Cell Physiology

Tastants evoke cAMP signal in taste buds that is independent of calcium signaling

Kristina R. Trubey, Schartess Culpepper, Yutaka Maruyama, Sue C. Kinnamon, Nirupa Chaudhari


We previously showed that rat taste buds express several adenylyl cyclases (ACs) of which only AC8 is known to be stimulated by Ca2+. Here we demonstrate by direct measurements of cAMP levels that AC activity in taste buds is stimulated by treatments that elevate intracellular Ca2+. Specifically, 5 μM thapsigargin or 3 μM A-23187 (calcium ionophore), both of which increase intracellular Ca2+ concentration ([Ca2+]i), lead to a significant elevation of cAMP levels. This calcium stimulation of AC activity requires extracellular Ca2+, suggesting that it is dependent on Ca2+ entry rather than release from stores. With immunofluorescence microscopy, we show that the calcium-stimulated AC8 is principally expressed in taste cells that also express phospholipase Cβ2 (i.e., cells that elevate [Ca2+]i in response to sweet, bitter, or umami stimuli). Taste transduction for sucrose is known to result in an elevation of both cAMP and calcium in taste buds. Thus we tested whether the cAMP increase in response to sucrose is a downstream consequence of calcium elevation. Even under conditions of depletion of stored and extracellular calcium, the cAMP response to sucrose stimulation persists in taste cells. The cAMP signal in response to monosodium glutamate stimulation is similarly unperturbed by calcium depletion. Our results suggest that tastant-evoked cAMP signals are not simply a secondary consequence of calcium modulation. Instead, cAMP and released Ca2+ may represent independent second messenger signals downstream of taste receptors.

  • calcium-sensitive adenylyl cyclase
  • capacitative entry
  • cross talk
  • taste transduction

during the past decade, numerous advances have been made in our understanding of taste transduction mechanisms. Most tastants of the sweet, bitter, and umami classes are thought to activate G protein-coupled taste receptors and their heterotrimeric G proteins. The Gγ13 subunit appears to be taste specific and associated with Gβ1 (14). This Gβ1γ13 dimer has been shown directly to activate a phospholipase C (PLCβ2), stimulating the production of inositol 1,4,5-trisphosphate (IP3) and eventually triggering an increase in cytoplasmic Ca2+ (14, 23, 35, 38). Strong support for this sequence of signaling events is derived from the pronounced taste deficit that results from genetic ablation of PLCβ2 (45). Although release of stored Ca2+ (triggered by PLCβ2 activity) is essential, taste-evoked calcium transients in taste cells may also include a component of capacitative entry from the extracellular medium (28).

Despite the recent emphasis on this IP3-mediated calcium release pathway, there is evidence that changes in cAMP and cGMP concentration occur after tastant stimulation. Cyclic nucleotide modulation has been demonstrated after stimulation with some sweet, bitter, and umami stimuli (1, 39, 44), and exogenously applied cAMP appears to mimic tastant-evoked activity (8). Nevertheless, the significance and source of cAMP and cGMP in taste transduction are unclear, given the essential role of IP3-mediated signaling.

Cells can modulate cytoplasmic cAMP levels by regulating the function of cAMP-synthesizing adenylyl cyclases (ACs), cAMP-hydrolyzing phosphodiesterases (PDEs), or both. The membrane ACs are integral membrane proteins that comprise a family of nine isoforms, each with distinct regulatory properties. G protein-coupled receptors regulate ACs through G protein subunits such as the stimulatory Gαs, the inhibitory Gαi, or Gβγ dimers that exhibit heterogeneous functional interactions (12, 30). Various AC isoforms are also responsive to modulators such as calcium and calmodulin. Of these, our previous studies (1) demonstrated the presence of mRNA and protein for AC4, AC5/6, and AC8 in rat taste buds. AC5/6 and AC8 are all sensitive to calcium concentrations, based on in vitro assays; AC5 and 6 are inhibited (16), whereas AC8 is stimulated, by calcium (5). The presence of calcium-sensitive ACs in taste cells suggests that tastant-evoked cAMP synthesis might simply be a secondary consequence of the elevation of cytoplasmic calcium. Such interaction between the two pathways has been reported in neural tissue, in neurally derived pheochromocytoma PC12 cells, and in certain endocrine cells (4). In many neuronal cell types, AC activity is stimulated by physiological concentrations of Ca2+ on activation of the PLC pathway or activation of voltage-gated calcium channels (7, 19).

