1Department of Biology and The Center for Integrated BioSystems, Utah State University, Logan, Utah; 2Brain Research Centre, University of British Columbia, Vancouver, British Columbia, Canada; and 3Department of Pediatrics, University of Arkansas for Medical Sciences, Little Rock, Arkansas
Submitted 11 March 2005 ; accepted in final form 20 May 2005
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ABSTRACT |
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transduction
TRCs, like many neurons, apparently encode stimulus intensity in terms of the frequency of action potentials generated during chemostimulation (13, 22). As such, the biophysical properties of delayed rectifying K+ (DRK) channels may play direct roles in the rapidity of repolarization during an action potential and in the chemosensory response itself. While K+ channels play a role in the action potential and the electrical excitability of the taste cell, they also have been implicated either directly or indirectly in a number of taste transduction pathways in the mammalian gustatory system. Inhibition of K+ channels, particularly DRK channels, has been demonstrated in response to acids (16, 37), sweeteners (14, 32), bitter tastants (65), and free fatty acids (26). While evidence for DRK channel inhibition by tastants in some cases is directly at the level of the channel (e.g., acids, fatty acids, some bitters), this effect may also be indirect, occurring through second-messenger cascades that ultimately result in the closure of DRK channels and depolarization. For example, protein kinase-induced phosphorylation of members of the Shaker (KCNA) family results in closure of these channels (46). Thus an understanding of the types and properties of delayed rectifying K+ channels in mammalian TRCs is necessary to generate a more complete picture of the physiology of the peripheral taste system.
Nine types of DRK channels have been well characterized to date with the use of heterologous expression in Xenopus oocytes or mammalian expression systems; however, none have been characterized from or identified in the mammalian taste system. These nine DRK channels are related to the Shaker (KCNA), Shab (KCNB), and Shaw (KCNC) Drosophila genes and include the pore-forming -subunits of Kv1.1 (KCNA1), Kv1.2 (KCNA2), Kv1.3 (KCNA3), Kv1.5 (KCNA5), Kv1.6 (KCNA6), Kv2.1 (KCNB1), Kv2.2 (KCNB2), Kv3.1 (KCNC1), and Kv3.2 (KCNC2). While the focus of this study was on DRK channels, other members of these families encode transient, A-type K+ currents, including Kv1.4, Kv1.7, Kv3.3, and Kv3.4 (12). Although these currents were not investigated in this study, many functional K+ channels in native cells may be formed from heteromers of members within a subfamily (17), including those encoding transient K+ currents such as Kv1.4 (15). This diversity of K+ channel properties also may be enhanced by alternative splicing, interaction with auxiliary subunits [e.g.,
-subunits, minK, minK-related protein 1 (MiRP1); Refs. 52, 59], and regulation by intracellular signaling pathways, particularly those involving the phosphorylation state of the channels. Nonetheless, identification of the DRK channel
-subunits expressed, as well as their pharmacological and electrophysiological characterization, may provide insight into the functional properties of the native DRK channels (11), their molecular underpinnings in mammalian TRCs, and their potential involvement in taste transduction signaling pathways.
In the present study, we have used patch-clamp recording to characterize DRK currents, both electrophysiologically and pharmacologically, in rat fungiform (FF) TRCs to gain insight into the types of DRK channels present. RT-PCRs conducted in pooled taste buds and single taste receptor cells as well as quantitative real-time PCR (qPCR) were used to determine the relative expression of the nine DRK channels in these anterior taste buds. Consistent with both the physiological and molecular biological data is the interpretation that the major DRK channel functionally expressed in rat FF TRCs is a Shaker Kv1.5-like channel. Relatively high expression of the DRK channels from the Shab and Shaw families is consistent with the interpretation that mammalian TRCs may possess a rich array of DRK channel subtypes.
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METHODS |
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Voltage-activated currents were recorded from individual TRCs maintained in the taste bud by using the whole cell configuration of the patch-clamp technique. Patch pipettes were pulled to a resistance of 510 M when filled with intracellular solution. Series resistance and cell capacitance were compensated optimally before the recording. The holding potential in all experiments was 80 mV. For activation of DRK currents, the voltage was usually stepped from 80 to +40 mV in 10-mV increments. Commands were delivered and data were recorded using pCLAMP software (version 8) interfaced with an AxoPatch 200B amplifier with a DigiData 1322A analog-to-digital board (Axon Instruments, Union City, CA). Data were collected at 10 kHz and filtered online at 2 kHz.
