Articles in PresS. Am J Physiol Cell Physiol (March 1, 2006). doi:10.1152/ajpcell.00303.2005
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Tastants evoke a cAMP signal in taste buds that is independent of calcium signaling
Kristina R. Trubey1 , Schartess Culpepper1 , Yutaka Maruyama 1 , Sue C. Kinnamon 3 and Nirupa
Chaudhari1, 2
1
Dept. of Physiology and Biophysics, and 2 Neuroscience Program, University of Miami Miller
School of Medicine, 1600 NW 10 th Ave, Miami, FL 33136, USA.
3
Dept of Biomedical Sciences, Colorado State University, Fort Collins, CO 80523
Running Title: tastant-evoked cAMP signal in taste buds
Address all correspondence to:
Dr. Nirupa Chaudhari
Dept. of Physiology and Biophysics (R430)
University of Miami Miller School of Medicine
1600 NW 10th Ave., Miami, FL 33136, USA
phone: (305) 243-3427; fax: (305) 243-5931
email: [email protected]
Copyright © 2006 by the American Physiological Society.
2
ABSTRACT
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 :M thapsigargin or 3 :M A23187 (calcium ionophore),
both of which increase [Ca2+ ]i, lead to a significant elevation of cAM P levels. This calciumstimulation of AC activity requires extracellular Ca 2+ , suggesting that it is dependent on Ca2+
entry, rather than release from stores. Using immunofluorescence microscopy, we show that the
calcium-stimulated AC8 is principally expressed in taste cells that also express PLC$2 (i.e. cells
that elevate [Ca 2+ ]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 cAM P 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 MSG
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.
KEYWORDS:
calcium-sensitive adenylyl cyclase, capacitative entry, cross-talk, taste transduction
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INTRODUCTION
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 G(13 subunit
appears to be taste-specific and associated with G$1 (14). This G$1(13 dimer has been shown
directly to activate a phospholipase C (PLC$2), stimulating the production of inositol 1,4,5trisphosphate (IP 3 ), 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 PLC$2 (45). Although release of stored Ca2+ (triggered by
PLC$2 activity) is essential, taste-evoked calcium transients in taste cells may also include a
component of capacitative entry from the extracellular medium (28).
In spite of the recent emphasis on this IP3 -mediated calcium release pathway, there is
evidence that changes in cAMP and cGMP concentration occur following tastant stimulation.
Cyclic nucleotide modulation has been demonstrated following stimulation with some sweet,
bitter and umami stimuli (1, 39, 44). And exogenously applied cAM P appears to mimic tastantevoked activity (8). Nevertheless, the significance and source of cAMP and cGMP in taste
transduction is unclear, given the essential role of IP3 -mediated signaling.
Cells can modulate cytoplasmic cAMP levels by regulating the function of either cAMPsynthesizing adenylyl cyclases (AC) or cAM P-hydrolyzing phosphodiesterases (PDE) or both.
The membrane adenylyl cyclases are integral membrane proteins that comprise a family of nine
isoforms, each with distinct regulatory properties. G protein-coupled receptors regulate ACs
through G protein subunits such as the stimulatory G"s, the inhibitory G"i, or G$( dimers which
exhibit heterogeneous functional interactions (12, 30). Various AC isoforms are also responsive
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to modulators such as calcium and calmodulin. Of these, our previous studies demonstrated the
presence of mRNA and protein for AC4, AC5/6 and AC8 in rat taste buds (1). 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 neural-derived pheochromocytoma PC12 cells and in certain endocrine cells
(4). In many neuronal cell types, AC activity is stimulated by physiologic concentrations of Ca 2+
upon 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 taste buds. As with other excitable cell types, the calcium-stimulation is dependent
on entry from extracellular space, rather than by release from intracellular stores. We show that
AC8 is localized in the same cells as PLC$2 (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
MSG) persists even in the absence of extracellular calcium. 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
PLC$2.