Here we demonstrate the functional presence of a calcium-stimulated AC in rat circumvallate (CV) taste buds. As with other excitable cell types, the calcium stimulation is dependent on entry from extracellular space rather than release from intracellular stores. We show that AC8 is localized in the same cells as PLCβ2 (a key enzyme for taste transduction) and investigate its role in sweet and glutamate taste. However, we demonstrate that although AC activity is enhanced by Ca2+ influx into taste cells, the cAMP response to tastants [sucrose and monosodium glutamate (MSG)] persists even in the absence of extracellular Ca2+. Our results suggest that the cAMP signal for these tastants is a direct outcome of receptor activation, rather than simply a downstream consequence of the elevation of cytoplasmic Ca2+ levels following activation of PLCβ2.


Animals and tissues.

All experiments were carried out according to National Institutes of Health guidelines; protocols were approved by the University of Miami Animal Care and Use Committee. Adult male Sprague-Dawley rats, 6–8 wk old, were purchased from Charles River Laboratories (Wilmington, MA); adult male C57BL/6 mice were obtained from Jackson Laboratories (Bar Harbor, ME). Rodents were killed by exposure to CO2 and decapitated, and tongues were removed. A mixture of 1 mg/ml collagenase D, 2.5 mg/ml dispase II, and 1 mg/ml trypsin inhibitor was injected under the epithelium (10). After 30–45 min of incubation, the epithelium was peeled and trimmed to separate taste bud-enriched areas of the CV trench from surrounding nonsensory epithelium (39).

Reagents and solutions.

Formulations for physiological saline solutions were as follows: Tyrode buffer (mM: 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, 10 glucose, 10 sodium pyruvate, pH 7.4); Ca2+- and Mg2+-free (CMF) Tyrode buffer (mM: 140 NaCl, 5 KCl, 2 EGTA, 10 HEPES, 10 glucose, 10 sodium pyruvate, pH 7.4); phosphate-buffered saline (PBS; mM: 154 NaCl, 1 KH2PO4, 3 Na2HPO4, pH 7.4). The following reagents were purchased from Sigma (St. Louis, MO) and were dissolved in dimethyl sulfoxide: forskolin (Fsk) at 20 mM; 3-isobutyl-1-methylxanthine (IBMX) at 500 mM; cycloheximide at 100 mg/ml; thapsigargin and calcimycin (A-23187) at 1 mM. Aliquots of each were stored at −20°C and were freshly diluted into Tyrode buffer for each experiment. Fsk, IBMX, thapsigargin, and A-23187 were used at concentrations within twofold of published EC50 values. Taste stimuli were diluted into Tyrode buffer at concentrations that are taste effective and shown previously to elicit taste-specific cAMP signals (1, 39).

cAMP measurements.

Rat CV or nontaste epithelial sheets were cut in the midline to yield two equal halves, which served as paired control and treated samples. Pairs of epithelial sheets were subjected to stimulation paradigms as shown in Figs. 1 and 3 and described below. The supernatant solutions were then removed and replaced with 16% HClO4 in Tyrode buffer, and tissues were extracted by vortexing and freezing at −80°C. Subsequent steps for neutralization and clarification of the extract were according to Krizhanovsky et al. (18), and cAMP was quantified with an enzyme immunoassay kit (Amersham Biosciences, Piscataway, NJ) as reported previously (1). To measure protein levels in tissues, we first hydrolyzed them in 5 N NaOH (100 μl for CV samples; 500 μl for nontaste samples) at 60°C for 3 h. Aliquots (10 μl) of the hydrolysates were neutralized with 190 μl of 1 M Tris·HCl (pH 7.0) and assayed with a Nano-Orange Protein Quantitation Kit (Molecular Probes, Eugene, OR). Paired two-tailed t-test was performed with Prism (version 4.00, GraphPad Software, San Diego CA).

Fig. 1.