K+ current inhibitors and compounds used for pharmacological study and their sources are listed in Table 1. All compounds were dissolved directly in Tyrode physiological saline solution and applied by bath perfusion unless otherwise indicated. For intracellular perfusion of compounds [e.g., tetraethylammonium (TEA); see Fig. 3], a 2PK+ pipette perfusion system was used (ALA Scientific Instruments, Westbury, NY). A quartz microperfusion capillary was inserted into the recording pipette and positioned near the patch pipette tip. Intracellular perfusion was achieved by applying positive pressure to force the perfusion solution through the quartz capillary into the patch pipette. Negative pressure was applied simultaneously through the recording pipette to neutralize the pressure in the cell. For analysis, currents during drug application were averaged over a consistent time range corresponding to the steady-state condition and compared with currents in control Tyrode solution. Tyrode solution was perfused between each application of drugs and continued until currents returned to near pretreatment levels. For analysis of DRK channels, currents were measured at a command potential of +40 mV during the steady state. Significant effects of these compounds on K+ currents were determined using paired Student's t-tests ( = 0.05) compared with control currents immediately preceding the test stimulus. Data are presented as means ± SD unless otherwise indicated.
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RT-PCR. First-strand cDNA was synthesized using the OmniScript RT kit (Qiagen). The maximum volume of taste RNA or 50 ng of brain RNA was used for the reaction in a total volume of 20 µl. Reactions were also set up in which the reverse transcriptase enzyme was omitted as a control to detect genomic DNA contamination. After first-strand synthesis, 2 µl of cDNA were added to a PCR mixture [final concentration: 500 mM KCl, 100 mM Tris·HCl (pH 8.3), 2.0 mM Mg2+, 1x Taq Master PCR enhancer (Eppendorf, Westbury, NY), 200 µM 2-deoxynucleotide 5'-triphosphate (dNTP), 500 nM forward and reverse primers, and 1.25 U of Taq polymerase]. Primers and accession numbers of the various DRK channels studied are listed in Table 2. Primers were designed using Oligo 6.0 Primer Analysis software (Molecular Biology Insights, Cascade, CO). Amplification using regular PCR included an initial 5-min denaturation step followed by 40 cycles of a three-step PCR: 30-s denaturation at 95°C, 30-s annealing at 58°C, 45-s extension at 72°C, and concluding with a 7-min final extension step. Amplified sequences were visualized using electrophoresis in 2% agarose gels poured using 1x TAE buffer (40 mM Tris-acetate and 1 mM EDTA) or real-time technology. cDNA to be sequenced was either purified directly after PCR using the QIAquick PCR purification kit (Qiagen) or extracted from agarose gels using the QIAquick gel extraction kit. Sequences were determined using the dye terminator method with a model 3100 Automatic Sequencer (Applied Biosystems, Foster City, CA).
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Single, elongated cells were located using an inverted microscope at x400 magnification and were captured using borosilicate hematocrit capillary tubes pulled and fire polished to a bore of 10 µm (collection pipette). A micromanipulator was used to position the unfilled collection pipette, and gentle suction was applied to retrieve the individual taste cell. After capture, the tip of the pipette was broken in the bottom of a 0.5-ml microfuge tube containing 20-µl cell lysis buffer (Cells-to-cDNA II kit; Ambion, Austin, TX) and kept on ice. After all cells were captured, RNA was isolated according to procedures described in the instructions provided with the Cells-to-cDNA II kit, including a 2-h DNase I treatment. One-half of the volume of this isolated RNA was used for cDNA synthesis, which eventually was divided into eight equal parts for the PCR reactions, and the other half was used for the RT control to detect genomic DNA contamination. The RT reaction was performed according to the manufacturer's directions provided in the Cells-to-cDNA II kit. The PCR reaction followed the protocol described above.
qPCR. To quantify DRK channel mRNA levels in FF taste buds, we used a two-tube RT-PCR assay with the PCR step conducted in a real-time thermal cycler (SmartCycler; Cepheid, Sunnyvale, CA). First-strand cDNA synthesis was performed as described above, with the exception that the reaction was scaled up to 100 µl. Two microliters of cDNA were used for each qPCR reaction. The HotMaster Taq DNA polymerase kit (Eppendorf, Westbury, NY) was used with the following final concentrations: 1x reaction buffer, 3.5 mM Mg2+, 200 µM dNTP, 300900 nM sense and antisense primers, 300900 nM fluorescent probes, and 1.25 U of HotMaster Taq. A two-step PCR protocol was used for the qPCR assays: 15-s denaturation at 95°C, 60-s annealing, and extension at 60°C.
