1Departamento de Fisiología, Facultad de Medicina, and 2Departamento de Biofísica, Instituto de Fisiología Celular, Universidad Nacional Autónoma de México, Mexico City DF, 04510, Mexico
Submitted 25 February 2003 ; accepted in final form 9 December 2003
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ABSTRACT |
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kidney; Kv1.3; potassium channel; potassium transport; whole cell clamp; immunocytochemistry; confocal microscopy
Cation conductances of the IMCD membranes have been studied with microelectrodes and patch-clamp techniques. Microelectrode recordings in isolated and perfused IMCD have demonstrated the presence of a basolateral K+ and an apical amiloride-sensitive Na+ conductance (20, 41). Patch-clamp studies of the apical membrane have identified two nonselective cation channels: an amiloride-sensitive (in IMCD1) and an amiloride-insensitive cation channel (in IMCD2 and IMCD3), as well as an amiloride-sensitive Na+ channel (30, 35, 47). In a murine IMCD3 cell line, an amiloride-sensitive cation channel and two K+-selective channels have been observed in the apical membrane (36, 40). Cl conductances have been registered by whole cell clamp studies of cells from IMCD in primary culture (19, 48). The first functional findings indicating evidence of Kv channel expression in the kidney were reported in a rabbit papillary epithelial cell line and in the medulla (45, 50).
In the present work we have examined the K+-selective conductances by the perforated-patch and conventional whole cell clamp in a primary culture of IMCD from rat. Our results demonstrate the presence of a time-dependent voltage-activated outward K+ current with a high voltage activation threshold in these cells. We detected mRNAs encoding three members of the Kv1 (Shaker) family: Kv1.1, Kv1.3, and Kv1.6 in cell cultures. Western blot showed Kv1.1 and Kv1.3 protein expression in plasmatic and microsome membranes from inner medulla. Immunocytochemistry analysis confirmed Kv1.3 distribution and localization in the cytoplasm and at the basolateral membrane of collecting duct cells. Kv1.x channel subunits have been reported to assemble into heterotetrameric channels with distinct biophysical and pharmacological properties when expressed in vitro (9). We suggest that in IMCD, the functional expression of an heteromultimer of Kv1.3/Kv1.x gives rise to this outward K+ current.
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MATERIALS AND METHODS |
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Primary cultures of IMCD cells were obtained by using a modified hypotonic lysis method (30). Four male Wistar rats (175225 g) were intraperitoneally injected with furosemide (Lasix; 1 mg/100 g; Hoechst, Mexico) 20 min before death. Animals were anesthetized with chloroform and killed by decapitation. The inner medulla of the kidneys was aseptically removed and sectioned in its outer and inner halves, which were handled in parallel. The tissue was finely minced and incubated, at 37°C, for 60 min as described (30). After this incubation, 1.75 volumes of distilled water were added, and the mixture was centrifuged at 500 g for 6 min. The pellet was suspended as described (30) and centrifuged at 110 g for 2 min. This last procedure was repeated three times. The resulting pellet was suspended in DMEM (GIBCO, Grand Island, NY) supplemented as described (6) and seeded on glass coverslips contained in a 35-mm plastic petri dish (15 coverslips per dish) and kept incubated, at 37°C, in a humidified air-5% CO2 atmosphere, with medium changed every other day. With this procedure two types of IMCD cell cultures were obtained: one from the IMCD outer half and one from its inner half (IMCDo and IMCDi cultures, respectively). Five to six days after plating, both cultures showed confluent cell monolayers in at least five of the coverslips. These monolayers remained stable for at least another week. Only confluent cells were studied.
To evaluate our cultures (42), some monolayers were incubated for 40 min in PBS (GIBCO) with 0.25 mg/ml peroxidase-labeled Dolichos biflorus lectin (Sigma, St. Louis, MO). Lectin binding was observed with diaminobenzidine plus hydrogen peroxide and estimated with an inverted microscope (Diaphot 300; Nikon) equipped with Hoffman modulation optics (Modulation Optics, Greenvale, NY).