Entry of Ca2+ from extracellular space stimulates adenylyl cyclase (AC) activity in circumvallate (CV) taste buds. Paired halves of CV epithelium were treated as shown (dark gray or black bars below each graph). Stimuli were in Tyrode buffer (light gray) or in CMF-Tyrode buffer (white). Then, cAMP and protein were quantified. A: thapsigargin (Tg) treatment in normal Tyrode buffer caused a significant elevation of cAMP (P = 0.035; n = 4). Mean values of pmol cAMP/μg protein were 19.8 ± 4.0 for control [forskolin (Fsk only)] and 30.6 ± 3.4 for Tg. Tg treatment did not change cAMP concentration in nontaste tissue (P = 0.44; n = 4; values were control 3.7 ± 0.4 and Tg 3.0 ± 0.9 pmol cAMP/μg protein). B: Tg treatment in the absence of extracellular Ca2+ did not stimulate AC activity (P = 0.95; n = 8). Paired halves of CV epithelium were preincubated in Ca2+- and Mg2+-free (CMF) Tyrode buffer before stimulations. The mean values were 16.7 ± 4.6 pmol cAMP/μg protein for control and 16.2 ± 4.4 pmol cAMP/μg protein for Tg. For nontaste epithelial samples as well, values were not significantly different (P = 0.74; n = 3; control 2.1 ± 0.3, Tg 2.2 ± 0.4 pmol cAMP/μg protein). We noted the high value of cAMP in some samples, even for unstimulated controls. To ensure that this does not represent the maximum achievable cAMP ceiling, we stimulated some CV samples with forskolin (Fsk) for 6 min, processed them in the same manner, and observed considerably higher levels (mean 209 pmol cAMP/μg protein). C: Ca2+ entry via an ionophore in normal Tyrode solution also stimulated AC activity in taste buds. Mean values of cAMP were significantly elevated in the presence of 3 μM A-23187 (P = 0.019; n = 3; control 10.6 ± 0.6, A-23187 16.4 ± 1.7 pmol cAMP/μg protein). For nontaste epithelium, cAMP levels were not significantly different between control and treated samples (P = 0.38; n = 3; control 4.4 ± 2.0, A-23187 5.0 ± 0.8 pmol cAMP/μg protein). Significant difference between control and treated samples (*P ≤ 0.05) or (**P ≤ 0.01).

Fig. 2.

Calcium-sensitive AC8 is expressed in many taste cells that release Ca2+ in response to tastants. Cryosections of CV papillae were subjected to double-label immunofluorescence with antibodies against AC8 and phospholipase C(PLC)β2. AC8 immunoreactivity (A, E) was seen in 2–6 cells per taste bud section, whereas PLCβ2 immunofluorescence (B, F) was typically seen in 4–10 cells per taste bud section. Overlays at low and high magnification (C, G) demonstrate the extent of coexpression of these genes. A bright field image of the same field (D) is also shown. Immunofluorescence for PLCβ2 appears homogeneously throughout the cytoplasm, whereas AC8 exhibits a mottled appearance (asterisk). Examples of cells stained for only 1 of the 2 antigens (open arrowhead) or for both antigens (solid arrowhead) are indicated. H: control cryosections of CV from PLCβ2-knockout mice show no fluorescence when incubated with the same anti-PLCβ2 and secondary antibodies as used above. A bright field image of the same section is shown in I. J: cryosections of rat CV papilla, stained with anti-AC8, preincubated with antigenic blocking peptide before application, and followed by the same fluorescent secondary antibody as above. The lack of immunofluorescence in H and J indicates the specificity of antibodies against PLCβ2 and AC8, respectively. Scale bars: 50 μm for A–D, H–J; 25 μm for E–G. K: Venn diagram illustrates the extent of coexpression of AC8 and PLCβ2 in rat CV taste buds.

Fig. 3.