We used a TaqMan detection system (Applied Biosystems) in which the primer pairs for channel-specific sequences were multiplexed with the primer pairs for the housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), for comparison of expression levels in the FF taste buds (8). Channel-specific probes were labeled at the 5'-end with FAM as the reporter fluorophore and 2,5-di-(tert-butyl)-1,4-hydroquinone (BHQ)-1 at the 3'-end as the quencher. The GAPDH probe was labeled with carboxy-X-rhodamine fluorescent dye (ROX) as the reporter fluorophore and BHQ-2 as the quencher. All probes were obtained from Integrated DNA Technologies (Coralville, IA), and their sequences are listed in Table 2. All qPCR assays were performed in triplicate, and a minimum of three independent experiments were conducted.
For quantitative analysis, fluorescent signals in the samples were plotted against the respective qPCR cycle number. The cycle at which the growth curve crossed 30 fluorescent units was defined as the cycle threshold (CT). This user-defined threshold was selected to occur during the log-linear phase of the growth curve, which is inversely proportional to the starting amount of target in the sample. Exact cycle thresholds were measured for each DRK channel as well as for the housekeeping gene, GAPDH. The change in CT (CT) was calculated by subtracting the GAPDH CT from the individual DRK channel CT. Comparing
CT values allowed for detection of relative transcript abundance between different sets of pooled taste buds by normalizing DRK channel expression to a constitutively expressed gene; therefore, the smaller the
CT, the greater the DRK channel expression. As previously described (39), for relative quantitation of our samples, the arithmetic formula 2
CT was used to take into account the amount of target normalized to an endogenous reference and relative to a calibrator. The DRK channel with the highest expression (or the lowest
CT) for each set of pooled taste receptor cells was defined as the calibrator for that set. The calculation of
CT involved subtraction of the
CT for each channel from the
CT calibrator value. The relative amount of target expression was determined according to the following relationship (2):
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To determine whether the efficiencies of the target and reference (GAPDH) amplification values were consistent across template dilutions, we evaluated the CT values for each set of DRK primers and GAPDH in three separate multiplexed reactions. For each of the PCR reactions, the absolute value of the slope of the log input vs.
CT was <0.1, demonstrating equal amplification efficiencies for the different starting template concentrations (cf. Fig. 6, inset). There was no effect on CT values when the GAPDH primers were either limited or not limited in the reactions.
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Western blot analysis.
Taste buds were collected from lingual epithelium (FF, foliate, and circumvallated area) of six rats. The sample was homogenized in ice-cold sample preparation solution (0.32 M sucrose and 5 mM Na2HPO4, pH 7.4, containing protease inhibitor). The debris was removed by performing centrifugation at 1,000 g at 4°C for 20 min. The resulting supernatant was further sedimented at 17,000 g at 4°C for 1 h. The final pellet was resuspended in 20 µl of PBS and denatured at 90°C for 5 min with the addition of 40 µl of loading buffer with 5% -mercaptoethanol.
The prepared protein was separated using 10% SDS-polyacrylamide gel electrophoresis and transferred onto nitrocellulose membrane. The membrane was incubated with 1% nonfat dried milk in Tris-buffered saline (TBS; 20 mM Tris and 500 mM NaCl, pH 7.6) for 1 h at room temperature to block nonspecific binding. The membrane was then incubated with primary antibody (anti-Kv1.5) diluted 1:200 in TBS containing 0.1% Tween 20 for 2.5 h and then a secondary antibody, alkaline-phosphatase-conjugated goat anti-rabbit IgG (Bio-Rad, Hercules, CA), at 1:7,500 dilution in TBS with 0.1% Tween 20 for 1 h at room temperature. After transfer and incubation with primary and secondary antibodies, the nitrocellulose was washed twice with fresh TBS containing 0.1% Tween 20 for 25 min. Detection was performed using enhanced chemiluminescence with the Bio-Rad ECL kit according to the manufacturer's directions. For positive and negative controls, the proteins extracted from rat cardiac tissue using the same method were also loaded at approximately the same amount (46 µg) on the same gel. Typically, the control antigen supplied with antibody and the fusion protein preincubated with the antibody were also loaded simultaneously. Data presented in Fig. 5A represent typical results from three independent preparations.