RNA Purification and RT-PCR
Total RNAs from primary cultures of IMCD cells, kidney inner medulla, and brain from rats were extracted with CsCl (39). RNA (1 µg) from each sample was converted to cDNA, using the random primer p(dN)6 (Roche Diagnostics) and reverse transcribed with SuperScript II RT (Life Technologies), according to the manufacturer's instructions. Contaminating DNA was removed by using DNase I (Pharmacia).
PCR Amplification and DNA Sequencing
cDNA products of the above-described samples were used directly as templates for PCR amplification with Taq DNA polymerase (Life Technologies). Only primers 5'-GAG THC TTC TTC GAC CG-3' (sense) and 5'-CAT GGT CAC CAC AGC CCA CCA GAA-3' (antisense; Life Technologies) (Table 1) were designed according to the conserved amino acid sequences at the amino terminus of a Kv1.x and Kv3.x and the pore regions of all Kv channels. The first PCR assays were performed with Kv1.x and Kv3.x primers; the 900-bp fragments obtained (n = 4) were cloned into the pCR 2.1-TOPO vector by using the TOPO TA cloning kit (Invitrogen). Clones were sequenced by using an ABI PRISM 310 Genetic analyzer. Three of the clones yielded a sequence corresponding to a Kv1.3. Therefore, specific primers for each of the Kv1 family members were designed (Table 1) to continue the PCR screening. cDNA fragments were purified from the agarose gel (Marligen-Biosciences) to further sequencing (n = 6).
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Western Blot Analysis
Kidneys were removed immediately after perfusion with ice-cold saline phosphate solution (SPS). The inner medulla was dissected and homogenized in a solution containing 250 mM sucrose, 1 mM EDTA, 0.1 mM phenylmethylsulfonyl fluoride, protease cocktail inhibitor (Sigma-Aldrich), and 10 mM Tris·HCl buffer, pH 7.6. Large tissue debris and nuclear fragments were removed by two low-speed spins (1,000 g, 10 min each). To separate the plasmatic (high-density fraction, HD) and microsomal (low-density fraction, LD) membrane-enriched fractions, the supernatant was centrifuged at 17,000 g for 20 min to yield a HD fraction pellet. The supernatant was centrifuged again at 100,000 g for 1 h to obtain a LD fraction pellet (25).
Protein concentration was measured by using the Coomassie brilliant blue G-250 assay kit (Bio-Rad). Samples containing 100 µg of protein were electrophoretically size separated with a 10% polyacrylamide gel and electroblotted to nitrocellulose membranes. Potassium channels Kv1.1 and Kv1.3 were detected with the use of the antibodies rabbit anti-Kv1.1 (1:100) and rabbit anti-Kv1.3 (1:200) (Chemicon International), ECL Western blotting detection reagents, and an analysis system (Amersham Biosciences). The same protocol was followed for cerebellum as a positive control.
Confocal Microscopy
Kidneys from normal male Wistar rats were fixed by retrograde perfusion via the aorta with 4% paraformaldehyde in SPS, pH 7.4 (37). The inner medulla was removed and fixed overnight. For preparation of sections, tissue was cryoprotected in 30% sucrose. The tissue was cut into 1- to 2-mm-thick slices and frozen in liquid nitrogen. Serial sections (10 µm thick) were cut in a cryostat. Sections were incubated 1824hat4°C with a rabbit anti-Kv1.3 antibody (1:40), BSA (0.5 mg/ml), FBS (3 drops/10 ml), and 0.3% Triton in SPS. After three washes with SPS (10 min each), samples were incubated with FITC-conjugated secondary antibody (goat anti-rabbit IgG, 1:1,000; Vector Laboratories) for 1 h. For simultaneous labeling of different antigens (Kv1.3 and Na+-K+-ATPase), samples were incubated at the same conditions with mouse anti-Na+-K+-ATPase or H+-K+-ATPase antibody (1:200, Sigma). This labeling was visualized by using Texas red-conjugated goat antimouse secondary antibodies (1:1,000; Vector Laboratories). Positive controls for Kv1.3 were cerebellum slices, and negative controls were renal inner medullary slices, incubated without the anti-Kv1.3 or with anti-Kv1.3 preincubated with the antigen peptide. The samples were mounted with Vectashield (Vector Laboratories). The sections were assessed by using a Bio-Rad 1024 confocal system with a Nikon TMD 300 inverted microscope. FITC and Texas red fluorophores were excited by using the 490- and 570-nm lines of the krypton-argon laser, and emission was detected by using a 520- and 600-nm long-pass filter, respectively.
