Amiloride-sensitive sodium current in everted Ambystoma initial collecting tubule: short-term insulin effects

N. Yvonne Tallini and Larry C. Stoner

Department of Neuroscience and Physiology, State University of New York Upstate Medical University, Syracuse, New York 13210


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Whole cell patch-clamp techniques were used to investigate amiloride-sensitive sodium conductance (GNa) in the everted initial collecting tubule of Ambystoma. Accessibility to both the apical and basolateral membranes made this preparation ideal for studying the regulation of sodium transport by insulin. GNa accounted for 20% of total cell conductance (GT) under control conditions. A resting membrane potential of -75 ± 2 mV (n = 7) together with the fact that GT is stable with time suggested that the cells studied were viable. Measurements of capacitance and use of a known uncoupling agent, heptanol, suggested that cells were not electrically coupled. Thus the values of GT and GNa represented individual principal cells. Exposure of the basolateral membrane to insulin (1 mU/ml) for 10-60 min significantly (P < 0.05) increased the normalized GNa [1.2 ± 0.3 nS (n = 6) vs. 2.0 ± 0.4 nS (n = 6)]. Cell-attached patch-clamp techniques were used to further elucidate the mechanism by which insulin increases amiloride-sensitive epithelial sodium channel (ENaC) activity. In the presence of insulin there was no apparent change in either the number of active levels/patch or the conductance of ENaC. The open probability increased significantly (P < 0.01) from 0.21 ± 0.04 (n = 6) to 0.46 ± 0.07 (n = 6). Thus application of insulin enhanced sodium reabsorption by increasing the fraction of time the channel spent in the open state.

sodium conductance; epithelial sodium channel


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

FINE CONTROL OF SODIUM REABSORPTION occurs in the distal nephron, where sodium homeostasis is achieved via regulation of the rate of its reabsorption. Sodium reabsorption is a two-step process, the entry of sodium on the apical side and the extrusion from the basolateral side (46). The Na+-K+-ATPase located in the basolateral membrane extrudes the intracellular sodium, maintaining the electrochemical gradient for the apical entry of sodium. On the apical side, sodium enters the cell through the amiloride-sensitive epithelial sodium channel (ENaC).

The highly selective 5-pS ENaC, comprised of three subunits (alpha , beta , and gamma ), is the predominant sodium channel found in the distal nephron (8, 17, 18, 30-32, 42, 44). Mutations in ENaC can lead to salt-retaining or salt-wasting disorders (18). These observations have led to the general view that ENaC is responsible for most, if not all, of the fine tuning of sodium reabsorption in the late distal nephron. Although other sodium-carrying channels have been reported in cell lines derived from various epithelial cells (for review, see Ref. 18) and in collecting tubules of potassium-adapted amphibia (45), they have not been observed in native rat or salamander collecting tubules under control conditions, making evaluation of their significance to sodium homeostasis difficult.

There is a considerable body of literature on the modulation of ENaC by hormones such as aldosterone and vasopressin. However, little is known about the regulation of sodium reabsorption by insulin, a hormone known to increase sodium reabsorption in the mammalian kidney (11, 12, 16) and amphibian model systems of the distal nephron (2, 5, 6, 9, 15, 27, 34). Increases in sodium reabsorption by insulin could be caused by several factors: 1) an increase in the open probability of the channel, 2) an increase in the conductance of ENaC, or 3) an increase in the number of active channels. The latter may represent the activation of quiescent ENaCs already present in the membrane or the synthesis and/or insertion of ENaCs into the membrane from a cytosolic pool.

The mechanism(s) by which insulin regulates sodium balance is controversial. Marunaka et al. (27) used a cell-attached patch-clamp technique on A6 cells to demonstrate that insulin increases the open probability of ENaC without any increase in active channel number. However, investigators using noise analysis techniques show no increase in ENaC open probability but do report an increase in the number of active channels in the apical membrane of A6 cells (2, 6, 15). It is unclear whether the different findings are due to the use of tissue-cultured epithelial cells, electrophysiological methods, or duration of insulin exposures.

The ability of insulin to increase sodium reabsorption occurs at physiological concentrations that are independent of changes in glucose, aldosterone, or vasopressin (10). Plasma insulin levels are known to be elevated in clinical conditions such as diabetes mellitus type II or obesity. In the obese Zucker rat, a model for type II diabetes, the expression of the beta -subunit of ENaC is upregulated in rats at 2 and 4 mo of age (3). It is not known whether the increase in this subunit correlates to an increase in sodium reabsorption. It is possible that over time (weeks, months, or years), the effects of the hyperinsulinemic condition on ENaC could increase sodium reabsorption, expand extracellular volume, and lead to hypertension (3, 10, 20).

