimura et al., 2014; Iwamoto et al., 2020). Similar results were observed in a recent study on sodium taste. According to that study, CALHM1/3 expressing taste cells can generate a depolarization driving action potential by activating the amiloride-sensitive epithelial sodium channel (Nomura et al., 2020). Furthermore, a high salt concentration recruited two aversive taste pathways that activate the bitter- and sour-taste-sensing cells (Oka et al., 2013). Bitter-sensing type II cells may generate oscillating depolarization in response to 1 M NaCl as bitter substance (10 mM denatonium) generated oscillation (Figure 1C). Further studies are needed to clarify the relationships between salty taste transduction mechanisms, including two aversive taste pathways and the two types of receptor potential changes, oscillation and depolarization.

Overall, type II taste receptor cells generate oscillating depolarization in response to sweet, bitter, and umami tastants that activate G protein-coupled receptors. Additionally, the frequency and magnitude of oscillations were increased in a concentration-dependent manner. These results suggest that the information on taste “thickness” can be converted into both firing frequency and magnitude of oscillations. Further studies are needed to elucidate the relationship between the ATP-secretion patterns and receptor potential changes, besides elucidation of molecular mechanisms that generate the oscillations.