cAMP modulation in response to tastants is independent of calcium signaling in taste cells. A: incubating taste tissue in CMF Tyrode buffer for 20 min effectively depletes Ca2+ stores. Ca2+ responses of individual taste cells to 20 μM cycloheximide (focally applied as indicated by arrowheads) were recorded in slices of CV papillae in normal Tyrode buffer (left). The slice was then bathed in CMF Tyrode buffer for 20 min. On restimulation with cycloheximide, no elevation of cytoplasmic calcium is apparent (middle). Finally, the slice was again bathed in regular Tyrode buffer for 30 min. Taste cells recovered the ability to elevate cytoplasmic calcium when stimulated with cycloheximide (right). B–D. paired halves of CV epithelium were treated with 0.3 mM 3-isobutyl-1-methylxanthine (IBMX) with or without added tastant [500 mM sucrose or 20 mM monosodium glutamate (MSG)] as shown in the schematics (right), after which cAMP and protein were quantified. B: in normal Tyrode solutions, sucrose stimulation resulted in a significant elevation of cAMP (P = 0.006; n = 5). The mean values were control 6.3 ± 1.5 and sucrose 15.6 ± 3.9 pmol cAMP/μg protein. MSG stimulation resulted in a significant decrease in cAMP (P = 0.013; n = 5). The mean value for each set was control 9.5 ± 1.8 and MSG 6.4 ± 1.0 pmol cAMP/μg protein. C: in CV epithelium, preincubated in CMF Tyrode buffer for 15–20 min to deplete intracellular Ca2+ stores, sucrose and MSG continue to elicit changes in cAMP concentration. Sucrose stimulation resulted in a significant increase of cAMP (P = 0.001; n = 4), with mean values for control being 6.7 ± 1.9 pmol cAMP/μg protein and for sucrose 14.7 ± 3.7 pmol cAMP/μg protein. Under these conditions, MSG stimulation led to a significant decrease in cAMP level (P = 0.001; n = 5). The mean values for control samples were 21.8 ± 5.4 and for MSG samples 13.8 ± 3.4 pmol cAMP/μg protein. D: tastant-elicited cAMP signals persisted in taste buds preincubated with 5 μM U-73122, which blocks PLC-mediated Ca2+ release. Specifically, sucrose gave rise to a significant elevation of cAMP (P = 0.003; n = 3). The mean values were control 4.8 ± 0.7 and sucrose 9.7 ± 1.5 pmol cAMP/μg protein. MSG stimulation resulted in a significant decrease in cAMP (P = 0.0095; n = 3). The mean value for each set was control 8.8 ± 0.6 and MSG 6.6 ± 0.3 pmol cAMP/μg protein. Nontaste lingual epithelium showed no significant change in cAMP concentration when it was stimulated with tastants, in the presence or absence of extracellular Ca2+. With the same paradigms as in B, values for control and sucrose-stimulated samples were 1.3 ± 0.7 and 1.9 ± 1.1 pmol cAMP/μg protein, respectively (P = 0.31; n = 5). Similarly, control and MSG-stimulated samples contained 0.9 ± 0.3 and 0.7 ± 0.2 pmol cAMP/μg protein, respectively (P = 0.39; n = 5). Depleting extracellular Ca2+ as in C had no impact on cAMP in nontaste epithelial samples: control and sucrose-stimulated values were 2.3 ± 1.2 and 2.7 ± 1.5 pmol cAMP/μg protein, respectively (P = 0.60; n = 4), and control and MSG-stimulated samples yielded 2.9 ± 0.7 and 2.2 ± 0.2 pmol cAMP/μg protein, respectively (P = 0.32; n = 4).


CV and foliate papillae from rats or mice were fixed in 4% paraformaldehyde, cryoprotected in 30% sucrose, and cryosectioned (25 μm). Sections were blocked in 7% donkey serum, 0.025% Triton X-100 in PBS for 1 h. Sections were incubated overnight in polyclonal anti-AC8 (1:300 in blocking solution) and/or polyclonal anti-PLCβ2 (1:4,000), both from Santa Cruz Biotechnology (Santa Cruz, CA). Secondary antibodies (Molecular Probes) used for detection were donkey anti-goat IgG-Alexa 488 (1:1,000) for AC8 and donkey anti-rabbit IgG-Alexa 594 (1:1,000) for PLCβ2. Fluorescence images were captured on an Olympus Fluoview scanning confocal microscope. We previously confirmed (1) by immunoblot that the anti-AC8 antibody reacts against an antigen of the appropriate size in brain membrane extracts. The specificity for the AC8 antibody was further confirmed by preincubation with the corresponding blocking peptide (Santa Cruz Biotechnology). The specificity of the PLCβ2 antibody was confirmed in parallel by immunostaining sections of taste papillae from PLCβ2-knockout mice; no fluorescence was detected.

Functional imaging.

