Department of Anatomy and Neurobiology, Colorado State University, Fort Collins, Colorado 80523; and Rocky Mountain Taste and Smell Center, University of Colorado Health Sciences Center, Denver, Colorado 80262
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ABSTRACT |
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Ogura, Tatsuya and
Sue C. Kinnamon.
IP3-Independent Release of Ca2+ From
Intracellular Stores: A Novel Mechanism for Transduction of Bitter
Stimuli.
J. Neurophysiol. 82: 2657-2666, 1999.
A variety of substances with different chemical
structures elicits a bitter taste. Several different transduction
mechanisms underlie detection of bitter tastants; however, these have
been described in detail for only a few compounds. In addition, most studies have focused on mammalian taste cells, of which only a small
subset is responsive to any particular bitter compound. In contrast,
~80% of the taste cells in the mudpuppy, Necturus maculosus, are bitter-responsive. In this study, we used
Ca2+ imaging and giga-seal whole cell recording to compare
the transduction of dextromethorphan (DEX), a bitter antitussive, with
transduction of the well-studied bitter compound denatonium. Bath
perfusion of DEX (2.5 mM) increased the intracellular Ca2+
level in most taste cells. The DEX-induced Ca2+ increase
was inhibited by thapsigargin, an inhibitor of Ca2+
transport into intracellular stores, but not by U73122, an inhibitor of
phospholipase C, or by ryanodine, an inhibitor of ryanodine-sensitive Ca2+ stores. Increasing intracellular cAMP levels with a
cell-permeant cAMP analogue and a phosphodiesterase inhibitor enhanced
the DEX-induced Ca2+ increase, which was inhibited
partially by H89, a protein kinase A inhibitor. Electrophysiological
measurements showed that DEX depolarized the membrane potential and
inhibited voltage-gated Na+ and K+ currents in
the presence of GDP--S, a blocker of G-protein activation. DEX also
inhibited voltage-gated Ca2+ channels. We suggest that DEX,
like quinine, depolarizes taste cells by block of voltage-gated K
channels, which are localized to the apical membrane in mudpuppy. In
addition, DEX causes release of Ca2+ from intracellular
stores by a phospholipase C-independent mechanism. We speculate that
the membrane-permeant DEX may enter taste cells and interact directly
with Ca2+ stores. Comparing transduction of DEX with that
of denatonium, both compounds release Ca2+ from
intracellular stores. However, denatonium requires activation of
phospholipase C, and the mechanism results in a hyperpolarization rather than a depolarization of the membrane potential. These data
support the hypothesis that single taste receptor cells can use
multiple mechanisms for transducing the same bitter compound.
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INTRODUCTION |
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Many compounds with diverse structures elicit a
bitter taste in humans (Belitz and Weiser 1985). This
structural diversity implies the presence of multiple transduction
mechanisms and/or multiple receptor proteins in taste receptor cells.
Several mechanisms have been proposed, and even multiple mechanisms
have been reported for single compounds. For example, quinine has been
reported to stimulate inositol-1,4,5-triphosphate
(IP3) production (Spielman et al.
1996
), decrease intracellular cAMP (Ming et al.
1998
) and directly block K+ channels
(Cummings and Kinnamon 1992
), but whether these
mechanisms coexist in the same taste cells has not been determined.
Accordingly, we have undertaken a study using a variety of
methodologies in a single species to examine mechanisms of bitter transduction.
Several investigators have reported bitter transduction mechanisms
involving activation of G proteins. The first involves a
G-protein-mediated activation of phospholipase C (PLC), which results
in increased intracellular levels of IP3. The
IP3 elicits release of Ca2+
from intracellular stores; this is proposed to lead directly to
neurotransmitter release (Akabas et al. 1988;
Hwang et al. 1990
; Ogura et al. 1997a
;
Spielman et al. 1994
). Another mechanism involves a
G-protein-mediated activation of phosphodiesterase (PDE), leading to a
decrease in intracellular levels of cyclic nucleotides
(Ruiz-Avila et al. 1995
; Wong et al.
1996
). The decrease in cyclic nucleotides may depolarize taste
cells by relieving block of a direct cyclic nucleotide-blocked cation
channel (Kolesnikov and Margolskee 1995
). These two
hypotheses are not mutually exclusive. Recent studies suggest that the
bitter compound denatonium elicits both increases in
IP3 and decreases in cAMP in the same tissue (Yan et al. 1999
). However, whether these mechanisms
occur in the same taste cells has not been determined. Activation of
second messengers presumably requires the binding of bitter compounds to G-protein-coupled receptors. Recently, a putative bitter taste receptor protein was cloned (Hoon et al. 1999
), but the
protein has not been characterized functionally.