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RESULTS |
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DRK channels exhibit C-type inactivation during sustained depolarizations (51). Inactivation kinetics were determined by holding at prepulse potentials ranging from 120 mV to +40 mV (in 20-mV increments) for 20 s and then depolarizing to a test potential of +40 mV for 5 s (Fig. 1E). The voltage dependence of inactivation with a 20-s prepulse (n = 28) is shown in Fig. 1F. The inactivation curve is plotted as the current elicited by the test potential after each prepulse potential normalized to the current elicited on stepping to the test potential in the absence of a prepulse potential, and the curve is fit with a Boltzmann function. The prepulse voltage required for half inactivation (V1/2) is 35.5 mV, and the slope factor is 9.49.
To determine deactivation rates for the K+ current as a function of voltage, tail currents were elicited by stepping from a conditioning pulse of +40 mV to test potentials from 90 to 20 in 10-mV increments (Fig. 1G). The closing rate of the channels increased at more negative potentials. Tail-current time constants at different deactivation potentials were obtained by fitting single-exponential functions to the decay of the K+ current during repolarization (Fig. 1H).
Pharmacological characterization of the DRK channels in TRCs.
In this study, we examined 23 commercially available compounds that have activity as blockers of various types of K+ channels. The pharmacological profile of the K+ channels in rat FF TRCs and the concentrations of inhibitors tested are summarized in Table 1. The K+ channels in anterior TRCs are resistant to specific Ca2+-activated K+ channel blockers such as apamin and iberiotoxin (7, 20), although these currents have been reported in posterior (i.e., vallate) taste cells (10). They are also insensitive to a number of voltage-activated K+ channel inhibitors, including agitoxin (ATX), charybdotoxin (CTX), dendrotoxin (DTX), margatoxin (MTX), and mast cell degranulating peptide (MCDP), which were reported to induce inhibition in with varying affinities most DRK channels in the Shaker family, except Kv1.5 (9, 28, 56). Tityustoxin K (TsTX), a noninactivating K+ channel inhibitor that inhibits Kv1.2 and Kv1.3 channels (54), was inactive as well. However, the K+ channels in FF TRCs are sensitive to a number of known Kv1 inhibitors, including 4-aminopyridine (4-AP), bupivacaine, flecainide, nifedipine, perhexiline, quinidine, quinine, terfenadine, and TEA (Table 1). All of the aforementioned compounds inhibited >67% of the total outward current at +40 mV. Compounds known to share a common feature in their ability to inhibit Shaker Kv1.5 channels were all effective in inhibiting DRK currents in rat FF taste cells (Fig. 2), and these compounds (terfenadine, perhexiline, bupivacaine, and quinidine) were able to inhibit K+ currents in every cell examined. The Kv2-specific toxin, stromatoxin-1, had a more modest, yet highly variable, effect reflective of cell-to-cell variability (i.e., only 6 of 13 cells showed stromatoxin-1 sensitivity). Inhibition of the total outward current in the six responsive cells at +40 mV was 32 ± 17% (n = 6), but the relative block was greater at 0 mV (47 ± 21%), consistent with the voltage-dependent nature of its inhibition (19).
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In addition to the K+ channel inhibitors listed in Table 1, the effect of rose bengal, a classic generator of reactive oxygen species, on the K+ channels of FF taste cells before and after photoactivation was also tested in the present study. Photoactivated rose bengal is known to inhibit Kv1.5 channels functionally expressed in Xenopus oocytes, while expressed Kv1.2, Kv2.1, and Kv2.2 channels are insensitive to the same treatment (18). In this experiment, 100 nM rose bengal was first extracellularly perfused for 510 min in the dark, and then the recording chamber was illuminated using the microscope illuminator for 5 min during continual perfusion of rose bengal. Figure 3C shows that the DRK channels in these TRCs were significantly inhibited by photoactivated rose bengal (100 nM), while the same concentration rose bengal in darkness blocks only <15% of K+ currents. On average (n = 8), photoactivated rose bengal inhibits >60% of the total K+ current in FF taste cells. This result implies that the main DRK channels in anterior TRCs are hypoxia-sensitive channels, consistent with the presence of functional Shaker Kv1.5 channels in these cells (18). The residual current with slower activation kinetics may represent a hypoxia-insensitive current such as that carried through Kv2 channels, consistent with the positive effects of the Kv2 inhibitor stromatoxin-1 (Table 1).
