Expression and characterization of delayed rectifying K+ channels in anterior rat taste buds

Lidong Liu,2,* Dane R. Hansen,1,* Insook Kim,3 and Timothy A. Gilbertson1

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


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Delayed rectifying K+ (DRK) channels in taste cells have been implicated in the regulation of cell excitability and as potential targets for direct and indirect modulation by taste stimuli. In the present study, we have used patch-clamp recording to determine the biophysical properties and pharmacological sensitivity of DRK channels in isolated rat fungiform taste buds. Molecular biological assays at the taste bud and single-cell levels are consistent with the interpretation that taste cells express a variety of DRK channels, including members from each of the three major subfamilies: KCNA, KCNB, and KCNC. Real-time PCR assays were used to quantify expression of the nine DRK channel subtypes. While taste cells express a number of DRK channels, the electrophysiological and molecular biological assays indicate that the Shaker Kv1.5 channel (KCNA5) is the major functional DRK channel expressed in the anterior rat tongue.

transduction


THE INITIAL EVENTS in taste transduction involve the interaction of taste stimuli with receptors or ion channels localized to the apical membrane of taste receptor cells (TRCs). In most cases, the result of this interaction is the development of a depolarizing receptor potential, activation of voltage-dependent Na+ and K+ channels that underlie the generation of action potentials, and eventually the activation of voltage-dependent Ca2+ channels, leading to a rise in intracellular Ca2+ and release of neurotransmitters onto gustatory afferents (for review, see Refs. 6, 23, 31, 41).

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 {alpha}-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., {beta}-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 {alpha}-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.


    METHODS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Patch-clamp recording. Individual FF taste buds were isolated from the anterior tongues of 2- to 5-mo-old male Sprague-Dawley rats that were killed by CO2 asphyxiation followed by cervical dislocation, as approved by the Utah State University Animal Care and Use Committee. To isolate taste buds, tongues were injected beneath the lingual epithelium with ~1–1.2 ml of the following enzyme solution: 2.5 mg/ml dispase II, 1 mg/ml collagenase A (Roche, Indianapolis, IN), and 1.0 mg/ml soybean trypsin inhibitor (Sigma, St. Louis, MO) dissolved in Tyrode solution containing (in mM) 140 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, 10 glucose, and 10 Na+ pyruvate. The pH was adjusted to 7.4 with NaOH. The tongue was incubated for 25 min in Ca2+- and Mg2+-free Tyrode solution that contained 2 mM BAPTA in place of CaCl2 and MgCl2 and bubbled with O2. After incubation, the tongue was rinsed several times with Tyrode solution and the lingual epithelium was removed and pinned out serosal side up in a Sylgard-lined petri dish filled with Ca2+- and Mg2+-free Tyrode solution. Individual taste buds from the FF papillae were removed by gentle suction using a fire-polished pipette with a 100- to 150-µm bore (25). Once isolated, taste buds were placed onto Cell-Tak tissue adhesive (BD Biosciences, San Jose, CA)-coated microscope slides fitted with a Sylgard O-ring that served as a recording chamber. Cells were perfused continually with Tyrode solution containing tetrodotoxin (0.5 µM) to inhibit voltage-gated Na+ currents. In addition, use of this solution, which contained only 1 mM CaCl2, prohibited us from observing contaminating voltage-gated Ca2+ channels, which typically require 10–100 mM Ca2+ or Ba2+ to be evident in anterior taste cells (Ref. 5 and Gilbertson TA, unpublished observation). Intracellular (pipette) solution contained (in mM) 140 KCl, 1 CaCl2, 2 MgCl2, 10 HEPES, 11 EGTA, and 3 ATP. The pH was adjusted to 7.2 with KOH.

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 5–10 M{Omega} 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 ({alpha} = 0.05) compared with control currents immediately preceding the test stimulus. Data are presented as means ± SD unless otherwise indicated.