Several bitter compounds do not require G-protein-coupled receptors for
transduction. In the mudpuppy, Necturus maculosus, quinine
and CaCl2 directly block voltage-gated
K+ channels that are located on the apical
membrane of taste cells (Bigiani and Roper 1981;
Cummings and Kinnamon 1992
; Kinnamon and Roper
1988
); K+-channel block leads to membrane
depolarization and transmitter release. In frog, quinine activates
secretion of intracellularly accumulated Cl
through Cl
pumps on the apical receptive
membrane, resulting in membrane depolarization (Sato et al.
1994
). Recent studies have shown membrane permeant bitter
compounds can increase intracellular cGMP by direct inhibition of PDE
(Rosenzweig et al. 1999
), which may depolarize cells by
opening cyclic nucleotide-gated cation channels (Misaka et al.
1997
). Other receptor-independent events, such as changing the
phase-boundary potential at the outer surface of the membrane, have
been reported in response to lipophilic bitter compounds (Koyama
and Kurihara 1972
). It is not known if these nonspecific effects contribute to bitter transduction in taste cells.
One of the difficulties in revealing general mechanisms for bitter
transduction is that they have resulted from studies using different
bitter compounds, different animal species, and a variety of
techniques. To investigate whether the same bitter compound can use
multiple transduction mechanisms in the same taste cells, it is
reasonable to compare data obtained in a single species and with
similar techniques. The mudpuppy, N. maculosus, is an ideal
model for studying bitter transduction (Bowerman and Kinnamon 1994; Kinnamon 1992
; Ogura et al.
1997a
). First, mudpuppy taste cells are large, easily isolated,
and amenable to both Ca2+ imaging and patch-clamp
recording. In addition, most mudpuppy taste cells are bitter sensitive
(Ogura et al. 1997a
). This situation is in sharp
contrast to mammals, where only a small fraction of taste cells
responds to any particular bitter stimulus (Bernhardt et al.
1996
). Low responsivity in mammalian taste cells makes detailed
pharmacological dissection of transduction extremely difficult.
Our previous studies on bitter transduction in mudpuppy taste cells
have revealed two different transduction mechanisms for bitter
compounds, one relying on release of Ca2+ from
intracellular stores and the other involving direct block of apical
K+ channels. Denatonium, an extremely bitter
compound to humans, increased intracellular Ca2+
levels ([Ca2+]i) by an
IP3-mediated release of
Ca2+ from intracellular stores. The elevation in
[Ca2+]i activated
Ca2+-dependent K+ and
Cl currents, which hyperpolarized the membrane
potential. Intracellular cAMP levels did not significantly modify the
Ca2+ responses (Ogura et al.
1997a
). In contrast, quinine (Kinnamon 1992
) and
CaCl2 (Bigiani and Roper 1991
)
depolarized mudpuppy taste cells by direct block of the apical
K+ conductance. In the present report, we studied
the transduction of dextromethorphan bromide (DEX) in mudpuppy taste
receptor cells by means of using Ca2+ imaging and
whole cell patch recording. DEX, an antitussive, is avoided by pigs in
behavioral tests and causes a strong bitter taste in humans (at 8.1 mM)
(Nelson and Sanregret 1997
). To test whether DEX uses
transduction mechanisms similar to either quinine or denatonium, we
used recording procedures similar to those used previously
(Ogura et al. 1997a
). Some of these results have been published in abstract form (Ogura et al. 1997b
, 1998
).
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METHODS |
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Isolation of taste receptor cells
Mudpuppies (N. maculosus) were obtained from
commercial sources and housed in large aquaria at 10°C. Minnows were
provided weekly as a food source. Taste receptor cells were isolated as described previously (Kinnamon and Roper 1988;
Ogura et al. 1997a
). Briefly, mudpuppies were
decapitated after anesthesia in ice-cold water, and the lingual
epithelium was separated from the underlying connective tissue. The
apical surface of the stripped epithelium was then incubated for 15 min
in fluorescein-conjugated wheat germ agglutinin (Molecular Probes: 0.5 mg/ml in amphibian physiological saline, APS), so that mature taste
cells could be distinguished from other cell types after isolation
(Kinnamon et al. 1988
). The epithelium then was
incubated in APS containing collagenase (1 mg/ml; Sigma, Type1),
albumin (1 mg/ml), and glucose (5 mM) until the epithelium could be
separated gently from the underlying connective tissue, leaving the
taste buds atop their connective tissue papillae. The taste buds were
dissociated in Ca2+-free APS. Isolated taste
cells were removed by gentle suction and plated onto recording chambers
made with cover slips (for Ca2+ imaging) or glass
slides (for whole cell recording), both coated with Cell-Tak
(Collaborative research).