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MATERIALS AND METHODS
Animals and Tissues. All experiments were carried out according to NIH Guidelines; protocols
were approved by the University of Miami Animal Care and Use Committee. Adult male
Sprague- Dawley rats, 6-8 weeks 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 CO 2 , decapitated, and tongues were removed.
A mix of 1mg/ml collagenase D, 2.5mg/ml dispase II, 1mg/ml trypsin inhibitor was injected
under the epithelium (10). After 30-45 min of incubation, the epithelium was peeled and
trimmed to separate tastebud-enriched areas of the circumvallate trench from surrounding
nonsensory epithelium (39).
Reagents and solutions: Formulations for physiological salines were: Tyrode’s buffer (in 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’s buffer (in mM:140 NaCl, 5 KCl, 2 EGTA, 10 Hepes, 10 glucose,
10 sodium pyruvate, pH 7.4); phosphate-buffered saline (PBS, in mM: 154 NaCl, 1 KH 2 PO 4 , 3
Na 2 HPO 4 , pH 7.4). The following reagents were purchased from Sigma Chemical Co. (St. Louis,
MO), and were dissolved in dimethylsulfoxide (DMSO): forskolin (FSK) at 20mM; 3-Isobutyl-1methylxanthine (IBMX) at 500mM; cycloheximide at 100mg/mL; thapsigargin and calcimycin
(A23187) at 1mM. Aliquots of each were stored at -20°C and were freshly diluted into Tyrode
buffer for each experiment. FSK, IBMX, thapsigargin and A23187 were used at concentrations
within 2-fold of published EC50 values. Taste stimuli were diluted into Tyrode’s buffer, at
concentrations that are taste effective, and shown previously to elicit taste-specific cAMP signals
(1, 39).
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cAMP measurements: Rat CV or non-taste 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 Figures 1 and 3 and accompanying text. The
supernatant solutions were then removed, replaced with 16% HClO 4 in Tyrode’s and tissues were
extracted by vortexing and freezing at -80°C. Subsequent steps for neutralization and
clarification of the extract were according to Krizhanovsky (18), and cAM P was quantified using
an enzyme immunoassay kit (Amersham Biosciences, Piscataway, NJ) as reported previously
(1). To measure protein levels in tissues, we first hydrolyzed them in 5N NaOH (100:l for CV
samples; 500:l for non-taste samples) at 60°C for 3h. Aliquots (10:l) of the hydrolysates were
neutralized with 190:l of 1M Tris-HCl (pH 7.0) and assayed using a Nano-Orange Protein
Quantitation Kit (Molecular Probes, Eugene, OR). Paired two-tailed t-test was performed using
Prism (v 4.00), GraphPad Software, San Diego CA.
Immunocytochemistry. Circumvallate 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 in phosphate-buffered saline (PBS) for 1 h.
Sections were incubated overnight in polyclonal anti-AC8 (1:300 in blocking solution) and/or
polyclonal anti-PLC$2 (1:4000) , both from Santa Cruz Biotechnology (Santa Cruz, CA).
Secondary antibodies (Molecular Probes, Eugene, OR) 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 PLC$2.
Fluorescence images were captured on an Olympus Fluoview scanning confocal microscope. We
have previously confirmed by immunoblot that the anti-AC8 antibody reacts against an antigen of
the appropriate size in brain membrane extracts (1). The specificity for the AC8 antibody was
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further confirmed by pre-incubating with the corresponding blocking peptide (Santa Cruz
Biotechnology, Santa Cruz, CA). The specificity of the PLC$2 antibody was confirmed in
parallel by immunostaining sections of taste papillae from PLC$2 knockout mice; no
fluorescence was detected.
Functional Imaging. Circumvallate papillae were loaded with calcium green-dextran (CGD), cut
into 100 :m thick slices and functionally imaged for Ca2+ exactly as previously described (33).