CV papillae were loaded with calcium green-dextran, cut into 100-μm-thick slices, and functionally imaged for Ca2+ exactly as previously described (33). Imaging was carried out with argon laser excitation (488 nm) and an FITC filter set to scan >3.5-μm-thick optical sections once every 1.1 s. Taste stimuli were focally and transiently applied to the vicinity of the taste pore. The concentration of tastant reaching the taste pore was estimated by monitoring the dilution of a fluorescent indicator dye included in the stimulation pipette. Recordings are displayed as change of fluorescence normalized to initial fluorescence. It should be noted that this measurement yields relative changes in Ca2+ concentration rather than the absolute concentrations of Ca2+ found within cells.


Calcium-stimulated AC activity in taste buds.

Previously, we demonstrated (1) that mRNAs for the membrane-bound AC isoforms, AC4, AC5/6, and AC8, are expressed in taste cells. In vitro and in many neurons, AC8 activity is potentiated by calcium (once the enzyme has been activated physiologically by G proteins or pharmacologically by Fsk). To test for the functional presence of a calcium-stimulated AC in taste cells, we measured cAMP in intact taste buds, using treatments that elevate intracellular Ca2+ levels. Thapsigargin, an inhibitor of the endoplasmic reticulum Ca2+-ATPase, causes the passive emptying of intracellular Ca2+ stores (40). In previous studies (27, 28, 33), thapsigargin transiently elevated cytoplasmic Ca2+ in taste cells, especially during the first minute of exposure. Hence, we examined Fsk-stimulated AC activity in the absence and presence of 5 μM thapsigargin. Taste epithelium from rat CV papillae was pretreated with 10 μM Fsk to raise basal cAMP levels. Half of the epithelium was then stimulated with 5 μM thapsigargin for 1 min while the other half served as a control. As shown in Fig. 1A, cAMP levels increased significantly in the presence of 5 μM thapsigargin (162 ± 17%) compared with paired control (Fsk only) samples. This Ca2+-mediated stimulation was taste specific insofar as nontaste epithelium (devoid of taste buds) did not demonstrate a similar increase in cAMP (Fig. 1A).

Thapsigargin treatment in many cells induces capacitative entry of Ca2+ across the plasma membrane (32). We asked whether the above stimulation of AC activity resulted directly from the release of stored Ca2+ or was dependent on a secondary (capacitative) Ca2+ influx. Paired halves of taste bud-containing CV epithelium were pretreated with 10 μM Fsk in Tyrode solution (containing 2 mM Ca2+) for 1 min. One piece of taste epithelium (control) was then bathed in 10 μM Fsk in CMF Tyrode buffer. The other paired sample was bathed in 10 μM forskolin + 5 μM thapsigargin in CMF Tyrode buffer. Both tissue samples were incubated for one additional minute (see schematic, Fig. 1B). Thapsigargin failed to elevate cAMP levels when it was applied in a calcium-free condition; cAMP levels in treated samples were 99 ± 9% (n = 8) of those in control samples. The result suggests that the thapsigargin-induced stimulation of AC activity depends on Ca2+ entry across the plasma membrane, which would be lacking in the present paradigm. We noted the high animal-to-animal variability in cAMP levels in the short-term calcium-free condition. Taste buds transferred to an extracellular buffer devoid of calcium often exhibit unstable fluctuations of cytoplasmic Ca2+ (Ref. 28; unpublished observations in our laboratories). Perhaps the variability of cAMP relates to such fluctuations, but our data do not directly address this. Nevertheless, CV taste epithelia tested in pairs derived from individual animals show that thapsigargin fails to elevate cellular cAMP in the absence of extracellular calcium.

We also tested the ability of 3 μM A-23187, a calcium ionophore, when applied in Tyrode solution (i.e., with 2 mM extracellular Ca2+), to stimulate AC activity in taste buds (Fig. 1C). The cAMP concentration in A-23187-treated samples, as a percentage of control, was 154 ± 8%, similar to the effect of 5 μM thapsigargin. Parallel stimulations of nontaste epithelial sheets under all three conditions of calcium modulation showed no evidence of enhanced AC activity (see Fig. 1).

Immunolocalization of calcium-stimulated AC8 in taste cells.

The above results demonstrate that taste buds contain a calcium-stimulated AC that is dependent on Ca2+ entry rather than release from intracellular stores. When taste receptor cells are stimulated with many tastants, receptor-activated PLCβ2 causes the release of Ca2+ from intracellular stores (3, 14, 45). In addition, during taste stimulation, there is evidence of Ca2+ entry across the plasma membrane, through capacitative or voltage-gated mechanisms (3, 28, 31). Hence, we examined whether the calcium-sensitive AC of taste cells is expressed in the same cells that undergo calcium dynamics triggered by tastants.