Intracellular calcium measurement
[Ca2+]i in
isolated taste receptor cells was measured using the membrane-permeable
Ca2+ -sensitive dye fura-2 AM as described
previously (Ogura et al. 1997a). Briefly, cells were
loaded with fura-2 AM (2 µM, Molecular Probes) in the presence of a
dispersing reagent, Pluronic F-127 (final <0.02%, Molecular Probes)
for 10 min, then washed with normal APS for 20 min. Images were
acquired with an intensified CCD camera (IC100-ICCD, Paultek Imaging)
through an oil-immersion objective lens (Fluor ×40, 1.3 NA, Nikon) of
an inverted microscope (Diaphot TMD, Nikon). The video signal from the
camera was captured using a frame grabber board (Quick Capture, Data
Translation) on a Macintosh computer (Quadra 800, Apple Computer). For
dual-wavelength ratiometric Ca2+-measurements,
pairs of fluorescent images were recorded at 350- and 380-nm excitation
using a filter wheel (EMPIX Imaging). A 510- to 580-nm band-pass filter
(Chroma Technology) collected emitted light. Intracellular
Ca2+ concentration was calculated in selected
areas from the ratio of 350- and 380-nm images (Grynkiewicz et
al. 1985
), using the public domain NIH Image program (developed
at the National Institutes of Health and available on the Internet at
http://rsb.info.nih.gov/nih-image/). The calcium calibration kit II
(C-3009, Molecular Probes) was used to obtain
Ca2+ calibration curves. Time-course measurements
were obtained by plotting the time course of
[Ca2+]i averaged over
most of the cell area excluding the edges of the cell. Generally, <10
s. was required for total exchange of solutions in the 200-µl chamber
by superfusion.
Cells were bathed in normal APS until the resting intracellular calcium level was stable. The bath then was perfused with APS containing dextromethorphan hydrobromide (DEX, 2.5 mM, Sigma and gift from the Procter and Gamble) or denatonium benzoate (2.5 mM, Sigma). Washing with normal APS followed until the intracellular calcium again reached prestimulus levels. Other treatments included: Ca2+-free APS, ryanodine (100 µM in APS for 5-10 min, Calbiochem), thapsigargin (1 µM in APS for 12-15 min, Sigma), U73122 (5 µM in APS for 5-10 min, Calbiochem), H89 (10 µM in APS for 15-20 min, Calbiochem), and a mixture of isobutyl methoxyxanthine (IBMX, 100 µM, Sigma) and 8-chlorophenylthio-cAMP (8-cpt-cAMP, 1 mM, Sigma) in APS for 3 min.
Each taste cell was considered to respond to DEX if it experienced an increase in [Ca2+]i that was >2 SD above the mean resting level. The effects of drug treatments on the DEX response were assessed using Student's t-tests. Statistical values are presented as mean [Ca2+]i ± SE.
Giga-seal whole cell recording
Membrane currents were measured using giga-seal whole cell
recording (Hamill et al. 1981). Electrodes were made
from microhematocrit capillary tubes (American Scientific Products,
McGaw Park, IL) pulled on a two-stage vertical puller (PB-7,
Narishige). When filled with intracellular saline, electrode resistance
ranged from 5 to 7 M
. Cells were viewed at a magnification of ×400
using a Nikon Diaphot inverted microscope fitted with Hoffman optics. Seals of 1-10 G
were obtained by gentle suction, and entry into the
cell was achieved by delivery of a short depolarizing pulse to the
pipette. Whole cell currents were measured at room temperature using an
Axopatch 200B patch-clamp amplifier (Axon Instruments). Signals were
filtered at 5 kHz and recorded digitally at 100 µs. Data were stored
using a laboratory computer (11/23, Digital Equipment Corporation)
equipped with a Cheshire data interface and Basic 23 software (Indec
Systems). All voltage commands were computer generated. Unless
otherwise noted, leak and linear capacitative currents were subtracted
from all records by computer. Series resistance, which was typically
<10 M
, was not compensated. Gravity-fed stimuli were bath-applied
to the 0.5-ml recording chamber. To prevent loss of the seal and to
prevent perfusion artifacts during whole cell recording, the perfusion
rate was lowered to 2-3 ml/min.
After membrane capacitance was compensated electronically, membrane
currents were recorded in APS in response to depolarizing commands from
a holding potential of 80 mV. The bath then was perfused with DEX
(2.5 mM), and membrane currents were measured again. Finally, DEX was
washed out of the bath with normal APS and the currents measured again.