In native renal tissue, the mechanism that insulin utilizes to increase sodium transport at the single-cell level is unknown. The distal tubule segment of the amphibian nephron is a model system of the mammalian distal nephron and shares many similarities in structure and function (22, 38-41). It is possible to evert the initial collecting tubule of the tiger salamander, Ambystoma tigrinum, and perform electrophysiological techniques on the apical membrane while separately controlling the contents of the saline bathing the apical and basolateral membranes (14, 42, 44, 45). The everted tubule is ideal for studying the effects of hormones on the cellular mechanisms of sodium reabsorption.

A whole cell patch-clamp technique was applied to the apical membrane of the everted initial collecting tubule to characterize some of the electrophysiological properties of principal cells. We found the resting membrane potential to be near the calculated equilibrium potential for potassium. The sodium conductance (GNa) was ~20% of total cell conductance (GT) and remained stable with time. These data indicate the viability of the principal cells of the everted initial collecting tubule.

We used both the whole cell and cell-attached patch-clamp techniques on everted initial collecting tubules treated with insulin in vitro. We demonstrate that insulin nearly doubled the whole cell sodium conductance. In cell-attached patch-clamp experiments, insulin caused a significant increase in the open probability of ENaC.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

General. Western variety, aquatic-phase Ambystoma tigrinum (Charles Sullivan, Nashville, TN) were housed in an aquarium at 55°F with circulating tap water and were fed crickets daily. Immediately before an experiment, a salamander was pithed and the kidney was rapidly removed. Several cross-sectional slices of kidney, 3-4 mm thick, were cut for dissection. Initial collecting tubules were dissected and trimmed to a length of 1-1.5 mm. The segment was then transferred to a modified renal tubule perfusion chamber that was mounted on the stage of an inverted microscope (Zeiss). To prevent the tubule from rolling during patch-clamp experiments, the bottom of the perfusion chamber was pretreated with Cell-Tak (Collaborative Research, Boston, MA).

Eversion and perfusion. The technique and experimental setup used to evert and perfuse kidney tubules were a modification of those developed by Burg et al. (7) and have been described in detail previously (14, 42, 43). Briefly, specialized glass pipettes were fabricated to turn the initial collecting tubule inside out, exposing the apical membrane (14, 42). To prevent the collecting tubules from sticking to the glass pipettes during the eversion process, these pipettes were soaked in a saline solution containing 1 g/100 ml albumin. A system of valves and reservoirs permitted the investigator to superfuse a variety of saline solutions over either the apical or the basolateral surface of the everted tubule. Via gravity feed, saline flowed continuously to exchange fluid in the inner perfusion pipette during the experiment, providing fluid to the basolateral surface. Both apical and basolateral flow rates were adjusted such that the exchange of solution was 95% complete within 1 min.

Patch-clamp methods. Methods to fabricate patch-clamp pipettes and form seals for cell-attached and whole cell patch-clamp studies were described previously (17, 21, 42-45). Pipettes were fabricated from 100-µl Microcaps (Drummond Scientific, Broomall, PA) on a Brown-Flaming P-80/PC puller (Sutter Instrument, San Rafael, CA) immediately before use. The pipette tips were fire polished on a Narishige microforge (Narishige, Tokyo, Japan). Pipette resistances ranged between 2.5 and 3.3 MOmega .

For both cell-attached and whole cell experiments, gigaohm seals were formed at room temperature (22-23°C). To form a seal, the pipette was positioned above an area of the everted tubule that appeared smooth and flat with a mechanical micromanipulator (WPI, Sarasota, FL). The final approach to the everted tubule was made with a Narishige hydraulic manipulator. Gigaohm seals were formed by applying suction. Only seals with a resistance of >1.0 GOmega were used.

Whole cell recordings. Whole cell voltage-clamp studies were performed by forming a gigaohm seal on the apical membrane as described in Patch-clamp methods. Intrinsic capacitance of electrodes was compensated on-line. The cell membrane was ruptured by applying suction to the pipette until an abrupt decrease in resistance and increase in capacitance was observed on the oscilloscope. The holding potential was -80 mV. Membrane potentials were stepped in 20-mV increments ranging from -100 mV to 0 mV with a pulse duration of 100 ms. The current at each voltage was determined 30-40 ms after pulse onset. GT was determined from the slope of the current-voltage plot in the linear range of -100 to 0 mV. At the end of the experiment, withdrawal of the pipette from the cell with a concurrent loss of capacitive transient demonstrated that the whole cell patch had been maintained throughout the experiment.