Imaging was carried out using argon laser excitation (488nm) and an FITC filter set to scan ~10
:m-thick optical sections once every 1.1 second. 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
pipet. Recordings are displayed as change of fluorescence normalized to initial fluorescence
()F/F). It should be noted that this measurement yields relative changes in Ca2+ concentration,
rather than the absolute concentrations of Ca2+ found within cells.
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RESULTS
Calcium-stimulated AC activity in taste buds
Previously, we demonstrated 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) (5). 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 Ca 2+ -ATPase, causes the passive
emptying of intracellular calcium stores (40). Hence, we examined forskolin-stimulated AC
activity in the absence and presence of 5 :M thapsigargin. In previous studies, we showed that
in taste cells, thapsigargin transiently elevates cytoplasmic Ca2+ , especially during the first
minute of exposure(27, 28, 33). Taste epithelium from rat circumvallate papillae was pre-treated
with 10 :M forskolin to raise basal cAMP levels. Then, half the epithelium was stimulated with
5 :M thapsigargin for one minute while the other half served as a control. As shown in Fig.1A,
cAMP levels increased significantly in the presence of 5 :M thapsigargin (162 ± 17%) when
compared to paired control (forskolin 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 legend).
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 pre-treated with 10 :M forskolin in Tyrode’s
solution (containing 2mM Ca 2+ ) for one minute. One piece of taste epithelium (control) was then
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bathed in 10 :M forskolin in CMF Tyrode’s buffer. The other paired sample was bathed in 10
:M forskolin+5 :M thapsigargin in CMF Tyrode’s 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 shortterm calcium-free condition. Taste buds transferred to an extracellular buffer devoid of calcium
often exhibit unstable fluctuations of cytoplasmic Ca2+ (28; unpublished observations in our
laboratories). Perhaps the variability of cAMP relates to such fluctuations, but our data do not
directly address this. Nevertheless, circumvallate 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 :M A23187, a calcium ionophore, when applied in
Tyrode’s solution (i.e. with 2mM extracellular Ca2+ ), to stimulate AC activity in taste buds
(Fig.1C). The cAMP concentration in A23187-treated samples, as a percentage of control was
154 ± 8%, similar to the effect of 5 :M thapsigargin. Parallel stimulations of nontaste epithelial
sheets under all three conditions of calcium-modulation showed no evidence of enhanced AC
activity (see legend, 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 upon Ca2+ -entry rather than release from intracellular stores. When taste receptor cells
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are stimulated with many tastants, receptor-activated phospholipase C$2 causes the release of
Ca 2+ 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 that of the two known calcium-stimulated AC isoforms, only AC8 is
expressed in rodent taste buds (1). Hence, we carried out immunocytochemistry to determine if
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, PLC$2 (6). Using a well-characterized antibody against PLC$2, we found robust
staining in a large subset of taste cells in circumvallate (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 circumvallate papillae from PLC$2 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 pre-incubating the primary antibody with its corresponding antigenic peptide prior
to application; each of these controls consistently yielded no staining (not shown).
Immunoreactivity for AC8 was also detected in a subset of taste cells which were broad
and had rounded large nuclei (Fig2A). 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
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unevenly as patches. Very similar immuno-staining patterns were also observed in rat foliate,
mouse CV and mouse foliate papillae. When anti-AC8 was pre-incubated with antigenic peptide
before application on tissue sections, all immunfluorescence was lost (Fig.2J,K).
To quantify the co-expression 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 circumvallate papilla from each of 3 rats. Out of 146 cells positive for AC8,
119 cells were also strongly positive for PLC$2 (79 ± 5% across three animals). Conversely, of
390 cells positive for PLC$2, 119 cells were also strongly positive for AC8 (33 ± 8% across three
animals). We also scored taste buds from one immunolabeling experiment using mouse CV
papillae and detected a similar frequency of co-expression of AC8 and PLC$2 as in the rat.