To record the time course of the DEX effect, only two voltage steps
were used: one for inducing inward current and the other for inducing
outward current. For some recordings, the tip of the pipette was filled
with the normal pipette solution but back-filled with a solution
containing 1 mM GDP-
-S (Sigma) to inhibit activation of G proteins.
Solutions
Normal APS contained (in mM) 112 NaCl, 2 KCl, 8 CaCl2, and 3 HEPES, buffered to pH 7.2 with NaOH. Ca2+-free APS contained either 1 mM BAPTA (for cell isolation) or 1 mM EGTA (for Ca2+ imaging) without CaCl2 in normal APS. Patch pipette solution contained (in mM) 114 KCl, 2 NaCl, 0.09 CaCl2, 2 MgCl2, 1 BAPTA, 1 ATP, 0.4 GTP, and 10 HEPES, buffered to pH 7.2 with KOH. For recording Ba2+ currents, 30 mM NaCl was replaced with 20 mM BaCl2, 10 mM TEA Cl, and 0.001 mM TTX were added in APS, and KCl was replaced by CsCl in patch pipette solution.
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RESULTS |
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DEX increases [Ca2+]i in taste receptor cells
We measured DEX-induced changes in
[Ca2+]i in isolated
mudpuppy taste cells using calcium imaging with the
Ca2+-sensitive fluorescent dye fura-2.
Previously, we showed that >80% of mudpuppy taste cells respond to
denatonium with an increase in
[Ca2+]i (Ogura et
al. 1997a). More than 90% of these denatonium-sensitive taste
cells also responded to DEX with an increase in
[Ca2+]i (67 of 72 cells);
cells that did not respond to denatonium usually did not respond to
DEX. Figure 1 compares the time course of
responses to denatonium and DEX in the same taste cells. Responses to
denatonium reached a peak rapidly and began declining, even in the
continued presence of the stimulus. In contrast, responses to DEX were
slower in most cells (Fig. 1, A-D and G) and did
not decline until the stimulus was removed from the bath (Fig. 1, A, C, and D). A few cells, however, showed
responses to DEX that resembled the kinetics of the denatonium response
(Fig. 1F). An interesting feature of some DEX responses was
a temporal increase in
[Ca2+]i immediately after
the wash-out of DEX; these OFF responses were larger than
the ON response in some cells (Fig. 1G).
OFF responses were not usually observed in response to
denatonium. The peak
[Ca2+]i elicited by 2.5 mM DEX was usually 20-80% above resting
[Ca2+]i (43 ± 5%,
n = 67). Repeated applications of DEX to the same cells
resulted in similar increases in
[Ca2+]i, as long as the
[Ca2+]i was returned to
resting levels after each wash (1st application, 53 ± 11%
increase; 2nd application, 56 ± 13% increase, n = 11, see Fig. 2, control).
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DEX releases Ca2+ from intracellular stores
Intracellular Ca2+ can increase by release
from intracellular stores or by Ca2+ influx. To
determine whether extracellular Ca2+ was
required, we compared the DEX-induced increase in
[Ca2+]i in normal and in
Ca2+-free APS in the same cells. The
DEX-induced increase in
[Ca2+]i persisted even in
Ca2+-free APS. The magnitude of the response,
although slightly smaller, was not significantly different from in
normal APS (Figs. 1B and 2; 0 Ca2+,
paired 1-tailed Student's t-test = 0.62, df = 13, P = 0.27). These data suggest that intracellular stores
are the primary source for the increase in
[Ca2+]i. To investigate
further the role of Ca2+ stores in the response,
we used thapsigargin, a Ca2+-ATPase inhibitor.
Thapsigargin inhibits the reloading of Ca2+
stores, resulting in their gradual depletion of
Ca2+ (Meyer and Stryer 1990;
Thastrup et al. 1990
). Thapsigargin (1 µM) increased
[Ca2+]i to a variable
extent in all taste cells tested. This increase in
[Ca2+]i was slow, as has been shown previously
(Ogura et al. 1997a
). After incubation with thapsigargin
for 10-15 min, which should be sufficient for store depletion, neither
2.5 mM DEX nor 2.5 mM denatonium increased
[Ca2+]i in all cells
tested (Fig. 1C). The effect of thapsigargin on the response
to DEX was statistically significant (paired t- test = 3.37, df = 14, P < 0.003, Fig. 2, thapsigargin).
These data strongly suggest that DEX releases
Ca2+ from intracellular stores. Because
denatonium also releases Ca2+ from intracellular
stores (Akabas et al. 1988
; Hwang et al.
