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Am J Physiol Renal Physiol 287: F1030-F1037, 2004. First published July 27, 2004; doi:10.1152/ajprenal.00169.2004
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Apical potassium channels in the rat connecting tubule

Gustavo Frindt and Lawrence G. Palmer

Department of Physiology and Biophysics, Weill Medical College of Cornell University, New York, New York 10021

Submitted 6 May 2004 ; accepted in final form 23 July 2004


   ABSTRACT
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METHODS
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Apical membrane K channels in the rat connecting tubule (CNT) were studied using the patch-clamp technique. Tubules were isolated from the cortical labyrinth of the kidney and split open to provide access to the apical membrane. Cell-attached patches were formed on presumed principal and/or connecting tubule cells. The major channel type observed had a single-channel conductance of 52 pS, high open probability and kinetics that were only weakly dependent on voltage. These correspond closely to the "SK"-type channels in the cortical collecting duct, identified with the ROMK (Kir1.1) gene product. A second channel type, which was less frequently observed, mediated larger currents and was strongly activated by depolarization of the apical membrane voltage. These were identified as BK or maxi-K channels. The density of active SK channels revealed a high degree of clustering. Although heterogeneity of tubules or of cell types within a tubule could not be excluded, the major factor underlying the distribution appeared to be the presence of channel clusters on the membrane of individual cells. The overall density of channels was higher than that previously found in the cortical collecting tubule (CCT). In contrast to results in the CCT, we did not detect an increase in the overall density of SK channels in the apical membrane after feeding the animals a high-K diet. However, the activity of amiloride-sensitive Na channels was undetectable under control conditions but was increased after both 1 day (90 ± 24 pA/cell) or 7 days (385 ± 82 pA/cell) of K loading. Thus one important factor leading to an increased K secretion in the CNT in response to increased dietary K is an increased apical Na conductance, leading to depolarization of the apical membrane voltage and an increased driving force for K movement out into the tubular lumen.

ROMK; BK; potassium adaptation; ENaC; sodium channels

RENAL K SECRETION HAS BEEN studied in most detail in the cortical collecting duct (CCD). This is, at least in part, because this segment is relatively easy to isolate and study in vitro using microperfusion and electrophysiological techniques. However, in vivo micropuncture results suggest that most net secretion in the rat kidney occurs upstream of the CCD (14). Thus the K content of tubular fluid leaving the late distal tubule was not much different from that of the final urine for animals on diets with normal or high-K contents.

The segment just proximal to the CCD is the connecting tubule (CNT). This part of the nephron is a good candidate for the site of K secretion. Immunocytochemical studies have documented the expression of ROMK channels (10, 15, 26), which would provide a pathway for K movement from cell to lumen, and of ENaC (2, 12, 13), which would confer an apical Na conductance and increase the driving force for K secretion by depolarizing the apical membrane. Electrophysiological data on this segment are minimal. In the only such study of which we are aware, Taniguchi and Imai (21) identified high conductance or BK channels in the apical membrane of the rabbit CNT.

We recently established techniques for the isolation of CNT's from the rat kidney and have studied the expression of apical Na channels in this segment (5). In the current paper, we have followed a similar approach to characterize and quantify the expression of K channels in the apical membrane of the rat CNT. We report that the predominant K channel is the SK or ROMK type.


   METHODS
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Animals. Sprague-Dawley rats of either sex (120–150 g) raised free of viral infections (Charles River Laboratories, Kingston, NY) were fed with either standard rat chow or a high-K diet containing 10% KCl (Harlan-Teklad, Madison, WI) for either 24 h or 1 wk.

Isolation of the CNT. The technique used to dissect and open segments of connecting tubules was described previously (5). Briefly, rats were anesthetized with an intraperitoneal injection of 150 mg/kg inactin. The kidneys were perfused in situ through the abdominal aorta with ~10 ml of cold dissection solution containing heparin. Thin coronal slices of the kidney were made with a razor blade. CNTs were isolated from thin wedges of cortex containing mostly cortical labyrinth around a radial artery.

Electrophysiology. After dissection, tubules were opened manually with a very fine needle and forceps to expose the luminal surface. The split tubules were attached to a small plastic rectangle coated with Cell-Tak (Collaborative Research, Bedford, MA) and placed in a perfusion chamber mounted on an inverted microscope. The perfusate was prewarmed to 37°C. For cell-attached recordings, the perfusing solution consisted of (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 2 glucose, and 10 HEPES adjusted to pH 7.4 with NaOH. Pipettes were made from hematocrit tubing, pulled in a three-step process, coated with Sylgard and fire polished with a microforge. Pipette resistances ranged from 2 to 5 M{Omega}. They were filled with solution containing (in mM) 140 KCl, 1 MgCl2, and 10 HEPES adjusted to pH 7.4 with KOH.