Effect of Calcium on tastant-evoked cAMP modulation
Because the majority of AC8-positive cells also express PLC$2, a key marker of taste
transduction, we examined whether there was 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 sweettaste blockers or non-umami analogs have previously confirmed that in both sweet and umami,
the cAMP signals were taste-related and not simply responses to osmotic perturbations (1, 39).
The mechanism for altering cAMP following sweet or umami have not been explored, and is the
focus of the following set of experiments. Although bitter tastants also modulate cyclic
nucleotides, these signals are very transient (less than 1 sec) (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
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Ca 2+ (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 phosphodiesterases (G"s, G"i or G"gust 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
upon application of 100 :M cycloheximide (Fig.3A, left trace). The CV slice was then bathed in
CMF-Tyrode buffer for 20 min. and was re-stimulated with 100 :M cycloheximide. The absence
of a detectable Ca2+ response (Fig.3A, middle trace) suggested that cytoplasmic calcium 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’s for 30 minutes (Fig. 3A,
right trace).
Paired halves of CV epithelium in Tyrode’s solution were stimulated with either 0.3mM
IBMX + 500mM sucrose for six minutes or 0.3mM 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 same stimulation paradigm under conditions established above
wherein calcium was lacking both in stores and in the extracellular milieu. Paired CV samples
were pre-incubated in CMF Tyrode’s solution for 15-20 min and were stimulated with 0.3mM
IBMX with or without 500mM sucrose in CMF Tyrode’s solution. Even in the absence of
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calcium-release or calcium-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’s solution were treated with either 0.3mM IBMX (control) or
0.3mM IBMX + 20mM MSG, both for 6 min at 30BC. 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 following 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 :M U73122, an inhibitor of PLC$ (41). Taste
epithelium was pre-treated with the drug for 15 min prior to 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’s 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 :M U73122 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
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foregoing taste stimulations, either in the presence or in the absence of Ca 2+ , resulted in any
significant alteration of cAMP levels between paired samples of non-taste lingual epithelium (see
Fig.3 legend).
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DISCUSSION
We previously showed, using RT-PCR and immunocytochemistry, that AC8, a calciumstimulated cyclase, is expressed in many rat taste cells (1). 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 co-localize 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 G$1(13-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). PLC$2 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 PLC$2-positive cells) is present only in non-sweet,
non-umami cells. We note that capacitative entry has been demonstrated only for bitter, not
sweet or umami tastants (28). Secondly, AC8 in taste cells may be localized in cellular
microdomains distinct from those for taste receptors. Hence, if taste receptors are apically located
16
and if Ca2+ -diffusion is tightly restricted, as it is in most neurons, AC8 may not be subjected to
capacitative Ca2+ -entry during taste stimulation. However, during thapsigargin treatment,
intracellular Ca2+ increases globally across the cell, which would impact ACs sequestered in other
microdomains. Other types of receptors found on taste cells, e.g., muscarinic (27), purinergic (2)
or cholecystokinin receptors (13), all of which activate PLC signaling, might be the physiologic
triggers for AC8 activity.
Our results suggest that the positive or negative cAMP signals that we and others have
reported for tastants are not simply a downstream consequence of Ca 2+ -mobilization following
taste stimulation. Instead, we propose that downstream of taste receptors, two parallel and
independent G protein triggered pathways are activated. The G$( arm stimulates PLC$2 and
results in calcium elevation. The G" arm may trigger cAMP modulation via the action of G"s,
G"i or G"gustducin, each of which is expressed in taste buds (20, 23, 25). Specifically, G"s, 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 G"i to inhibit AC, or G"gustducin to activate phosphodiesterases.
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, cAM P-mediated phosphorylation is known to decrease the activity of both PLC$2
(22) and IP 3 R3 (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 PLC$2 and IP 3 R3, 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).