Whole Cell Clamp Recordings
Membrane currents were studied mainly with the perforated-patch whole cell clamp technique (17, 38); only some experiments were performed with the conventional whole cell clamp technique (34). Coverslips containing confluent cell monolayers were placed on a superfusion chamber (0.3 ml) fixed to the stage of an inverted microscope provided with Hoffman modulation optics. Cells were maintained in a bath solution containing (in mM) 157 gluconic acid, 146 NaOH, 5 KOH, 2 Ca(OH)2, 1 Mg(OH)2, 10 HEPES, and 10 glucose, pH 7.4. Other bath solutions were used in some experiments: 1) a "Ca2+-free bath solution" containing 1 mM EGTA and no Ca(OH)2, with a calculated free Ca2+ concentration of <108 M; 2) a "15 mM K+ bath solution" containing 136 mM NaOH and 15 mM KOH; and 3) a "45 mM K+ bath solution" containing 106 mM NaOH and 45 mM KOH. All experiments were performed at room temperature (2025°C) and with 30 µM amiloride in the bath. Recording of membrane currents was performed through Ag-AgCl electrodes by using an Axopatch-1D amplifier with a CV-4 (500 M) head stage (Axon Instruments, Foster City, CA). Micropipettes from Kimax-51 glass (Kimble, Vineland, NJ) were fabricated in a two-step vertical pipette puller (PB7; Narishige). The tips were fire-polished in a microforge (MF9, Narishige). Micropipettes were filled from the tip, up to a distance of 0.40.5 mm, with a pipette solution composed of (in mM) 156 gluconic acid, 141 KOH, 10 NaOH, 1.54 Ca(OH)2, 1 Mg(OH)2, 2.3 EGTA, and 10 HEPES, pH 7.4, with a calculated free Ca2+ concentration of 3 x 107 M. Pipette filling was completed, from the back, with a pipette solution with or without 200 µg/ml amphotericin B. In some experiments a "Ca2+-free pipette solution" was employed, containing 0.1 mM Ca(OH)2 and 2 mM EGTA, with a calculated free Ca2+ concentration of <108 M. Once filled, micropipettes had a resistance of 23 M
. With the use of a hydraulic micromanipulator (WR-60; Narishige), pipettes were applied to the cells to contact the membrane, and a gentle suction was applied and maintained until a seal of at least 1 G
was obtained. Perforated-patch whole cell configuration was reached 48 min after the membrane contact as monitored when a voltage square pulse (20 mV, 5 ms) evoked a capacitative current transient shorter than 4 ms and a series resistance (Rs) smaller than 20 M
. In conventional whole cell recordings the membrane was ruptured with suction. Membrane potential was clamped at 50 mV. Membrane capacitance and Rs were compensated (80%) and then measured by using the Axopatch-1D compensation systems. The voltage-clamp protocols were generated, and the membrane currents were acquired by the Axopatch-1D under the control of pCLAMP software (version 6, Axon Instruments) running on a 486 personal computer (Gateway 200, N. Sioux City, SD) and using a Digidata 1200 analog-to-digital converter (Axon Instruments). Membrane currents were low-pass filtered at 2 kHz, digitized, and stored on the hard disk of the computer for subsequent analysis. A basic stimulation protocol was used in every cell: from a holding potential of 50 mV, a series of 720-ms voltage steps between 160 and 80 mV were applied in 20-mV increments and with 4-s intervals between the steps. Other protocols are described below. Membrane currents were analyzed by using the Clampfit module of pCLAMP, and curve fitting was performed by using SigmaPlot (Jandel Scientific, San Rafael, CA).