1990
; Ogura et al. 1997a
), we examined whether
the mechanism of Ca2+ release in response to DEX
is the same as the response to denatonium. These results are described
in the following section.
Intracellular signaling pathway for DEX response
Two types of Ca2+ stores have been
identified in many types of cells: one coupled to an
IP3 receptor and the other coupled to a ryanodine
receptor (Sharp et al. 1993; Simpson et al.
1995
). To investigate whether ryanodine receptor-coupled
Ca2+ stores are involved in the DEX response, we
applied DEX in the presence of ryanodine, which inhibits
Ca2+ release from the stores (Sutko et al.
1985
). Ryanodine (100 µM for 5-10 min) had no effect on
resting [Ca2+]i
and did not inhibit the
[Ca2+]i increase in
response to DEX (Figs. 1E and 2; paired
t-test = 0.88, df = 13, P = 0.20).
Thus ryanodine receptor-coupled Ca2+ stores are
not likely to be involved in the response to DEX.
To determine whether activation of PLC is involved in the DEX response,
we used U73122, a PLC inhibitor (Salari et al. 1993; Thompson et al. 1991
). Incubation with U73122 (5 µM
for 10-15 min) completely inhibited the denatonium-induced response,
as shown previously (Ogura et al. 1997a
), but the
response to DEX was unaffected (Figs. 1D and 2; paired
t-test = 0.88, df = 7, P = 0.21).
These results suggest that U73122-sensitive PLC is not involved in the
response to DEX.
To determine if DEX and denatonium release Ca2+ from the same intracellular stores, we applied DEX immediately after stimulation with denatonium without an intervening wash. If DEX and denatonium release Ca2+ from different Ca2+ stores, the two Ca2+ responses should be mutually independent. However, as shown in Fig. 1F, DEX failed to induce a Ca2+ response when applied subsequent to denatonium. These data suggest that DEX and denatonium both release Ca2+ from the same IP3-coupled Ca2+ stores, but the DEX response apparently does not require activation of PLC.
OFF responses to DEX
An OFF response, involving a temporal increase in
[Ca2+]i immediately after
wash out of DEX, was observed in 45% (57 of 126 cells) of the taste
cells that responded to DEX. Interestingly, we seldom observed
OFF responses to denatonium. In some cells, the
OFF response to DEX was larger than the DEX response itself (cf. Fig. 1G). The OFF response persisted in
Ca2+-free saline (n = 10, data not shown), suggesting that the off response involves release of
Ca2+ from intracellular stores.
Electrophysiological OFF responses to taste stimuli have
been described previously (Cummings et al. 1993;
Tsunenari et al. 1996
) but the mechanisms involved are
not known. It is possible that electrophysiological off responses may
result from the transient increases in
[Ca2+]i due to release
from intracellular stores.
Intracellular cAMP levels enhance DEX responses
A possible involvement of cAMP in the response to DEX was examined
by increasing intracellular cyclic nucleotide concentrations because
cAMP levels are modulated by some bitter compounds (Kinnamon and
Margolskee 1996). Incubation with a mixture of IBMX (100 µM, a phosphodiesterase inhibitor) and 8-cpt-cAMP (1 mM, a cell permeant cAMP analogue) did not affect resting
[Ca2+]i (Fig.
3). However, the DEX-induced increase in
[Ca2+]i was enhanced
under these conditions (Fig. 3). In the presence of IBMX and 8-cpt
cAMP, the increase in
[Ca2+]i reached peak
levels faster than under control conditions, and the response began to
decline in the continued presence of DEX. The increase in peak
[Ca2+]i was significant
(paired t-test = 3.3, df = 24, P < 0.002, Figs. 3 and 4, cAMP).
Incubation with either IBMX (100 µM) or 8-cpt-cAMP (1 mM) alone did
not enhance the DEX response (data not shown). The enhanced response
persisted in Ca2+-free bath solution (paired
t-test = 2.2, df = 3, P =0.12, Fig. 4,
cAMP in 0 Ca2+), suggesting that enhanced portion
of the Ca2+ response was due to release of
Ca2+ from intracellular stores. The enhanced
response was blocked partially after incubation in H89 (10 µM, a PKA
inhibitor) in the presence of IBMX and 8-cpt-cAMP. The effect of H89 on
the response to DEX was statistically significant (paired
t-test = 3.6, df = 7, P < 0.005, Figs. 3B and 4, cAMP + H89). There was no effect of H89
itself on the DEX response (paired t-test = 0.37, df = 5, P = 0.36, Fig. 4, H89). These data suggest
that DEX is unlikely to increase cAMP levels directly because
incubation with the mixture of IBMX and 8-cpt-cAMP alone did not mimic
the DEX response. Thus it is likely that cAMP levels (and PKA) are
modulated by other regulatory mechanisms, which subsequently modulate
the response to DEX.