Expression of DRK channels in FF taste buds. To further identify the subtypes of DRK channels expressed in rat FF taste buds, we performed RT-PCR on mRNA isolated from pooled rat FF taste buds. PCR products for seven of the nine DRK channels were identified in pooled FF taste buds in a minimum of three independent experiments. With the exception of Kv1.1 and Kv1.6, the remaining DRK channels were expressed in rat taste buds (Fig. 4A). Sequencing of the PCR products confirmed the identity of each of the seven DRK channels found in RNA isolated from FF taste buds (data not shown). The rapidly inactivating member of the Shaker family, Kv1.4 (KCNA4), was not found in FF taste buds upon performing RT-PCR (data not shown).
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To confirm expression of Kv1.5 protein, we performed Western blot analysis and immunocytochemistry on rat FF taste buds (Fig. 5). In addition to the detection of Kv1.5 protein in taste bud lysate from pooled FF taste buds of six rats (Fig. 5A), multiple cells in the taste buds were labeled with an anti-Kv1.5 antibody, while cells ectopic to the taste bud, presumably epithelial cells, did not show obvious membrane labeling (Fig. 5B). We did not attempt to verify protein expression of other DRK channels in the present study, owing to the difficulty of obtaining sufficient protein from taste buds for Western blot analysis, coupled with the lack of commercially available, highly specific antibodies for immunostaining in taste buds.
Quantification of expression of DRK channels using qPCR.
To quantify expression of DRK channels in FF taste buds, a series of multiplexed Taqman-style qPCR reactions were run on pooled cDNA isolated from several rats. In a single tube, primer sets for one of the nine DRK channels and a dual-labeled fluorogenic probe specific for a region within the DRK primer boundaries were multiplexed with primers and a probe for the housekeeping gene GAPDH. Each of the nine DRK channels was analyzed in this manner to determine its expression relative to GAPDH by calculating the CT values for each replicate as described by Eqs. 1-4. To compare expression among the various DRK channels, the relative expression of each channel type was determined with respect to an internal calibrator (i.e., the most highly expressed DRK channel within an individual experiment) for a minimum of three independent experiments. As shown in Fig. 6, the most highly expressed DRK channel in rat FF taste buds appeared to be Kv1.5. Four other channels, Kv2.2 (P = 0.9), Kv3.2 (P = 0.99), Kv3.1 (P = 1.0), and Kv1.3 (P = 0.26), also showed fairly high expression levels that were not significantly different in expression from Kv1.5 as analyzed using one-way ANOVA (F = 8.667; P < 0.001; df = 8), which did show that the remaining DRK channels (Kv1.1, Kv1.2, Kv1.6, and Kv2.1) had significantly lower (<3% of the expression level of Kv1.5) or undetectable expression levels in these qPCR assays compared with Kv1.5 (P < 0.05 each; Bonferroni post hoc test).
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DISCUSSION |
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Physiological properties of DRK currents in anterior taste cells. The biophysical properties of DRK channels in mammalian taste cells have been the focus of several earlier studies; yet, to date, none have attempted to determine the molecular identity of specific types of channels that contribute to the outward current. For example, Béhé et al. (5) performed the first series of studies on rat anterior taste buds, and their results showed TEA and 4-AP sensitivity and activation properties close to those reported herein. To identify specific functional expression of DRK subtypes, however, numerous studies have compared the biophysical properties of K+ currents in native cells with the properties of single DRK channels expressed in heterologous systems. We have characterized activation, use-dependent inactivation, and voltage-dependent inactivation and deactivation in rat taste cells both to determine the properties of these channels and as a first approach to identifying the types of DRK channels contained in these cells.