Current Kinetics Analysis
Time course of the capacitative current. Current was evoked by a voltage pulse from 50 to 60 mV, and its time course was fitted with a single exponential function of the form
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Time course of the voltage-activated outward current. The time course of the voltage-activated outward current was adjusted to a Hodgkin-Huxley-type model with one activation gate and one inactivation gate (15, 16). Leak current was measured [from the recordings obtained in the presence of 10 mM tetraethylammonium (TEA) or 1 mM quinidine in the bath] and subtracted. Currents were evoked by the basic stimulation protocol. The time course of current activation was fitted to a single exponential function of the form
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Voltage dependence of the outward current activation. The voltage dependence of the outward current activation was studied with a voltage protocol departing from a holding potential of 50 mV to depolarizing voltage pulses, applied every 4 s for 300 ms, from 40 to 65 mV in 5-mV increments. Leak current was subtracted as before. The peak outward current (I) recorded after each voltage step (V) was transformed into conductance (g) according to the relation I = g(V Vrev), where Vrev is the reversal potential of the current, which was assumed to be 50 mV (see RESULTS). Conductance values were fitted with the following (Boltzmann type) equation
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Tail currents. In tail current experiments, the conventional whole cell clamp technique was employed. From a holding potential of 50 mV, the current was activated by a prepulse to 40 mV during 180 ms, and then voltage was returned to various test potentials ranging from 100 to 30 mV in 10-mV increments for 32 ms. When high-K+ concentration bath solutions were used, the test potentials ranged from 100 to 0 mV. Leak current was measured (from the recordings obtained with a similar protocol in which a prepulse to 35 mV was used) and subtracted. Tail currents were fitted with a single exponential of the form
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The tail currents measured in the first 2 ms were averaged and plotted against voltage. The reversal potential of the tail currents was measured at the point where the best curve that fitted the plotted current points crossed the voltage axis.
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RESULTS |
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The monolayers observed in IMCD cultures (IMCDo and IMCDi) were mainly formed by 20- to 40-µm-diameter cells showing an almost uniform appearance: flat and polygonal, with a large nucleus (Fig. 1). This morphology is similar to that previously described for cells from the IMCD in primary culture (30, 35). Smaller cells (1020 µm) were present in some monolayers, with oval or rounded appearance. However, >85% of the cells in IMCDo cultures and >90% of the cells in IMCDi cultures exhibited positive binding for D. biflorus lectin (not shown). All these cells were flat and polygonal, indicating that these were IMCD cells in the IMCDi cultures or principal and IMCD cells in the IMCDo cultures (7, 42). Only these cells were used for the electrophysiological recordings. Therefore, electrophysiological recordings were performed in IMCD and principal cells. The monolayers exhibited blisters, providing evidence that cells were polarized and capable of transepithelial transport (29).
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Capacitance and Resistance of Cells from IMCDi and IMCDo Cultures
According to the criterion established in MATERIALS AND METHODS, perforated-patch whole cell configuration was reached in 6.2 ± 0.4 min, the time at which Rs was 13.9 ± 0.3 M (mean ± SE; n = 121). Rs became smaller during the following minutes and reached a minimum value of 9.9 ± 0.4 M
. When studied (n = 15), the time course of the capacitative current was well described by Eq. 1. A good fit of the time course of the capacitative current with a single exponential function is indicative of the absence of electrical coupling between cells (6, 19), an indispensable condition for obtaining space clamp.
Membrane capacitance was 24.0 ± 0.9 and 24.8 ± 1.0 pF in IMCDi (n = 77) and IMCDo cells (n = 44), respectively, values identical to those previously reported (19). Cell input resistance, measured by means of the current change elicited by a voltage change from 60 to 40 mV, was 1.35 ± 0.17 and 1.07 ± 0.15 G in IMCDi (n = 77) and IMCDo cells (n = 44), respectively. The difference is not statistically significant.