For whole cell clamp measurements, tubules were superfused with solutions containing (in mM) 135 Na methane sulfonate, 5 K methanesulfonate, 2 Ca(NO3)2, 1 MgCl2, 2 glucose, 5 mM Ba acetate and 10 HEPES adjusted to pH 7.4 with NaOH. The patch-clamp pipettes were filled with solutions containing (in mM) 2 KCl, 128 aspartic acid, 20 CsOH, 20 TEAOH, 5 EGTA, 10 HEPES, 3 MgATP and 0.3 NaGDP{beta}S with the pH adjusted to 7.4 with KOH. Amiloride-sensitive currents were measured as the difference in current with and without 10 µM amiloride in the bath solution.

Data analysis. In most patches the number of K channels was estimated from the number of current levels observed. When the number of channels was >10, it was usually difficult to observe the state in which all the channels were closed. We therefore estimated N at a pipette potential of 0 mV. Under these conditions the current across patches with no open channels (measured in patches with no channel activity or with small number of channels) was reasonably constant from patch to patch, averaging 2 pA. We assumed that this value also represents the current level in very active patches which would remain if all the channels were to close. The number of channels was then estimated using the equation: Imax – 2 = iN, where Imax was the maximal observed current level in pA and i was the single-channel current.

In a few patches the number of channels appeared to be so large that unitary events could not be resolved. In these cases we assumed that the single-channel current (i) and the open probablility (Po) were similar to values obtained for patches with only one channel. We then measured the mean current at a pipette potential of 0 mV and used the equation: Imean – 2 = iNPo, where 2 pA was again used to correct for basal current through the patch in the absence of channel activity.

In these patches, the standard deviation of the current was also measured over a period of 3–10 s, and the number of channels was estimated from the equation (7, 11): {sigma}2 = i2N(1 – Po)Po.

Statistical analysis was carried out using GraphPad InStat. Data are presented as means ± SE.


   RESULTS
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Cell-attached patches were formed on the apical membranes of flat, polygonal cells of the CNT. We presume these to be principal and/or connecting tubule cells. Their appearance contrasted with the more rounded, raised image of intercalated cells. Although we could not always identify cell types with certainty using these morphological criteria, in a previous study (5) nearly all of the putative principal/connecting tubule cells had measurable amiloride-sensitive currents, as expected for these cell types. The most abundant channel type observed in cell-attached patches with high-K solution in the pipette is illustrated in Fig. 1. This was one of a small number of patches with a single active channel; most patches had either no channel activity at all or multiple channels (see below). Currents in the absence of a pipette potential were inward, presumably due to a cell-negative electrical potential with similar K concentrations in the cell and the pipette solution. Currents reversed when the pipette voltage was more negative than –80 mV. The slope conductance for inward currents was 48 pS; outward conductance was somewhat smaller, around 30 pS. This mild inward rectification is characteristic of small-conductance K channels in the CCT. The mean value of the inward conductance for 6 similar patches was 52 ± 4 pS.



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  		Fig. 1. Current traces from a patch with a single SK channel. Dotted lines indicate the level at which the channel was closed. Upward deflections indicate inward current. Voltages indicate the negative of the pipette potential relative to the bath. Right: single-channel current is plotted as a function of voltage. Inward currents are negative.

 
Two different closed states are evident from the traces based on the length of closures. This is quantified in Fig. 2. Closed times required two exponentials, with the majority of the closures lasting < 3 ms, but a significant population > 10 ms. Open times could be described by a single exponential distribution, with a time constant of 20 ms. These kinetics were only mildly dependent on voltage. Open times increased monotonically with membrane depolarization, with a three-fold increase over the 120-mV range studied. The long closed times responded biphasically to voltage changes over this span, with maximal levels at potentials slightly hyperpolarized with respect to the resting membrane voltage. The shorter closures also decreased with strong hyperpolarizations and were fairly constant for depolarizations. Open probabilities were high, decreasing slightly with hyperpolarization (Fig. 3). This effect was due primarily to the increase in the number of long closures as illustrated in Fig. 1. These kinetic properties are very similar to those of ROMK2 expressed in oocytes (1). Thus we have identified these channels as equivalent to "SK" channels in the CCT, and presume they are the product of the ROMK gene.



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  		Fig. 2. Histograms of open and closed times. Pipette voltage was 0 mV. Open times were fit with a single exponential with a time constant of 11.4 ms. Closed times were fit with 2 exponentials with time constants of 0.72 (95% of events) and 37 ms (5% of events).