Because the DRK channels in anterior rat TRCs activate very rapidly and are slowly inactivating, quickly inactivating channels such as Kv1.4, Kv3.3, Kv3.4, Kv4.1, Kv4.2, and Kv4.3, which underlie A-type current, are unlikely to contribute significantly to the outward current in FF TRCs. The time constants for activation and inactivation of the channels in taste cells, the half-activation potential, the slope of the voltage dependence of activation, and the deactivation time constant are all similar to those of Kv1.5 channels in cardiac myocytes (45, 63) and stably expressed Kv1.5 channels in mammalian cell lines (28, 48, 56). Compared with other DRK channels, the currents in TRCs show a much faster time course of activation than those observed with Kv1.2, Kv2.1, Kv2.2, Kv3.1, and Kv3.2 channels (12, 28, 35, 45, 53, 55), although as mentioned above, we did observe a small percentage (<5%) of cells with a much slower activation time constant that appeared to be qualitatively similar to the type II cell currents in the mouse reported by Medler et al. (43). The half-activation potential for the currents in taste cells is more positive than the values for Kv1.1 and Kv1.3 but more negative than the values for Kv1.2, Kv3.1, and Kv3.2 (28, 45, 53). The peak current in TRCs does not exhibit use-dependent inactivation. However, this is in contrast to Kv1.3 channels, which show use-dependent inactivation, and Kv1.2 channels, which show a modest increase in current magnitude after repetitive stimulation (28). The K+ currents in TRCs inactivate slowly, a property that differs from that of Kv2.1 and Kv2.2 channels, which are largely noninactivating (35, 55, 64). The DRK channels in taste cells also have a considerably slower tail current compared with Kv3.1 (28). Taken together, the kinetics of the K+ currents in FF TRCs are more similar to those of native and expressed Kv1.5 channels than to those of any other single subtype of DRK channel. Unfortunately, because rat TRCs appear to contain a variety of K+ channels and the single-channel conductances of many DRK channels overlap considerably (12), single-channel analyses have been largely inconclusive in terms of identifying K+ channel subtypes in taste cells by unitary conductance measurements (Gilbertson TA, unpublished observation).
Pharmacological properties of DRK currents in anterior taste cells. Like their physiological properties, the pharmacological properties of K+ channels in FF TRCs are consistent with the interpretation that the Shaker Kv1.5 channel contributes significantly to the outward K+ currents in anterior rat TRCs. The most effective inhibitors of K+ currents in these TRCs, including terfenadine, bupivacaine, and perhexiline, appear to be those that preferentially inhibit the native or expressed Kv1.5 channels (50, 60, 61). Quinidine and photoactivated rose bengal, which is known to inhibit hypoxia-sensitive Kv channels, including Kv1.5 (3, 18, 47), also efficiently block K+ currents in TRCs. Compounds known to inhibit K+ channel subtypes other than DRK channels, such as MTX, MCDP, CTX, and DTX (9, 28, 45, 56), were ineffective in reducing K+ currents in taste cells. Thus the Kv1.5 channel is a strong candidate that contributes predominantly to the DRK current in rat FF taste cells. The only major discrepancy is the inhibitory response of the channels in TRCs to external TEA because Kv1.5 is considered an external TEA-resistant channel with an arginine replacing a tyrosine at the COOH-terminal end of the P-region (9, 28). The moderate sensitivity of the K+ currents in taste cells to external TEA may be due to the contribution of other channels. Kv2.1, for example, is one of the channels potentially blocked by external TEA because of the presence of tyrosine in the binding site (9). The fast permeation or transport of external TEA across the cell membrane or the accessibility of the TEA binding site from either side of the membrane may be alternative explanations.
Ca2+-sensitive K+ channels do not contribute to these DRK currents in FF TRCs as evidenced by the ineffectiveness of the Ca2+-sensitive K+ channel inhibitors, such as CTX, apamin, and iberiotoxin (7, 9, 20). Herness and Chen (29, 30) reported Ca2+-sensitive K+ currents in rat taste cells from the foliate and vallate papillae, however, reflective of potential differences between K+ channel expression in the anterior and posterior tongue or, alternatively, the conditions under which the present data were collected, which limited macroscopic voltage-gated Ca2+ currents. ATP-sensitive K+ channels may contribute to the total K+ conductance but are unlikely the main K+ channels in TRCs, because glyburide, an ATP-sensitive K+ channel inhibitor (1), inhibits only 1030% (maximum) of the outward K+ current in taste cells (data not shown). Most DRK channels in the Shaker family, including Kv1.1, Kv1.2, Kv1.3, and Kv1.6 can also be excluded as the predominant K+ channels, because MCDP and DTX (blockers of Kv1.1, Kv1.2, and Kv1.6 with high affinity; Refs. 9, 28), recombinant CTX (specific inhibitor of Kv1.2 and Kv1.3; Refs. 9, 28), and MTX (blocker of Kv1.3 and Kv1.6; Ref. 9) are all ineffective in significantly reducing K+ currents in taste cells. Although Kv1.2 was expressed in roughly one-half of individual taste cells examined (Fig. 4), inhibitors of Kv1.2 had no effect on DRK currents in the present experiments. This may be due to the fact that when expressed, Kv1.2 is always part of a heteromer with Kv1.5 (36, 58) with an altered pharmacological profile, or, as the qPCR data show (Fig. 6), Kv1.2 expression is exceedingly low and may not contribute significantly to the total outward K+ current.