Membrane Currents in Cells
At the voltage range explored (from 160 to 80 mV), cells showed both inwardly and outwardly rectifying currents (Fig. 2). The present work focuses on voltage-dependent outward current.
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A time- and voltage-dependent outward current (Ivto) was observed in 27% of the cells (IMCDi: 29%, n = 77; IMCDo: 25%, n = 44). Figure 2A shows that these currents activate at potentials between 20 and 0 mV, require
100 ms to complete their activation at 0 mV, and activate more quickly with larger depolarizations. Figure 2B shows the mean current-voltage (I-V) relationship obtained in cells with Ivto. The outward rectification is due to Ivto. Considering the ionic conditions in our experiments (virtual absence of any potentially permeant anion) and the outward direction of Ivto, it can be expected that this is a K+ current. Figure 2B also shows that these cells exhibited an inward rectification and a leak component that reverses close to 0 mV, which are mainly due to an inward rectifying Cd2+-sensitive current (unpublished results).
Effect of Inhibitors on Ivto
To test whether Ivto is a K+ current, we explored its sensitivity to four K+ channel blockers. The effect of the inhibitors studied, expressed as the mean percentage block of the total outward currents recorded at membrane potentials between 0 and 80 mV, was TEA (10 mM), 82 ± 4% (n = 7); TEA (1 mM), 57 ± 2% (n = 2); quinidine (1 mM), 96 ± 1% (n = 5); Ba2+ (5 mM), 25 ± 4% (n = 2); and 4-aminopyridine (4-AP; 10 mM), 54 ± 10% (n = 2). TEA, quinidine, 4-AP, and Ba2+ inhibition of similar voltage-activated outward K+ currents has been observed in other classes of epithelial cells (6, 18, 21, 27, 45, 49). Therefore, the present experiments suggest that Ivto is a voltage-activated K+ current present in IMCD cells. Figure 3 illustrates the reversible effect of TEA on Ivto. TEA does not affect the Cd2+-sensitive cationic current, and its apparent inhibition is mainly due to its rundown (unpublished results).
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Ca+ Independence of Ivto
Time- and voltage-dependent outward K+ currents similar to Ivto may be Ca2+ dependent (6, 21). To test this possibility, we employed the conventional whole cell clamp technique with a Ca2+-free intracellular solution (pCa > 8; see MATERIALS AND METHODS). Extracellular Ca2+ removal did not reduce the Ivto current amplitude after 6 min (not shown), as should happen when the current is Ca2+ dependent (21). Also, the presence of 5 mM Cd2+ in the bath did not inhibit the current. These results suggest that Ivto is not a Ca2+-dependent current (1).
Time-Dependent Kinetics of Ivto
Current activation was followed by a slow inactivation. Figure 4A shows that the time course of current activation and inactivation can be well described by Eq. 2 or 3. Current activation shows voltage dependence, with a becoming smaller as depolarization increases (Fig. 4B). Inactivation was observed in 73% of the currents recorded at potentials between 20 and 80 mV.
i exhibited a weak voltage dependence, from 5.1 ± 1.1 s (at 20 mV) to 3.3 ± 0.8 s (at 80 mV; n = 9). Ivto activation and inactivation had
a and
i values comparable to those reported by other authors in delayed rectifier-type K+ outward currents observed in other classes of epithelial cells and fitted with similar equations (18, 21, 45, 49, 50). The similarities extend to the voltage dependence of
a and
i, which appears to be a common characteristic for this type of current when observed in epithelial cells. Therefore, these experiments suggest that Ivto is a K+ current of the delayed rectifier type. This current was the only one with delayed rectifier characteristics recorded in IMCD cultures.