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DEX inhibits voltage-gated currents
We used giga-seal whole cell recording to examine the effect of
DEX on membrane potential and voltage-dependent currents. Cells were
voltage-clamped at a holding potential of 80 mV, and current was
measured in response to voltage step pulses from
50 to +70 mV. A
representative I-V relationship of peak inward current and
steady outward current measured at 17.5 ms is plotted in Fig. 5A. Perfusion of DEX for 1 min
completely blocked the transient inward Na+
current and significantly reduced outward K+
currents (Fig. 5A, right). The average inhibition of outward currents elicited by voltage steps to +40 mV was 47 ± 8.5%
(n = 18). To examine the time course of the response,
peak inward and steady state outward currents were monitored by voltage
steps to
20 and +40 mV, respectively (Fig. 5B). These
electrophysiological responses to DEX reached maximum levels more
rapidly than responses in Ca2+-imaging
experiments. In addition to Na+ and
K+ currents, mudpuppy taste cells express a
prominent, slowly inactivating Ca2+ current that
is usually hidden in the large sustained outward current
(Kinnamon and Roper 1988
). Using
Ba2+ as a charge carrier for
Ca2+ channels, we measured
Ba2+ currents elicited by voltage steps from
30
to +50 mV from a holding potential at
80 mV (Fig.
6A). The average inhibition of
Ba2+ currents was 79 ± 6.1%
(n = 5). To monitor the time course of the response,
peak inward currents were elicited by a voltage step to +10 mV. DEX
significantly reduced the Ba2+ currents. Compared
with DEX-modulated inward and outward currents in normal saline, the
reduction was slower in Ba2+ saline (Fig.
6B).
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|
G proteins are not involved in the response to DEX
To examine whether G proteins are involved in the DEX-induced
reduction of voltage-gated currents, we used a nonhydrolyzable GDP
analogue, GDP--S (1 mM) in the patch pipette solution. The effect of
GDP-
-S on peak inward and outward currents was monitored by voltage
steps to
20 and +40 mV, respectively, from a holding potential of
80 mV (Fig. 7). Previously, we showed
that the response to denatonium was inhibited completely by GDP-
-S
15 min after establishment of the whole cell configuration
(Ogura et al. 1997a
). However, even after 17 min of
whole cell recording, DEX continued to block inward and outward
currents, similar to control responses. There was no significant
difference between the amplitude of responses at 2 and at 17 min
(paired t-test = 2.34, df = 2, P = 0.07). The time course of responses at 2 and 17 min was quite similar,
suggesting that G proteins are not involved in the DEX-induced block of
voltage-gated currents. Thus DEX apparently blocks voltage-gated
channels directly.
|
DEX depolarizes the membrane potential
Previously, we showed that another bitter compound, quinine,
blocks voltage-gated outward currents, causing membrane depolarization in mudpuppy taste cells (Kinnamon and Roper 1988). In
contrast, denatonium increases outward currents and hyperpolarizes the
membrane potential (Ogura et al. 1997a
). To determine
the effect of DEX on membrane potential, we used giga-seal whole cell
recording in current-clamp mode. As expected, DEX depolarized taste
cells (Fig. 8). The time course was
similar to the DEX-induced reduction of outward current. The amplitude
of depolarization was 25-49 mV from the resting potential (resting
potential:
73 ± 3.9 mV, after DEX:
27 ± 6.8 mV,
n = 8).
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DISCUSSION |
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In this study, we used Ca2+ imaging and
giga-seal whole cell recording to examine the transduction mechanism of
DEX in isolated mudpuppy taste cells. The results suggest that DEX
depolarizes mudpuppy taste cells by direct block of an apically located
K+ conductance. In addition, DEX causes a slow
release of Ca2+ from
IP3-sensitive Ca2+ stores
by a PLC-independent mechanism (see Fig.
9 and Table
1). This latter mechanism is novel and
may be due to the lipophilic DEX entering taste cells and interacting
directly with the Ca2+ stores. Because many
bitter compounds are lipophilic, this may represent a general mechanism
for bitter transduction that may be associated with the persistent
bitterness of some compounds. The transduction mechanism for DEX is
distinct from that of denatonium (Ogura et al. 1997a)
but has some similarity to that of quinine (Kinnamon and Roper
1988
) in the same species. These data support the hypothesis
that single taste receptor cells can use multiple mechanisms for
transducing the same bitter compound (Spielman et al.