 


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  		Fig. 3. Voltage dependence of kinetics. Open probability (Po), mean open times ({tau}open) and mean closed times ({tau}closed1, {tau}closed 2) were measured at pipette potentials between +80 and –40 mV. Results show means ± SE for 6 patches.

 
A second type of channel observed under these conditions corresponds to the "BK" or "maxi" K channel seen in the CCT (3, 9) and in many other cell types including the rabbit CNT (21). These channels have a much higher single-channel conductance than do the SK channels, but usually also have much smaller open probability under physiological conditions. An example is illustrated in Fig. 4A. In this patch no channel activity was observed at pipette voltages of –80 mV or higher. When the pipette voltage was –100 mV, corresponding to a patch potential depolarized by 100 mV, brief openings appeared, which became more numerous as the depolarization increased to 140 mV. These openings revealed large unitary currents of 20 pA or more. Similar kinetics and conductances were found in the CCT (3). However, in two of five patches in which they were observed, large-conductance channels had openings in the absence of a depolarizing pipette potential. One of these patches, which contained at least 5 channels, is shown in Fig 4B. Channel activity could be observed with no potential applied to the pipette, and appeared to be cyclical with a period of about 2 s. Since these channels are activated by cell Ca+2 (3, 21), it is likely that the cell studied in Fig. 4B had an elevated cytoplasmic Ca+2 concentration. This might have reflected an activated physiological state, but the possibility of damage to the cell during the isolation procedure cannot be ruled out. Overall, these channels were much less abundant in the CNT. Such current traces were seen in only 5 of 162 patches, compared with an incidence of ~40% for the SK channels. On the basis of these observations, we conclude that the BK channels mediate a minor component of K secretion in these tubules.



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  		Fig. 4. BK channel activity. A: traces of a channel which required large depolarization of the patch to be activated. Channel openings (downward deflections) result in outward currents. B: traces of a patch with channels which were open at the spontaneous membrane potential. Channel openings produce upward deflections or inward currents.

 
The distribution of SK channels is shown in histogram form in Fig. 5. The overall density of channels was 2.0 channels/patch, but the distribution was complex. More than half (~60%) of the patches were devoid of channel activity. However, there were many more patches with large numbers (7–12) of channels than would be expected for a random sampling of a homogeneous distribution. In addition, 6 patches shown to the far right of the figure had unitary events that were similar to those of the rest of the distribution, but whose activities were difficult to estimate by counting current levels. In these patches the maximum currents measured divided by the unitary current provided a minimum estimate of channel number of 20–25/ patch. Finally, some patches had so many channels that their currents appeared only as noisy, "macroscopic" traces (Fig. 6). These currents presumably represent K movement into the cell because they disappeared when the pipette potential was –80 mV, similar to the single-channel recordings (e.g., Fig. 1). In these cases the number of channels was estimated from the mean current divided by the unitary current (3.92 pA) and the open probability (0.79) at pipette potential = 0 for 6 patches containing single SK channels. These estimates ranged from 25 to 50 channels/patch. We do not know for certain that SK channels were responsible for these large currents. However, a simple analysis of noise was at least consistent with this assumption. For 6 patches, the mean channel number estimated from the mean current was 37 ± 4. Using the standard deviation of the current, the mean number was 41 ± 7. If these patches with macroscopic currents are included, the estimate of the overall SK channel density increases to 3.3 channels/patch.



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  		Fig. 5. Histogram of channel number. Black bars represent patches from control animals. Gray bars represent patches from animals on a high-K diet. Right: bar represents patches with more than 20 channels.

 


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  		Fig. 6. Current traces from a patch with a large number of channels. In the absence of a pipette potential, the mean current was large (150 pA) and the trace was noisy. Both the mean current and the noise decreased as the membrane was depolarized, reducing the driving force for inward K current. Numbers to the right of each trace represent pipette voltages.

 
To investigate the basis for the inhomogeneity of channel density, we performed a series of measurements in which two different patches from the same cell were examined with successive pipettes. As shown in Fig. 7, there was a weak correlation between the two measured densities. For example, there were two cases where 1 patch was very active, with 6 conducting channels, whereas the paired patch on the same cell had no activity at all. Linear regression analysis provided a slope of 0.14 and a regression coefficient of r = 0.27, which was not statistically significant. However, a Spearman nonparametric correlation gave an r value of 0.42, significant at a level of P = 0.017. From this we conclude that the distribution of channels may in part be cell based, with some cells expressing higher levels of SK channels than others. However, it is also attributable to variations in channel density within the same cell.