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Voltage Dependence of Ivto Activation
The voltage dependence of Ivto activation is well described by Eq. 4 (Fig. 4C). The mean values of the activation parameters were Vo = 5.6 ± 3.3 mV and k = 6.8 ± 1.0 mV (n = 4). The Vo value is similar to (18, 24) or 1525 mV more positive than (45, 49) the Vo value reported for delayed rectifiers in other epithelial and nonepithelial cells. The value of k compares to those already reported (24, 46). The relative conductance tends to decline at potentials more positive than that at which gmax is reached (not shown). This phenomenon also has been observed in delayed rectifiers of other cells (12, 24).
K+/Na+ Selectivity of Ivto
To further characterize the ionic selectivity of Ivto, a tail current analysis of its reversal potential was performed. The conventional whole cell clamp technique was employed for these experiments. Figure 5A shows the tail currents recorded in a representative cell bathed in the control solution. The I-V relationship of these currents (Fig. 5B) exhibited a reversal potential of about 52 mV, which is much closer to EK (85 mV) than to ENa (68 mV). The reversal potential of the tail currents depended on the external K+ concentration. Figure 5C shows the mean reversal potentials measured in the presence of 5 (n = 9), 15 (n = 2), and 45 mM (n = 3) external K+. The straight line that joins the points is the best fit obtained with the minimum squares method; it has a slope of 30.4 that does not suggest a complete K+ selectivity. However, from this slope, the Goldman equation predicts that Ivto channels are 11.7 times more selective to K+ than to Na+. This value compares with that obtained for the delayed rectifier of the squid axon (5).
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RT-PCR Analyses of Kv.x mRNA Expression in Rat IMCD Cells
Delayed rectifying K+ channels are found in the Kv1 and Kv3 voltage-gated channel families. Therefore, our first strategy was to design primers to amplify a conserved region from the pore toward the amino-terminal end of both families. A single base band of roughly 900 bp was amplified with the IMCD cell culture and kidney inner medulla cDNA samples (Fig. 6). Sequence analysis of the 900-bp cDNA identified it as a Kv1.3 channel. Because the kinetic properties of the Ivto did not correspond to a Kv1.3 homomultimer (see above and Ref. 9), we carried out further RT-PCR reactions with specific primers for all of the Kv1.x family members (see Table 1). We observed amplification of the expected cDNA fragments for Kv1.1 and Kv1.6 (Fig. 6). GAPDH cDNA from IMCD cells and GAPDH and Kv1.x cDNAs from brain samples were always amplified as positive controls of the PCR reaction.
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Kv -Subunit Protein Expression and Localization in Inner Medulla Cells
To determine whether Kv1.x -subunits are expressed as protein, we performed Western blot experiments with commercially available antibodies directed against Kv1.1 and Kv1.3 in plasmatic membrane and microsome fractions. A representative blot (Fig. 7) identifies proteins with masses of
70 kDa that were specifically labeled with antibodies to these Kv
-subunits in both fractions.
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Confocal microscopy images of the inner medullary slices and cultures (Fig. 8) were helpful in studying the Kv1.3 distribution. Fluorescence was observed mainly in the intracellular perinuclear zones. Colabeling of Kv1.3 and Na+-K+-ATPase proteins was observed, supporting the expression of Kv1.3 -subunits at the basolateral membrane. Interestingly, this colabeling was also observed at the basolateral membrane of the papillary epithelial cells (Fig. 8F), in agreement with the hypothesis raised by others (50).
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DISCUSSION |
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The cells of IMCDi and IMCDo cultures exhibited the same type of currents. IMCDi cultures must be formed mainly by IMCD cells from the IMCD2 and IMCD3, whereas IMCDo cultures must be enriched with principal cells from the IMCD1. Both cultures exhibited positive binding for D. biflorus lectin in at least 85% of the cells, indicating that they are formed mainly by these cellular types (7). It is known that cells from different IMCD segments show important functional differences (32, 33). However, with regard to the cationic conductances we observed, all cells behaved similarly. Therefore, functional differences between IMCD segments may involve other conductance types.