17
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
PLC$2-mediated calcium release suggested by studies on PLC$2-knockout mice (45). A
solution to this conundrum may lie in the presence of two second messenger pathways triggered
by sweeteners, (G$(-mediated Ca2+ modulation and G"-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 (where the G$( pathway predominates) and rats or hamsters
(where the G" 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.
18
ACKNOWLEDGMENTS
We thank Dr. Eugene R. Delay for assistance on statistical analyses. This work was supported by
grants from NIH/NIDCD (DC03013, DC06021).
19
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FIGURE LEGENDS
Figure 1: Entry of Ca2+ from extracellular space stimulates AC activity in CV taste buds.
Paired halves of CV epithelium were treated as shown in the paradigms, and cAMP and protein
were quantified.
A. Thapsigargin treatment in normal Tyrode’s caused a significant elevation of cAMP (p=
0.035; n=4). Mean values of pmol cAMP/:g protein were: 19.8 ± 4.0 for control; 30.6 ± 3.4 for
Tg. Thapsigargin treatment did not change cAMP concentration in non-taste tissue (p= 0.44;
n=4; values were: control, 3.7 ± 0.4 and Tg, 3.0 ± 0.9 pmol cAMP/:g protein).
B. Thapsigargin treatment in the absence of extracellular Ca2+ did not stimulate AC activity
(p=0.95; n=8). Paired halves of CV epithelium were pre-incubated in CMF Tyrode’s before
stimulations. The mean values were 16.7 ± 4.6 pmol cAMP/:g protein for control and 16.2 ±4.4
pmol cAMP/:g protein for Tg. For non-taste epithelial samples as well, values were not
significantly different (p= 0.74; n=3; control, 2.1 ± 0.3 and Tg, 2.2 ± 0.4 pmol cAMP/:g
protein). We noted the high value of cAMP in some sample, even for unstimulated controls. To
ensure that this does not represent the maximum achievable cAM P ceiling, we stimulated some
CV samples with FSK for 6 min, processed them in the same manner, and observed considerably
higher levels (mean 209 pmol cAMP/ug protein).
C. Ca2+ entry via an ionophore in normal Tyrode’s solution also stimulated AC activity in taste
buds. Mean values of cAMP were significantly elevated in the presence of 3 :M A23187 (p=
0.019; n=3; control, 10.6 ± 0.6 pmol cAMP/:g protein; A23187, 16.4 ± 1.7 pmol cAMP/:g
protein). For non-taste epithelium, cAMP levels were not significantly different between control
and treated samples (p=0.38; n=3; control, 4.4 ± 2.0; A23187, 5.0 ± 0.8 pmol cAMP/:g
protein).
26
Figure 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 using
antibodies against AC8 and PLC$2. AC8 immunoreactivity (A, E) was seen in 2-6 cells per taste
bud section whereas PLC$2 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 PLC$2 appears homogeneously throughout the cytoplasm, whereas
AC8 exhibits a mottled appearance (*). Examples of cells stained for only one of the two
antigens ()) or for both antigens (•) are indicated.
H. Control cryosections of CV from PLC$2 knockout mice show no fluorescence when
incubated with the same anti-PLC$2 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, pre-incubated with antigenic blocking
peptide prior to 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 PLC$2
and AC8 respectively. Scale bars: 50 :m for A-D, H-J; 25 :m for E-G.
K. Venn diagram illustrates the extent of co-expression of AC8 and PLC$2 in rat CV taste buds.
Figure 3. cAM P modulation in response to tastants is independent of calcium-signaling in
taste cells.
A. Incubating taste tissue in Ca2+ , Mg2+ -free (CMF) Tyrode’s for 20 min effectively depletes Ca2+
stores. Ca2+ -responses of individual taste cells to 20 :M cycloheximide (focally applied at •)
27
were recorded in slices of CV papillae in normal Tyrode’s (left). The slice was then bathed in
CMF-Tyrode for 20min. Upon re-stimulation with cycloheximide, no elevation of cytoplasmic
calcium is apparent (middle trace). Finally, the slice was again bathed in regular Tyrode’s buffer
for 30 min. Taste cells recovered the ability to elevate cytoplasmic calcium when stimulated with
cycloheximide (right trace).