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Table 1. Pharmacological characterization of DRK currents in fungiform taste receptor cells

 


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Fig. 3. No significant difference in the inhibition of DRK channel currents was observed upon intracellular and extracellular application of TEA. A: whole cell K+ currents recorded at command potential of +40 mV during intracellular administration of 1 mM TEA or the same concentration TEA applied on both sides simultaneously. B: relative steady-state outward K+ currents at 40 mV compared with control during intracellular, extracellular, and simultaneous application of 1 mM TEA on both sides of the cell membrane. C: photoactivated rose bengal, which inhibits Kv1.5 but not Kv2 channels, inhibits a majority of DRK channel currents in taste receptor cells. Whole cell recordings of K+ current at 40 mV show a significant sensitivity to photoactivated rose bengal. More than 60% of the total K+ current in taste cells was blocked by 100 nM photoactivated rose bengal.

 
Isolation and purification of taste bud RNA. Taste buds were isolated from the FF papillae of the Sprague-Dawley rat tongue as described above. Taste buds were washed to remove nonadherent cells and immediately placed into 1.5-ml microfuge tubes with 200 µl of RNAlater (Ambion, Austin, TX). The taste buds were centrifuged at 6,000 rpm (3,300 g) for 7 min. The resulting pellet was resuspended in lysis buffer from the RNeasy Mini Kit (Qiagen, Valencia, CA), mixed rapidly on a vortex for 2 min, and then passed through a prefiltration column (MiniPrefilter column 5188-2736; Agilent, Wilmington, DE) to remove any genomic DNA. RNA was then extracted according to the instructions included with the RNeasy Mini Kit, including a 2-h DNase I treatment. For positive controls, RNA was extracted from ~100 mg of brain tissue using TRI reagent (MRC, Cincinnati, OH) according to the manufacturer's instructions.

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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Table 2. Nucleotide sequences for the primers and probes used in the PCR assays

 
Single-cell RT-PCR. Single cells from FF taste buds were obtained by injecting the FF region of male Sprague-Dawley rat tongues with a slightly different enzyme cocktail containing 1) 2.7 mg/ml dispase II (Roche, Indianapolis, IN), 2) 1.08 mg/ml collagenase A (Roche), and 3) 1.0 mg/ml trypsin inhibitor (Sigma, St. Louis, MO). The tongue was then incubated in Tyrode buffer for 45 min at room temperature. After incubation, the lingual epithelium was peeled from the tongue and pinned out in a Sylgard-lined petri dish with the serosal side up in Tyrode buffer. After the tongue was pinned, the buffer was removed and the aforementioned enzyme cocktail was added to cover the epithelium for a second digestion lasting 10 min. Next, the enzyme mixture was removed, and the epithelium was rinsed twice with Tyrode buffer before Ca2+- and Mg2+-free Tyrode buffer was added for an additional 10-min incubation. Taste buds were then extracted from the epithelium and placed on a charged slide fitted with a Sylgard O-ring and perfused with Ca2+- and Mg2+-free Tyrode buffer.

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, 300–900 nM sense and antisense primers, 300–900 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 ({Delta}CT) was calculated by subtracting the GAPDH CT from the individual DRK channel CT. Comparing {Delta}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 {Delta}CT, the greater the DRK channel expression. As previously described (39), for relative quantitation of our samples, the arithmetic formula 2{Delta}{Delta}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 {Delta}CT) for each set of pooled taste receptor cells was defined as the calibrator for that set. The calculation of {Delta}{Delta}CT involved subtraction of the {Delta}CT for each channel from the {Delta}CT calibrator value. The relative amount of target expression was determined according to the following relationship (2):

(1)

(2)

(3)

(4)
where CT is the cycle threshold for the DRK channels or GAPDH determined empirically. CTCAL is the cycle threshold for the calibrator, the most highly expressed channel in each assay. Mean relative expression values and standard deviations were calculated from the three individual sets of pooled taste bud types. To determine whether there were significant differences among the expression of DRK channels in the FF taste buds, multiple pairwise comparisons were conducted using one-way ANOVA, followed by the Bonferroni post hoc test to determine significance using SPSS version 13.0 software (SPSS, Chicago, IL).