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  		Fig. 7. Measurements of SK channel number in consecutive patches from the same cell. The number in the first patch is plotted on the abscissa, that in the second patch on the ordinate. The line indicates a linear regresssion (r = 0.27).

 
We did not detect an effect of dietary K on SK channel density. Mean densities were 3.3 channels/patch both for rats on a diet of normal chow (72 patches on 24 tubules from 6 animals), and for animals fed a high-K diet (84 patches on 20 tubules from 6 animals matched with those on a normal diet) for at least 6 days before the experiment. The distributions of densities were similar, although there were fewer patches with no channels and more patches with 1–4 channels in the K-loaded animals (Fig. 5). In particular, in both cases the fraction of patches with no channels was >50%, and there were identical numbers of patches with densities of 20–25/patch and of 25–50/patch. This was surprising since several studies have consistently found greater numbers of conducting SK channels in the CCT when animals were fed a diet rich in K (17, 19, 23, 24).

We had previously shown that elevation of dietary K for 1 wk increased the amiloride-sensitive current (INa), a measure of the number of conducting Na channels in the apical membrane, in these cells (5). To see whether upregulation of Na channels could be accomplished more rapidly, we measured INa in CNT's from rats fed a high-K diet for 24 h. An example of this measurement is shown in Fig. 8. Amiloride decreased inward currents, presumably Na currents through apical Na channels, but not outward currents. The mean INa in 31 cells was 91 ± 24 pA, with 16 cells (12 tubules, 3 animals) showing clear responses to the blocker. In control animals, 1 of 23 cells (8 tubules, 4 animals) responded to amiloride and the mean INa was 6 ± 6 pA. In animals fed a high-K diet for 6–8 days, 21/22 cells responded (10 tubules, 5 animals) and the mean INa was 385 ± 82 pA (Fig. 8).



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  		Fig. 8. Amiloride-sensitive currents in CNT cells. Typical current-voltage relationships are shown for cells from a rat on a control diet, a high-K diet for 24 h, and a high-K diet for 7 days. Squares and circles represent steady-state currents before and after application of 10 µM amiloride to the perfusate. Amiloride-sensitive currents measured at a cell potential of –100 mV are plotted as means ± SE for 23 (control), 31 (high K 1 day), and 22 (high K 7 days) determinations.

 

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K channel types in the CNT. In a previous study of apical K+ channels in the everted rabbit CNT, Taniguchi and Imai (21) found that BK channels were the predominant type in the apical membrane. These channels were recognized by their large single-channel conductance, activation by cytoplasmic Ca+2 and sensitivity to charybdotoxin. Although we also identified these channels in the rat CNT, they were rather rare, appearing in only 5 of 162 patches. In 3 of these 5, large depolarizing voltage jumps needed to be applied to the patch to open the channels. In the other two, channel activity was observed at the spontaneous membrane potential. It is unlikely that we would have missed BK channels that were spontaneously active given their large current amplitudes and characteristic gating patterns. Channels which required large depolarizations to observe their activity are much more likely to be undercounted. However, such channels would not contribute significantly to K secretion. It is possible that species differences account for part of the discrepancy of BK channel densities in the two studies. However, these channels can be activated by membrane stretch and/or flow (16, 21, 25). It is conceivable that differences in tissue preparation lead to alterations in mechanical stress on the membrane which could affect the number of conducting channels. In addition, at least in the CCT densities of BK channels were much higher in intercalated cells than in principal cells (16). The cell type expressing the BK channels in the study of Taniguchi and Imai (21) was not identified.

On the other hand, we did observe lower conductance "SK" type channels with much greater frequency. These were by far the most common K channel type in our preparations. These channels had characteristics that were essentially identical to those found in the CCT and which have been identified as ROMK channels (18). The single-channel inward conductance of ~50 pS is larger than that measured in either ROMK-expressing oocytes (8, 27) or in CCTs superfused at room temperature (4, 24) but is similar to that reported for CCTs at 37°C under similar conditions (6). The high open probability and the weak voltage-dependent kinetics of opening and closing are also similar to those observed in the CCT and in oocytes injected with ROMK2 cRNA (1). This identification of these channels is consistent with immunocytochemical localization of ROMK protein at the apical surface of CNT cells (10, 15, 26).