B-D. Paired halves of CV epithelium were treated with 0.3mM IBMX with or without added
tastant (500mM sucrose or 20mM MSG) as shown in the schematics, after which, cAMP and
protein were quantified.
B. In normal Tyrode’s solutions, sucrose stimulation resulted in a significant elevation of cAMP
(p= 0.006; n=5). The mean values were: control, 6.3 ± 1.5 pmol cAMP/:g protein; sucrose, 15.6
± 3.9 pmol cAMP/:g 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 pmol cAMP/:g protein; MSG,
6.4 ± 1.0 pmol cAMP/:g protein.
C. In CV epithelium, pre-incubated in Ca2+ , Mg2+ -free (CMF) Tyrode’s for 15-20 min to deplete
intracellular calcium 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/:g protein and for sucrose, 14.7 ± 3.7 pmol cAMP/:g
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 pmol cAMP/:g protein
and for MSG samples, 13.8 ± 3.4 pmol cAMP/:g protein.
D. Tastant-elicited cAMP signals persisted in tastebuds pre-incubated with 5:M U73122, 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 pmol cAMP/:g protein;
28
sucrose, 9.7 ± 1.5 pmol cAMP/:g 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 pmol cAMP/:g
protein; MSG, 6.6 ± 0.3 pmol cAMP/:g protein.
Non-taste lingual epithelium showed no significant change in cAMP concentration when it was
stimulated with tastants, in the presence or absence of extracellular Ca2+ . Using the same
paradigms as in Fig. 3B, values for control and sucrose-stimulated samples were 1.3 ± 0.7 and
1.9 ± 1.1 pmol cAMP/:g protein respectively (p= 0.31; n=5). Similarly, control and MSGstimulated samples contained 0.9 ± 0.3 and 0.7± 0.2 pmol cAMP/:g protein respectively (p=
0.39; n=5). Depleting extracellular Ca 2+ as in Fig.3C had no impact on cAMP in non-taste
epithelial samples: control and sucrose-stimulated values were 2.3 ± 1.2 and 2.7 ± 1.5 pmol
cAMP/:g 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/:g protein respectively (p= 0.32; n=4).
pmol cAMP / mg protein
A
B
C
40
40
40
30
30
30
20
20
20
10
10
10
*
0
0
FSK
FSK+Tg
1
0
FSK+Tg
FSK
FSK + A23187
Ca2+ Present
Ca2+ Free
Ca2+ Present
0
FSK
*
FSK
FSK
FSK
Tg
FSK
Tg
FSK
A23187
FSK
2 min
0
1
2 min
Figure 1, Trubey et al.
0
1
2 min
E
A
B
C
D
AC8
*
E
*
F
G
H
I
J
Figure 2, Trubey et al
K
PLCβ2
A
0.1 ΔF/F
in tyrode
20min in CMF tyrode
30min return to tyrode
10sec
Ca
present
C
2+
Ca
free
D
Ca2+
release
blocked
pmol cAMP / mg protein
2+
pmol cAMP / mg protein
B
pmol cAMP / m g protein
tastant
30
tastant
30
**
tastant
*
20
control
IBMX
tyrode
10
10
taste
tastant
IBMX
tyrode
0
0
20
0 2 4
control
30
sucrose
**
control
MSG
**
40
30
20
20
10
0
30
control
IBMX
CMF tyrode
taste
tastant
IBMX
CMF tyrode
10
control
sucrose
**
0
30
-15
control
control
10
10
taste
0
4 6 min
**
20
U73122 U73122+Sucrose
0 2
MSG
20
0
6 min
-15
U73122 U73122+MSG
Figure 3, Trubey et al
IBMX
tyrode+U73122
tastant
IBMX
tyrode+U73122
0
2
4 6 min