To determine whether the efficiencies of the target and reference (GAPDH) amplification values were consistent across template dilutions, we evaluated the {Delta}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. {Delta}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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Fig. 6. Relative expression of the nine DRK channel subtypes determined by performing multiplexed qPCR assays in pooled FF taste cell mRNA. Expression was determined relative to an internal calibrator as described in Eqs. 1-4. Data are means ± SD expressing the mean of a minimum of three independent experiments, with each experiment performed in triplicate. Letters (a, b) above bars in bar graph at bottom indicate the statistical grouping variables determined using one-way ANOVA, followed by the Bonferroni post hoc test. Inset: relative PCR efficiency plots for the nine DRK channels and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) plotting log total input RNA vs. change in CT ({Delta}CT). Absolute values of the slopes (in parentheses) of the relationships in all nine assays were <0.1.

 
Immunocytochemical staining. Isolated rat FF taste buds were plated onto a glass slide coated with Adhesive Protein (Sigma) and fixed with acid alcohol fixative (70% ethyl alcohol and 10% acetic acid) for 10 min at room temperature. After three rinses in phosphate-buffered saline (PBS; in mM: 8 Na2HPO4, 2 NaH2PO4, and 154 NaCl, pH 7.5), cells were permeabilized with 0.3% Triton X-100 in PBS for 10 min at room temperature. Nonspecific protein-binding sites were blocked using incubation in TBS-TM [5% (wt/vol) nonfat dry milk, 0.15 M NaCl, 20 mM Tris·HCl, 0.01% antifoam A, and 0.1% Triton X-100] for 30 min at room temperature. To reduce bacterial contamination, 0.08% sodium azide, 100 U/ml penicillin G, and 0.1 mg/ml streptomycin were added in TBS-TM. Cells were then incubated with polyclonal anti-Kv1.5 antibody (Alomone Laboratories, Jerusalem, Israel) diluted 1:50 in TBS-TM for 72 h at 4°C. After being washed several times with TBS-TM, cells were incubated with biotin-conjugated goat anti-rabbit IgG (Dako, Carpinteria, CA) diluted 1:200 in TBS-TM for 1 h at room temperature. After being washed in TBS-TM, cells were incubated for 2.5 h at 4°C with Texas Red-conjugated streptavidin (Vector Laboratories, Burlingame, CA) diluted 1:200 in TBS-TM. Cells were then washed three times in PBS and mounted in 100% glycerol supplemented with N-propyl-gallate and examined under a Zeiss confocal microscope. For negative controls, the primary antibody was omitted.

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% {beta}-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 (4–6 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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Fig. 5. Taste buds express Kv1.5 protein. A: Western blot analysis revealing the presence of Shaker Kv1.5 channel protein in taste cells. Anti-Kv1.5 antibody labeled one prominent band in the taste sample with a molecular weight of ~66–70 kDa, which closely matches the protein size of Kv1.5. Protein from rat heart ventricular myocytes, which are known to contain Kv1.5 channels, was used as a positive control. Lanes containing the control antigen supplied with antibody (37 kDa) and the fusion protein preincubated with the antibody are also shown. BD: rat FF taste buds labeled with an anti-Kv1.5 antibody shown in confocal images of fluorescently labeled taste cells within a FF taste bud with multiple positive cells. Dotted line comprises area containing putative epithelial cells that do not show significant Kv1.5 membrane labeling (B). The differential interference contrast (C) and overlay images (D) show the apparent taste bud specific labeling pattern for Kv1.5.