Although Kv2 mRNA was detected in FF TRCs and a subset of TRC K+ currents is inhibited by the Kv2-specific inhibitor stromatoxin-1, the channels in the Shab (KNCB) subfamily are unlikely the predominant DRK channels in the majority of taste cells in light of the electrophysiological (see above) and pharmacological properties of the DRK channels in these TRCs. In addition, Kv2.1 and Kv2.2 are known to be hypoxia-insensitive channels (3, 18), while the K+ channels in TRCs are very sensitive to hypoxia-sensitive channel inhibitors. Nonetheless, it is clear on the basis of our data that Kv2 channels, primarily Kv2.2, do contribute to K+ currents in rat TRCs. Indeed, it may be the variable expression of channels such as Kv2.2 that contribute to some of the heterogeneity reported in K+ currents in posterior taste cells (10, 43). Kv3.1 and Kv3.2 are also unlikely candidates for the major DRK channels in TRCs according to the kinetics of the currents, although pharmacological evidence is lacking because of the unavailability of specific inhibitors for members of the Shaw family.
Expression of DRK channels in anterior taste cells. While we currently favor the interpretation that the Shaker Kv1.5 (KCNA5) channel is the major functional DRK channel in rat FF taste buds on the basis of both pharmacological and physiological analyses, it is clear from RT-PCR on pooled taste buds that TRCs express a rich variety of DRK channels (Fig. 4A). Given this diversity of DRK expression, it was surprising that we saw little variability in the physiological properties of DRK channels (personal observation), especially in light of the fact that taste buds contain multiple cell types, including functionally distinct types I, II, and III cells (40, 44). Furthermore, as the data in Table 1 show, there was little apparent variability in pharmacological profiling of DRK currents, with the notable exception of stromatoxin-1. Medler et al. (43) characterized outward K+ currents in types I, II, and III cells and showed modest and variable functional or kinetic differences among the cell types, although the molecular identities of these channels were not explored. Moreover, to isolate the DRK currents, we purposely inhibited voltage-activated Na+ channels with TTX and used an extracellular solution that limited contaminating Ca2+ currents; therefore, we could not directly link expression of these currents and DRK phenotypes in this study. To determine the variability of DRK expression at the cellular level, we performed a series of RT-PCR assays on mRNA from individual FF TRCs. Because we were able to isolate only enough RNA to run eight PCR reactions per sample, we were not able to attempt to identify cell type and DRK expression simultaneously. While our sample was small, these analyses did demonstrate cell-to-cell variability in DRK expression (Fig. 4B), which may reflect the different cell types contained within the taste bud. Future studies are required to determine the correlation between these DRK expression patterns and markers for specific taste cell types at the single-cell level.
Interestingly, in agreement with the data implicating Kv1.5 channels as important in TRCs and the apparent lack of variability in our electrophysiological recordings, Kv1.5 was expressed in all cells examined. The finding that Kv2.2 was expressed in 8 (32%) of 25 cells using single-cell PCR dovetails nicely with our pharmacological experiments that showed partial sensitivity to the Kv2 inhibitor stromatoxin-1 in 6 (46%) of 13 TRCs.
In general, there was good agreement with the single-cell PCR results and the qPCR assays of pooled taste buds (Fig. 6). Notably, the Shaker Kv1.5 channel was the most highly expressed of all the DRK channels, which may be a direct reflection that it was expressed in all FF taste cells examined. Kv2.2, Kv3.1, and Kv3.2 also showed high expression levels on the basis of qPCR and were found to be expressed in relatively high percentages of cells using single-cell PCR. There were some disparities between the single-cell PCR and qPCR data, however. Kv1.2, for example, was found in 13 of 25 individual FF taste cells, yet its expression level was only 0.2% that of Kv1.5. Kv1.6, on the other hand, was never found to be expressed in single TRCs, yet it demonstrated low to moderate expression (2% of Kv1.5) on the basis of qPCR. There are several possible explanations for this incongruity. First, single-cell RT-PCR is not quantitative; it is merely a qualitative description of what is expressed at the mRNA level. Second, the qPCR assays were conducted on pooled FF taste buds, which, as described above, contain a variety of cell types. Pooling taste buds would be predicted to capture the entire population of cells within the taste bud, while our single cells were chosen on the basis of their elongated, receptor-like morphology. Thus the latter may likely represent a subset of cells within the greater taste bud population. Small, round cells indicative of basal cells (44) would not have been selected for single-cell PCR or for the electrophysiological assays, yet clearly they would be contained in the material used for RNA isolation in qPCR. Third, while few individual cells may express a particular DRK channel such as Kv1.6, when present, it is expressed at very high levels. Clearly, it was beyond the scope of the present study, in which our aim was the initial identification and characterization of DRK channels, to rectify all of these issues.