Our data suggest that a Kv1.3 channel protein is present at the basolateral membrane. In contrast, patch-clamp studies performed on the apical membrane of IMCD cells in primary cultures have only identified amiloride-sensitive Na+ channels (47) and two nonselective cation channels (one amiloride sensitive and the other amiloride insensitive; Refs. 30 and 35). Kv1.2 and Kv1.3 channels have been identified in the rabbit kidney medulla and in GRB-PAP1 cells (45, 50), although their activation occurs at voltages 10 mV more negative. However, the values of the activation time constants and their voltage dependence are equivalent to those reported here.
Even when the RT-PCR and immunocytochemistry assays identified the presence of Kv1.3 channels in the IMCD cells, the biophysical and pharmacological properties of the Ivto did not correspond to those reported for a Kv1.3 homomultimer. The voltage at which Ivto is half-maximally activated (V1/2) was 5.6 mV, in contrast to the V1/2 of about 30 mV of a Kv1.3 channel (13). The time constant of Ivto inactivation (5.1 s at 20 mV) was about five to more than seven times slower than that expected for a Kv1.3 (250600 ms; Ref. 9). Also, the Ivto is about 10 times more sensitive to TEA and about 2 orders of magnitude less sensitive to 4-AP, compared with a Kv1.3 homomultimer (ID50 of 10 mM TEA and 195 µM 4-AP; Ref. 13). To compare cloned channels with native channels in IMCD cells, we have to consider that heteromultimer channels might be able to form in these cells with the resulting biophysical and pharmacological differences. The other two Kv channels (Kv1.1 and Kv1.6) identified in the cell culture also correspond to delayed rectifier channels. Although we observed expression of Kv1.1 and Kv1.3 proteins in the inner medulla, further immunoprecipitation experiments are needed to determine the Kv -subunits that conform the heteromultimer that probably gives rise to Ivto.
The physiological role of Kv1.x channels in IMCD cells would depend on the membrane (apical or basolateral) where the channels are located and the membrane potential in physiological conditions. Colabeling of Kv1.3 and the Na+-K+-ATPase in our fluorescent assays suggests that Kv1.3 is expressed at the basolateral membranes of IMCD cells. However, there is no information about the membrane potential in IMCD in physiological conditions. Conflicting results are available from in vitro studies performed in extracellular K+ concentrations of 45 mM (not physiological for IMCD; Refs. 2 and 8). Meanwhile, mean basolateral membrane potentials of 82 and 51 mV have been reported (20, 41), and it was also observed that 60% of the cells exhibited a positive membrane potential (mean: 24 mV), whereas the remaining 40% exhibited a negative membrane potential (mean: 15 mV; Ref. 43). Because the transepithelial potential difference is very small, apical and basolateral membrane potentials must be approximately the same, at least in control conditions (20, 41). Because the IMCD is exposed to two different extracellular media (tubule and inner medullar interstitial fluids) that exhibit large variations in their ionic concentrations and osmolarity (2, 3, 8, 11, 14, 22, 23), it is possible that, depending on the physiological conditions, the IMCD membrane potential (apical and basolateral) undergoes large variations. Accordingly, there will be situations in which the basolateral membrane is sufficiently depolarized to activate these Kv1 channels. This activation, by inducing a K+ efflux, would oppose further depolarization. One can speculate that Kv1 channel activation may contribute to the maintenance of the driving force for Na+ reabsorption in conditions in which Na+ is reabsorbed at a high rate by the IMCD cells. Furthermore, because of the large variations in interstitial osmolarity, Kv1 channels might also play a role in volume regulation (10).
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ACKNOWLEDGMENTS |
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GRANTS
This work was partially supported by Dirección General de Asuntos del Personal Académico (DGAPA), Universidad Nacional Autónoma de México (UNAM) Grants IN206393 and IN216396 (to J. J. Bolívar) and DGAPA UNAM Grant IN233802, Consejo Nacional de Ciencia y Tecnología Grants 41365 (to L. I. Escobar).
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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.
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