 

    RESULTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
K+ currents in rat FF taste receptor cells. The properties of voltage-activated K+ currents in FF TRCs of Sprague-Dawley rats, including activation and inactivation kinetics, use-dependent inactivation and deactivation (tail currents) were studied using whole cell patch-clamp recording. Typical voltage-gated outwardly rectifying K+ currents in TRCs are shown in Fig. 1A and were representative of >95% of the >370 cells examined in this study. A subset of cells showed a more slowly activating profile; however, they were encountered so infrequently that it was not possible to characterize them in any detail pharmacologically or physiologically, and thus they were not included in the analyses. None appeared to show a more rapid inactivation profile. Currents were elicited with 400-ms depolarizing voltage steps from –80 to +40 mV in 10-mV increments with a threshold for activation near –40 mV. The channels opened rapidly upon depolarization to reach a peak with a mean (n = 24) activation time constant ({tau}a) of 4.7 ± 0.4 ms at +40 mV and then slowly inactivated, with average inactivation time constants of {tau}1 = 777.9 ± 73.1 ms and {tau}2 = 5,077.8 ± 426.2 ms for both quickly and slowly decaying components, respectively (n = 24). The currents display rapidly activating and slowly inactivating characteristics typical of DRK currents. The voltage dependence of normalized conductance (Fig. 1B) was fitted with a single, first-order Boltzmann function in the following form:

(5)
where I1 and I2 are the maximum and minimum normalized currents, respectively; V1/2 is the voltage at which half of the channels are activated (i.e., activation midpoint); and Vs is the slope of the voltage dependence. The best fit values for DRK channels in FF taste cells were V1/2 = –2.36 mV (n = 24) and Vs = 11.3 (n = 24).



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Fig. 1. Kinetics of voltage-gated outward K+ channels in taste receptor cells. A: K+ currents elicited by 400-ms voltage steps from –80 to +40 mV in 10-mV increments. B: voltage dependence of normalized conductance fitted with a Boltzmann function. C: use-dependent inactivation of K+ currents derived by a train of depolarizing steps to 40 mV at 1-s intervals from holding potential of –80 mV. D: normalized peak current related to depolarizing pulse number. E: inactivation of K+ currents tested by holding prepulse (20 s) potentials from –120 to +40 mV in 10-mV increments and then depolarizing to +40 mV for 1.5 s. F: voltage dependence of inactivation plotted as normalized current at different prepulse potentials. G: tail currents elicited by voltage steps from –90 to –20 mV after a 15-ms depolarizing prepulse to 40 mV. H: deactivation time course at different potentials.

 
No use-dependent inactivation was observed during nine repetitive depolarizing pulses to +40 mV delivered at 0.5-s intervals from a holding potential of –80 mV (Fig. 1C). The relationship between the normalized peak current and the depolarizing pulse number is shown in Fig. 1D (n = 26).

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{alpha} (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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Fig. 2. Whole cell recording of the outward delayed rectifying K+ (DRK) channel currents in taste receptor cells during application of several known Kv1.5 inhibitors. K+ currents elicited by 400-ms voltage steps from –80 to +40 mV in 10-mV increments. Data are from same cell washed with Tyrode solution between each compound. The concentrations used were 10 µM terfenadine, 50 µM bupivacaine, 10 µM perhexiline, and 20 µM quinidine.

 
Because some subtypes of DRK channels are differentially sensitive to external and internal TEA, we next compared application of TEA on both sides of the taste cell membrane using pipette perfusion. For example, Kv1.5 channels are relatively insensitive to external TEA but display sensitivity to internal TEA in the low millimolar range (4, 9, 28). Experiments comparing the effects of extracellular and intracellular TEA (Fig. 3) show that the DRK channels in anterior TRCs are moderately sensitive to TEA that is either internally or externally applied (each IC50 = ~2.0 mM). There is no significant difference in the inhibition of TEA on DRK currents between internal and external perfusion. Moreover, application of TEA at its IC50 concentration (2 mM) extracellularly after intracellular perfusion did not cause a significant increase in inhibition of DRK currents (Fig. 3B). The lack of an additive effect suggests that the same binding site for TEA is accessible to TEA applied from either side of the membrane.