Channel density and effects of K loading. The overall density of conducting channels was higher in the CNT than we reported previously for the CCT of control rats. The average number of channels per patch was 2 to 3, compared with ~0.5 for the CCT. As estimated from the previous study, the membrane area for a typical patch is about 1 µm2. Thus the observed channels could account for a K conductance of 8 to 14 mS/cm2 of true membrane area. We were surprised to find no effect of prior feeding of the rats with a diet high in KCl. In the CCT, this treatment increased channels densities three- to fourfold (17, 19, 23, 24). A lack of response of SK channel density to high loading has also been observed in the rat TALH, the other segment in which expression of ROMK channels is abundant (22). However, the role of the channels in the TALH is generally considered to be one of K recycling rather than K secretion, and their expression might not be expected to be regulated. If the CNT is an important site for controlling K balance, then the rate of K secretion should be variable and should respond to changes in K intake. One possibility is that given the heterogeneity of the distribution of channels, we missed an effect of the high-K diet due to sampling limitations. This would be of particularly importance if an increase were due to a larger number of hot spots with very high channel densities. More macroscopic recording methods will probably be required to settle this issue.

We also tested whether increasing dietary K might influence the driving force for K secretion in the CNT by upregulating the activity of apical Na channels. We had shown previously that chronic high-K intake had a large stimulatory effect on amiloride-sensitive currents (5). However, to be important in regulating K excretion, such an effect should occur at least in part within the first 24 h of K loading (20). We found that INa was significantly elevated after 24 h, although clearly to a smaller degree than after 7 days. These results suggest that during increased K intake, upregulation of Na channels is at least one important factor in stimulating K secretion by the CNT.

Basis for channel clustering. One of the most remarkable features of SK channel expression in the CNT is its inhomogeneity. More than 50% of patches contained no active channels at all. In other cases we estimated densities of up to 50 channels in a single patch. This distribution is clearly inconsistent with a single randomly distributed population of channels. In that case, the distribution would fall around a single peak at or near the mean density of 3.3 channels/patch. Instead, there appeared to be at least two populations, one with densities of 0–5/patch, and another with densities of 6–12. In addition, there was yet another population, albeit a small one, with >20 channels/patch. A similar, although less striking, inhomogeneity of SK channels was previously noted in the CCT (19).

In principle, the different densities could represent different populations of tubules, variability of expression from cell to cell, or clustering within the membrane of individual cells. These phenomena are not mutually exclusive. To test these possibilities we analyzed the data in several different ways. To see whether the inhomogeneity was mainly from tubule to tubule, we analyzed a subset of data including all tubules for which we had at least 3 measurements on 3 different cells. We reasoned that if the channels were clustered by tubule, then the standard deviations for individual patches or for individual tubules should be similar. The mean ± SD for individual patches was 2.7 ± 4.8, while that for individual tubules was 2.6 ± 2.7. The decrease in SD, which itself was statistically significant, suggests that the inhomogeneity is, at least in part, within tubules and that this is to some extent smoothed out by making multiple measurements on each tubule.

Within the population of tubules described above, there were 4/21 in which at least three patches all had zero channels. We asked whether this reflected a separate population of low-activity tubules. The overall probability of finding an empty patch was 51/93 = 0.55. Thus the chance of finding three empty patches in a row was 0.16, not very different from the observed frequency of 3 consecutive empty patches (0.19). This supports the idea that the heterogeneity of tubules is not the major factor determining the distribution of SK channels.

The probability of finding a patch with 4 or more channels was 37/156 = 0.24. If the inhomogeneity were due to variations among tubules, then the chance of finding a second patch within the same tubule with a high density should be greater. The measured probability of finding a second patch with a density of 4 or higher was 16/66 or 0.24, identical with the overall incidence rate.

All these analyses suggest that the clustering of channels is at least in part due to differences among cells within a tubule or differences in membrane patches within a cell. To distinguish these two possibilities we made a series of measurements of two patches from the same cell. If the clustering is cell based, we would expect a strong correlation between the two measured densities. In fact, as shown in Fig. 7, a linear correlation had a small slope and a nonsignificant r value of 0.27. This suggests that at least part of the clustering results from hot spots within individual cells. However, a nonparametric correlation analysis of the data gave an r value of 0.42 with P ~ 0.017. This correlation suggests the presence of some heterogeneity from cell to cell as well. Although it is possible that such cell-to-cell variation reflects the presence of intercalated cells in the CNT which would not be expected to express SK channels, more than 95% of the cells from K-loaded animals expressed amiloride-sensitive Na channel currents, indicating at most a small contamination of the sample by intercalated cells. The functional consequences of channel clustering is unclear.


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This work was supported by National Institutes of Health Grant DK-27847.


   FOOTNOTES
 

Address for reprint requests and other correspondence: L. G. Palmer, Dept. of Physiology and Biophysics, Weill Medical College of Cornell Univ., 1300 York Ave., New York, NY 10021 (E-mail: lgpalm{at}med.cornell.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.


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