1992
).
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|
A caveat in the interpretation of these data are that we have no direct
behavioral evidence that DEX is bitter to mudpuppies. A previous
behavioral study indicated that compounds that block the apical
K+ conductance in taste cells are rejected when
presented to mudpuppies in agar cubes (Bowerman and Kinnamon
1994). Given that DEX blocks the same conductance, it is likely
that DEX would be rejected in a similar manner.
Ca2+-imaging data
Whereas denatonium transduction relies of a G-protein-coupled
receptor-PLC-IP3 pathway (Ogura et al.
1997a), the response to DEX persists in the presence of the PLC
inhibitor U73122. Thus the DEX response appears to be independent of
IP3 formation. Yet, both compounds appear to
release Ca2+ from the same intracellular
Ca2+ stores. Release of
Ca2+ from intracellular stores is involved in the
transduction of many bitter compounds, including sucrose octaacetate,
caffeine, strychnine, and denatonium, all of which increase
IP3 levels in the taste tissue of rodents
(Hwang et al. 1990
; Spielman et al. 1994
). Clearly, the transduction of DEX must involve an
IP3-independent mechanism for
Ca2+ release. Although we cannot rule out a
mechanism involving a unique second-messenger pathway, we suggest that
DEX may penetrate the membrane and release Ca2+
directly from intracellular stores. Several bitter compounds are
lipophilic, and previous studies provided evidence for the plausibility
of direct interaction with intracellular targets by quinine
(Cummings and Kinnamon 1992
) and caffeine (Koyama
and Kurihara 1972
; Rosenzweig et al. 1999
). If
DEX uses this mechanism, specific membrane-receptors for DEX would not
be required. Such a mechanism could explain the relatively slow
increase in [Ca2+]i and
the insensitivity to U73122. It is not clear what effect the slow
increase in [Ca2+]i has
on transmitter release in taste cells. Presumably, transmitter is
released initially as a result of the DEX-induced block of the apical
K+ conductance. We hypothesize that the slow
increase in [Ca2+]i that
follows would prolong transmitter release, but further studies will be
required to determine the Ca2+ dependence of
vesicle release. Many lipophilic bitter compounds exhibit a bitter
aftertaste, and a prolonged period of vesicle release may contribute to
this phenomenon.
Effect of cAMP
Our data showed that increased cAMP levels enhance the
[Ca2+]i increase in
response to DEX by PKA phosphorylation. Similar modulatory effects of
cAMP on Ca2+ release have been reported
in hepatocytes and neuroblastoma cells (Bird et al.
1993; Wojcikiewicz and Luo 1998
). The
physiological relevance of the enhanced Ca2+
response in the presence of cAMP in taste cells is unclear. Because cAMP itself had no effect on intracellular Ca2+
levels, it is likely that other physiological processes modulate cAMP
levels and that this modulation affects the DEX response. It has been
shown that serotonin modulates voltage-dependent
Ca2+ channels in mudpuppy taste cells via an
increase in intracellular cAMP levels (Delay et al.
1997
). Further experiments will be required to determine if the
presence of serotonin affects Ca2+ responses in
response to DEX stimulation.
In contrast to the present experiments, previous physiological studies
suggest that a reduction in cAMP levels is associated with
bitter transduction. In frog taste cells, an inward conductance induced
by quinine is suppressed by membrane permeant cAMP analogs (Tsunenari et al. 1996), and the effect is antagonized
by inclusion of transducin-derived peptides in the patch pipette
solution (Kolesnikov and Margolskee 1995
). These data in
frog are considered to support a role for gustducin or transducin in
bitter taste transduction. Quinine is believed to activate gustducin or
transducin, which in turn activates PDE, reducing cAMP levels and
relieving block of a cyclic nucleotide suppressible cation conductance
(Kinnamon and Margolskee 1996
). It is not known,
however, if frog taste cells contain transducin or gustducin. In
contrast, there is no evidence for a role of gustducin or transducin in
mudpuppy taste cells. The denatonium Ca2+
response is unaffected by changes in intracellular cAMP (Ogura et al. 1997a
), and transducin-derived peptides have no effect on membrane conductance (Kinnamon, unpublished data).
Several studies suggest a role for gustducin (and/or transducin) in
bitter taste transduction in mammalian taste cells. Both G proteins are
present in taste cells, and gustducin knockout mice are less sensitive
to bitter compounds than control mice (Wong et al.