Earlier, we showed (1) that of the two known calcium-stimulated AC isoforms only AC8 is expressed in rodent taste buds. Hence, we carried out immunocytochemistry to determine whether AC8 is found in cells that exhibit Ca2+ mobilization and Ca2+ entry in response to taste stimuli. In taste buds, cells that express the G protein-coupled taste receptors also express the effector enzyme PLCβ2 (6). Using a well-characterized antibody against PLCβ2, we found robust staining in a large subset of taste cells in CV (Fig. 2B) and foliate papillae from both rats and mice. Generally, immunoreactivity was displayed throughout the cytoplasm of labeled cells (Fig. 2F). Immunoreactivity was specific to taste buds and was not present in the lingual epithelium surrounding taste buds. To confirm the specificity of immunoreactivity, we also immunostained sections of CV papillae from PLCβ2-knockout mice (15). As expected, no immunofluorescence was detected in any taste buds (Fig. 2I) or elsewhere in this tissue. Additional controls for immunofluorescence microscopy consisted of omitting the primary antibody and preincubating the primary antibody with its corresponding antigenic peptide before application; each of these controls consistently yielded no staining (not shown).

Immunoreactivity for AC8 was also detected in a subset of taste cells that were broad and had rounded, large nuclei (Fig. 2A). Fluorescence was concentrated in the cytoplasm, whereas nuclei were devoid of staining. We noted that in some AC8-immunopositive cells staining was concentrated in the apical portion of the cell. In most cells, AC8 reactivity was distributed unevenly as patches. Very similar immunostaining patterns were also observed in rat foliate, mouse CV, and mouse foliate papillae. When anti-AC8 was preincubated with antigenic peptide before application on tissue sections, all immunofluorescence was lost (Fig. 2, J and K).

To quantify the coexpression of these two genes, we scored immunopositive cells that were well defined, with visible nuclei. We counted all such cells within taste buds in both crypts of one section of a CV papilla from each of three rats. Out of 146 cells positive for AC8, 119 cells were also strongly positive for PLCβ2 (79 ± 5% across 3 animals). Conversely, of 390 cells positive for PLCβ2, 119 cells were also strongly positive for AC8 (33 ± 8% across 3 animals). We also scored taste buds from one immunolabeling experiment using mouse CV papillae and detected a frequency of coexpression of AC8 and PLCβ2 similar to that in the rat.

Effect of calcium on tastant-evoked cAMP modulation.

Because the majority of AC8-positive cells also express PLCβ2, a key marker of taste transduction, we examined whether there is a relation between calcium-sensitive AC activity and taste signaling. Increases of cAMP have been noted in response to sweet taste stimuli (39) and decreases in response to umami and bitter (1, 44) stimuli. Control stimulations using sweet taste blockers or nonumami analogs previously confirmed that in both sweet and umami the cAMP signals are taste related and not simply responses to osmotic perturbations (1, 39). The mechanism for altering cAMP after sweet or umami has not been explored and was the focus of the following set of experiments. Although bitter tastants also modulate cyclic nucleotides, these signals are very transient (<1 s) (18, 44) and hence are not amenable to the present method of analysis.

When taste buds are stimulated with sweet or umami tastants, there is release of stored Ca2+ (3, 24, 26) and the subsequent capacitative entry of Ca2+ (28). This suggests two alternative sources for the cAMP signals reportedly evoked by tastants. First, G proteins might directly activate or inhibit ACs or PDEs (Gαs, Gαi or Gαgustducin, respectively). Alternatively, calcium (released or entering capacitatively) might secondarily modulate a calcium-sensitive AC.

To distinguish between these alternatives, we carried out sucrose stimulation under normal physiological conditions, as well as under conditions of depleted Ca2+. We first identified conditions under which stores of cytoplasmic Ca2+ are depleted. Using a well-established paradigm, we imaged the calcium response of CV taste cells to focal application of the bitter tastant cycloheximide. As expected, a pronounced elevation of cytoplasmic Ca2+ was observed on application of 100 μM cycloheximide (Fig. 3A, left). The CV slice was then bathed in CMF Tyrode buffer for 20 min and was restimulated with 100 μM cycloheximide. The absence of a detectable Ca2+ response (Fig. 3A, middle) suggested that cytoplasmic Ca2+ stores had been depleted. Replenishment of stores, as evidenced by the recovery of a calcium response to cycloheximide, was achieved by bathing the slice in regular Tyrode buffer for 30 min (Fig. 3A, right).