Implications of Kv1.5 expression. Our physiological, pharmacological, and expression data are consistent with the interpretation that the Shaker Kv1.5 channel in particular may play important roles in rat FF taste buds. Interestingly, Kv1.5 was expressed in virtually every taste cell, but was not observed in non-taste bud cells using immunocytochemistry (Fig. 4B). Given our level of analysis, we cannot determine whether the native Kv1.5-like current in FF TRCs is due to homotetrameric Kv1.5 channels or heterotetrameric combinations of Kv1.5 with Kv1.2 or Kv1.3 to form the functional channel. The single-cell PCR data demonstrated that within the Shaker family, only one TRC expressed more than two Kv1 channels, one of which was always Kv1.5. To our knowledge, there have been no reports of heterotetrameric channels involving Kv1.3/Kv1.5. Heterotetramers of Kv1.2/Kv1.5 have been reported to underlie DRK currents in rabbit myocytes (36, 58). In this heterotetrameric form, the DRK currents are insensitive to homotetrameric Kv1.2 inhibitors such as apamin and charybdotoxin, which is similar to the pharmacology observed in rat TRCs. However, as our single-cell PCR and qPCR data show, Kv1.2/Kv1.5 heteromers, if they exist in mammalian TRCs, are likely to comprise the minority of taste cells.
Voltage-gated K+ channels, including Kv1.5, are functional targets for modulation by protein phosphorylation. Src family tyrosine protein kinases and cAMP-dependent protein kinase A (PKA) regulate the activity of Kv1.5 in a variety of tissues (42, 46). PKA activity and the production of cyclic AMP (cAMP) have been implicated for many years in taste transduction via the inhibition of outward K+ currents (for review, see Refs. 23, 41). In mammalian taste cells, Herness et al. (33) demonstrated a protein kinase C (PKC)-mediated inhibition of outward K+ currents, consistent with a functional role for DRK channel phosphorylation in vitro. Varkevisser and Kinnamon (62) implicated PKC activation as critical in the response to sweetener-induced inhibition of outward K+ currents. Recently, the functional coupling of the G protein-coupled taste receptors (T1R and T2R families) to inhibitory G proteins (Gi/Go) and decreases in intracellular cAMP levels have been demonstrated (49). The physiological consequences of cAMP decreases in mammalian taste cells have remained elusive (24, 41). It is plausible that a decrease in cAMP could be linked to alterations in DRK (Kv1.5) channel activity. A decrease in cAMP would in turn be predicted to decrease PKA activity and inhibit Kv1.5 currents, a pathway that has been demonstrated functionally using heterologous expression (42). Thus the link between these kinase-mediated pathways and Kv1.5 activity (and that of DRKs in general) remains an open question.
The transduction of fatty acids in the gustatory system has been suggested to mediate a taste for fat (21) by a direct inhibition of DRK channels by polyunsaturated fatty acids (PUFAs; Ref. 26). Because Kv1.5 appears to be the major DRK in FF taste buds and PUFAs inhibit >90% of the total outward K+ current in 95% of all taste cells (26), Kv1.5 is likely the predominant PUFA-sensitive channel expressed in these taste buds. A preliminary report has also demonstrated that trigeminal neurons, which may mediate the textural properties of fats in the oral cavity (57), highly express Kv1 (KCNA) channels, including Kv1.5, and respond similarly to PUFAs (27). Thus, similarly to bitter transduction in lower vertebrates (38), DRK channels in mammals may serve as a primary receptive mechanism for sapid molecules. Because fatty acids act as open channel blockers of the Kv1.5 channel (34) and because these channels are predominantly closed at taste cell resting potentials (Fig. 1B), we propose that fatty acids may act as a taste modulator by altering the time course of repolarization during and after chemostimulation. In this way, inhibition of DRK (Kv 1.5) channels may enhance taste stimulus-induced activation of taste receptor cells (26). In conclusion, the identification of Kv1.5 and the other DRK channels that we have found are highly expressed in the peripheral taste system will make it possible to try to pinpoint both direct and downstream targets that may ultimately be implicated in specific taste transduction pathways.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
* L. Liu and D. R. Hansen contributed equally to this work.
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