1996). These data are supported by biochemical measurements showing that several bitter compounds, including quinine and
denatonium, activate gustducin and transducin in the presence of taste
cell membranes (Ming et al. 1998
; Ruiz-Avila et
al. 1995
). These same compounds also activate
IP3 production in taste cells (Miwa et al.
1997
; Spielman et al. 1996
). The relative
importance of these second messenger systems in bitter taste has not
been examined.
OFF responses
We often observed OFF responses to DEX. Similar
OFF responses to taste stimuli have been observed
previously in electrophysiological recordings from taste receptor cells
(Cummings et al. 1993; Tsunenari et al.
1996
). It is possible that relief from a suppressive mechanism may be involved in the off response. We often recorded a decrease in
[Ca2+]i in response to
denatonium or DEX after inhibition of Ca2+
release with thapsigargin. Similar suppressive effects are proposed as
a mechanism for off responses in olfactory receptor neurons (Kurahashi et al. 1994
). Further studies on the
suppressive mechanism in taste receptor cells may reveal a role for off
responses in taste adaptation and taste coding mechanisms.
Electrophysiological data
Our electrophysiological data showed that DEX depolarized mudpuppy
taste cells by blocking the voltage-dependent K+
conductance, which is also a resting conductance in these cells (Cummings and Kinnamon 1992). Similar results were
obtained with quinine (Cummings and Kinnamon 1992
;
Kinnamon and Roper 1988
) and CaCl2
(Bigiani and Roper 1991
). In contrast, denatonium
hyperpolarized the membrane, due to activation of
Ca2+-dependent K+ channels
(Cummings and Kinnamon 1992
) and
Cl
channels (Taylor and Roper
1994
) due to the Ca2+ released from
intracellular stores (Ogura et al. 1997a
).
In brain, DEX has been shown to bind to Sigma receptors, which are
G-protein-linked receptors that affect a variety of targets, including
a resting K+ conductance (Su
1991). We do not believe that DEX is blocking K+ channels in taste cells by activating Sigma
receptors, however, because the DEX-induced block of
K+ channels in taste cells was unaffected by the
G protein inhibitor GDP-
-S.
DEX also blocked both inward Na+ and
Ca2+ currents. Similar results were obtained by
quinine, which blocked all voltage-gated conductances when bath-applied
to mudpuppy taste cells (Kinnamon and Roper 1988) as
well as to rat taste cells (Chen and Herness 1997
;
Ozeki 1971
; Sato and Beidler 1983
).
Blockage of inward currents is not likely to interfere with
transduction, however, because voltage- and
Ca2+-dependent K+ channels
are localized to the apical membrane in mudpuppy taste cells, whereas
Na+ and Ca2+ channels are
relatively evenly distributed on the taste cell membrane
(Kinnamon et al. 1988
). Thus bitter compounds would
likely block the K+ conductance and depolarize
the membrane potential before a substantial number of
Na+ and Ca2+ channels would
be blocked. Because both DEX and quinine are membrane permeant,
however, prolonged stimulation may result in blockage of
Na+ and Ca2+
channels due to interaction with basolateral channels. This could result in a prolonged inhibition of the taste cells and cause them to
be refractory to taste stimulation. This type of inhibitory effect has
been observed in afferent nerve recordings of catfish, where increased
nerve firing to quinine is followed by complete suppression of the
firing to all taste stimuli (Ogawa et al. 1997
). Quinine
suppression of taste responsivity also has been observed in mammals
(Formaker et al. 1997
). It is not known if prolonged DEX
stimulation causes taste cells to be in a refractory state.
Our electrophysiological data for the DEX response show some similarity
to that of quinine (Kinamon and Roper 1988) in the same
species. It will be interesting to examine
[Ca2+]i in response to
quinine in mudpuppy taste cells. Currently, however, the
autofluorescence of quinine prevents us from using conventional
ratiometric measurement of
[Ca2+]i with fura-2.
Using a different Ca2+-sensitive dye that does
not interact with the fluorescence of quinine would make it possible to
compare responses to quinine and other bitter compounds in the same
taste cells.
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ACKNOWLEDGMENTS |
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We thank Dr. Sandra L. Nelson for helpful discussions and Drs. Thomas Finger and Diego Restrepo for helpful comments on the manuscript.
This work was supported by National Institute of Deafness and Other Communication Disorders Grants DC-00244 and DC-00766 and by a grant from the Procter & Gamble Co.
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FOOTNOTES |
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Address for reprint requests: T. Ogura, Dept. of Anatomy and Neurobiology, Colorado State University, Fort Collins, CO 80523.
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.
Received 26 May 1999; accepted in final form 12 July 1999.
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REFERENCES |
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