Paired halves of CV epithelium in Tyrode solution were stimulated with either 0.3 mM IBMX + 500 mM sucrose for 6 min or 0.3 mM IBMX alone as a control. A substantial accumulation of cAMP resulted from sucrose stimulation (cAMP concentration for sucrose, as a percentage of control, was 244 ± 28%). These data are consistent with those previously reported (39). We then repeated this stimulation paradigm under conditions established above wherein calcium was lacking both in stores and in the extracellular milieu. Paired CV samples were preincubated in CMF Tyrode solution for 15–20 min and were stimulated with 0.3 mM IBMX with or without 500 mM sucrose in CMF Tyrode solution. Even in the absence of Ca2+ release or Ca2+ entry (Fig. 3C), sucrose-stimulated samples showed an increase in cAMP (cAMP concentration for sucrose, as a percentage of control, was 224 ± 10%).

We also considered that the decrease of cAMP previously reported (1) for umami stimulation could result from a calcium-inhibited tonic AC activity. To investigate this possibility, we carried out MSG stimulations under calcium-replete and calcium-depleted conditions. Paired halves of CV epithelium in Tyrode solution were treated with either 0.3 mM IBMX (control) or 0.3 mM IBMX + 20 mM MSG, both for 6 min at 30°C. The results (Fig. 3B) showed a decrease in cAMP induced by MSG (cAMP concentration for MSG, as a percentage of control, was 71 ± 7%), consistent with our previous observations (1). To assess whether modulation of cAMP is a downstream consequence of calcium signaling after umami stimulation, we repeated the MSG stimulation with calcium depleted from stores and extracellular medium. As shown in Fig. 3C, MSG elicited a decrease in cAMP concentration even under conditions that prevent Ca2+ release or entry (cAMP concentration for MSG, as a percentage of control, was 66 ± 7%).

As a further test of the independence of the taste-evoked cAMP and Ca2+ signals, we carried out taste stimulation in the presence of 5 μM U-73122, an inhibitor of PLCβ (41). Taste epithelium was pretreated with the drug for 15 min before application of IBMX and sucrose. The paradigm was essentially similar to that shown in Fig. 3C except that cells were in normal, not CMF, Tyrode solution. Sucrose-stimulated samples exhibited an increase of cAMP concentration (201 ± 6%; P = 0.003, n = 3) over the unstimulated controls (Fig. 3D). Similarly, MSG-stimulated samples exhibited a pronounced decrease of cAMP concentration (75 ± 2%; P = 0.01, n = 3). In contrast, in calcium imaging experiments, 5 μM U-73122 was almost completely effective at blocking the tastant-evoked calcium transient (not shown).

The cAMP responses to both sucrose and MSG were taste specific. None of the foregoing taste stimulations, either in the presence or in the absence of Ca2+, resulted in any significant alteration of cAMP levels between paired samples of nontaste lingual epithelium (see Fig. 3).


We previously showed (1), using RT-PCR and immunocytochemistry, that AC8, a calcium-stimulated cyclase, is expressed in many rat taste cells. Here we demonstrate the functional presence of a calcium-stimulated AC activity in taste cells. In neurons and other cell types, AC8 activity is highly influenced by Ca2+ entry through voltage-gated or capacitative channels (7, 19). Indeed, the calcium-sensitive AC isoforms have been demonstrated to colocalize in membrane microdomains with nicotinic receptors, voltage-gated Ca2+ channels, or capacitative channels, all of which mediate Ca2+ entry (7, 19, 29). Hence, we considered whether taste stimuli might be the physiological triggers that generate the Ca2+ needed to stimulate AC8 in taste cells. Bitter tastants, via Gβ1γ13 subunits, trigger the synthesis of IP3 and the release of stored Ca2+ (14, 36, 44). This IP3-mediated Ca2+ release also occurs when taste buds are stimulated with sweet or umami compounds (3, 24, 26). Tastant-mediated Ca2+ release (shown for denatonium, a bitter tastant) is thought to be coupled to a concurrent entry of Ca2+ from extracellular space, via capacitative channels (28). PLCβ2 is a marker for taste cells that exhibit Ca2+ release and, potentially, capacitative Ca2+ entry in response to tastants. Here we have shown that AC8 is predominantly expressed in this same population of tastant-responsive cells. Nevertheless, we found that tastant-mediated changes in cAMP levels were not perturbed when we altered the availability of Ca2+ for release or capacitative entry.