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 5–10 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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Fig. 4. Expression of DRK channels by RT-PCR. A: ethidium bromide-stained gel of PCR products for the nine DRK channel subtypes in pooled taste buds from the fungiform (FF) papillae. Only Kv1.1 was consistently negative in these assays. Positive control for Kv1.1 using brain cDNA is shown. The negative control lane [(–)DNA] represents the omission of cDNA from the PCR reaction. B, top: ethidium bromide stained gel of PCR products for eight DRK channel subtypes in single taste cells for cells 8 and 9; bottom, single-cell PCR assays from 25 individual taste cells from five rats reveal variability in DRK expression. Black circles indicate positive identification by PCR, blank column spaces indicate lack of detectable expression. NT indicates that Kv1.6 was not tested in this cell. Kv1.1 was not included in these assays, because it was never found to be expressed using PCR on samples from pooled taste buds. +/n, number of positive cells/total cells.

 
Because the pharmacological experiments suggested that there was cell-to-cell variability in the effectiveness of some DRK inhibitors (e.g., stromatoxin-1), we next attempted to characterize expression of DRK channels in individual FF taste receptor cells. Cells with the elongated morphology expected of taste receptor cells were harvested, and total RNA was isolated from a total of 25 individual taste cells from five rats in independent experiments. Using this approach, we were able to test DRK expression using up to eight sets of DRK primers per cell. Because Kv1.1 was never found to be expressed in pooled FF taste buds, we omitted this channel from this investigation. Figure 4B shows the expression pattern from 25 taste cells. On average, individual FF TRCs expressed 3.8 ± 0.21 (mean ± SE; n = 25) DRK channels per cell. Although we have not attempted to analyze these data statistically, owing to the relatively low number of cells examined, all cells express at least two and as many as six different DRK channels. In particular, Kv1.5 is expressed in every cell, while the Kv3 subfamily also appears to be highly expressed. Other channel transcripts, such as Kv1.3 and Kv2.1, were rarely identified in single taste receptor cells.

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 {Delta}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).


    DISCUSSION
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DRK channels play important roles in all excitable cells by means of their critical function in regulating cell excitability. In addition, in the mammalian taste system, DRK channels may also play more central roles in specific taste transduction pathways, in which they have been reported to serve as direct or indirect targets for modulation by a variety of taste stimuli, including acids, sweeteners, bitter stimuli, and fatty acids (for review, see Refs. 23, 24, 41). While we are cognizant of the fact that DRK channels may be formed by heteromeric complexes within the subfamilies, that auxiliary {beta}-subunits interact with the pore-forming {alpha}-subunits, and that these associations may alter their physiological properties and pharmacological sensitivity (cf. Ref. 12), identification of the types and physiological properties of DRK channels is necessary to gain a fuller understanding of signaling pathways within the peripheral gustatory system. Toward this end, we have used patch-clamp recording and PCR to characterize and identify the types of DRK channels in rat FF taste buds.

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 10–30% (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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 ABSTRACT
 METHODS
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This study was supported by National Institutes of Health Grants DK-59611 and DC-02507 and the Utah Agricultural Experiment Station Project 00630, Utah State University, Logan, UT (to T. A. Gilbertson).


    ACKNOWLEDGMENTS
 
We thank Huai Zhang and Nikki D. Siears for expert technical assistance. We also thank Jeffrey T. Klein for contributions to the electrophysiological assays involving the pharmacological characterization of DRK channels and Dr. Kristina Spray and Catherine Burks for constructive comments on the manuscript.


    FOOTNOTES
 

Address for reprint requests and other correspondence: T. A. Gilbertson, Dept. of Biology, Utah State Univ., 5305 Old Main Hill, Logan, UT 84322-5305 (e-mail: tag{at}biology.usu.edu)

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. Back


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