There are two possible explanations for this apparent conundrum. First, it is possible that AC8 (which is expressed in only a subset of PLCβ2-positive cells) is present only in nonsweet, nonumami cells. We note that capacitative entry has been demonstrated only for bitter, not sweet or umami, tastants (28). Second, AC8 in taste cells may be localized in cellular microdomains distinct from those for taste receptors. Hence, if taste receptors are apically located and if Ca2+ diffusion is tightly restricted, as it is in most neurons, AC8 may not be subjected to capacitative Ca2+ entry during taste stimulation. However, during thapsigargin treatment, intracellular Ca2+ increases globally across the cell, which would impact ACs sequestered in other microdomains. Other types of receptors found on taste cells, e.g., muscarinic (27), purinergic (2), or cholecystokinin (13) receptors, all of which activate PLC signaling, might be the physiological triggers for AC8 activity.

Our results suggest that the positive or negative cAMP signals that we and others have reported for tastants are not simply a downstream consequence of Ca2+ mobilization after taste stimulation. Instead, we propose that downstream of taste receptors two parallel and independent G protein-triggered pathways are activated. The Gβγ arm stimulates PLCβ2 and results in calcium elevation. The Gα arm may trigger cAMP modulation via the action of Gαs, Gαi, or Gαgustducin, each of which is expressed in taste buds (20, 23, 25). Specifically, Gαs, a subunit that stimulates all ACs, may give rise to increases of cAMP (for sucrose). Decreases of cAMP (for bitter and umami stimuli) may arise from activation of Gαi to inhibit AC or Gαgustducin to activate PDEs.

What are the downstream consequences of cAMP modulation in taste cells? Decreases in cellular cAMP may affect taste transduction by modulating the sensitivity of the PLC pathway. For instance, cAMP-mediated phosphorylation is known to decrease the activity of both PLCβ2 (22) and IP3R3 (11), the IP3 receptor located on taste cell stores (6). Thus we postulate that decreased intracellular cAMP levels following umami stimulation may diminish phosphorylation of PLCβ2 and IP3R3, thereby prolonging the Ca2+ transient. Consistent with this hypothesis, membrane-permeant analogs of cAMP blocked the electrophysiological and Ca2+ responses to umami stimuli in rat fungiform taste cells (21).

In the case of sweet transduction, the central role of cAMP is less clear. In hamsters, membrane-permeant analogs of cAMP and cGMP mimic the trains of action potentials elicited by sweet stimuli (8). Patch clamp recordings in frogs and hamsters indicate that cyclic nucleotides depolarize sweet-responsive taste cells by blocking a K+ current (9), possibly by direct action on the channel (17, 42). These data are difficult to reconcile with the seemingly exclusive role of PLCβ2-mediated calcium release suggested by studies on PLCβ2-knockout mice (45). A solution to this conundrum may lie in the presence of two second messenger pathways triggered by sweeteners, (Gβγ-mediated Ca2+ modulation and Gα-stimulated cAMP modulation). Indeed, convergence of calcium and cyclic nucleotide pathways onto a common target K+ channel has been demonstrated in the hamster (42). We postulate that the relative importance of these two pathways may differ between mice (in which the Gβγ pathway predominates) and rats or hamsters (in which the Gα pathway is prominent). It is important to note that, even in the mouse, the impact of cyclic nucleotide signaling is evidenced by the decrease in behavioral and neural responses to sweet, bitter, and umami stimuli when gustducin is knocked out (34, 37, 43). Further studies using single-cell approaches will be required to resolve the specific roles of these two signaling pathways in sweet transduction.


This work was supported by grants from the National Institute on Deafness and Other Communication Disorders (DC-03013, DC-06021).


We thank Dr. Eugene R. Delay for assistance on statistical analyses.


  • The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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