Two approaches were used to estimate the amiloride-sensitive sodium conductance. In the first, GT (slope from the linear regression plot of the current-voltage relationship from -100 mV to 0 mV) was determined in the presence of amiloride and subtracted from that measured before the application of drug. This difference, Delta GT, should represent the sodium conductance, assuming that amiloride does not affect other membrane conductances.

The second method measured the change in current caused by amiloride at a membrane voltage of -100 mV. Most of the change in current should be due to sodium at this voltage because it is close to the equilibrium potential for potassium (35). Thus
G<SUB>Na</SUB> = <FR><NU><IT>I</IT><SUB>amil</SUB></NU><DE>100 mV</DE></FR> (1)
where GNa is the sodium conductance, and Iamil is the amiloride-sensitive sodium current at -100 mV.

To reduce the possibility that a second messenger pathway could be disrupted because of a washout of a cytosolic factor during whole cell experiments, patches were made only after pretreating the initial collecting tubule with insulin (1 mU/ml) for 10-60 min. Others have observed the initial antinatriuretic effects of insulin by 5-15 min (37). Whole cell parameters of untreated tubules were compared with insulin (1 mU/ml)-pretreated tubules.

Cell-attached patches. The total current recorded in the cell-attached patch is defined as
I<SUB>Na</SUB> = <IT>i</IT><SUB>Na</SUB><IT>NP</IT><SUB>o</SUB> (2)
where INa is the total sodium current, iNa is the single-channel sodium current, N is the number of active channels in the patch determined as the total number of levels in a given patch, and Po is the open probability of the channel.

To eliminate the possibility that the presence of the patch pipette itself may restrict activation or insertion of ENaC into the patch membrane (26), the channel characteristics of patches on insulin-treated tubules were compared with controls. Tubules were pretreated with insulin (1 mU/ml) for 10-60 min before forming a cell-attached seal. In each condition, a series of pipette voltages between 0 and +100 mV in 20-mV steps were applied for 1-2 min. Single-channel conductance was determined as the slope of the current-voltage relationship.

Although the short recording times of 1-2 min mentioned above are adequate to determine the conductance of the channel, they may be insufficient for an accurate estimate of the number of active levels/patch or open probability. To increase the likelihood that all the levels observed were open at once, thus reducing errors in the estimate of number of levels in the patch (26), recordings were maintained at +40 or 0 mV for an average of 13 min. Recording times ranged from 5 (2 active levels/patch) to 26 (9 active levels/patch) min. Open probability was calculated from the following equation
NP<SUB>o</SUB> = <FR><NU><LIM><OP>∑</OP></LIM><IT>it<SUB>i</SUB></IT></NU><DE><IT>T</IT></DE></FR> (3)
where N is the maximum number of simultaneously active levels, i is an active level, ti is the time open at the ith level, and T is the total recording time.

Cell capacitance. Determination of capacitance from whole cell currents was described previously (17). Briefly, a 20-mV voltage step was given through the pipette command potential. Currents were recorded at 1-ms time intervals for 10 ms after the voltage step and were subtracted from the stepped current. The transient current was then fit to a single exponential decay. Capacitance is determined empirically
C<SUB>m</SUB> = <FR><NU>(<IT>I<SUB>∞</SUB>+&agr;</IT>)<SUP>2</SUP></NU><DE><IT>&agr;&bgr; · &Dgr;V</IT></DE></FR> (4)
where Cm is cell capacitance, Iinfinity is the steady-state current after the change in voltage, alpha  is the current at time zero calculated from the single exponential decay, beta  is the time constant of decay calculated from the single exponential decay, and Delta V is the change in voltage.

As observed by others, GT varied over a wide range (17). One possible explanation for this variability is differences in cell surface area. Because capacitance is believed to be a function of the membrane surface area of the cell, the value of capacitance can be used to normalize for differences in cell size. To normalize sodium conductance to surface area, GNa was multiplied by the capacitance ratio (average capacitance divided by the measured capacitance of that individual cell).

Acquisition and analysis. Single-channel data acquisition and analysis were performed as previously described (42, 44). The patch-clamp signals were monitored via an Axopatch-1D amplifier (Axon Instruments, Burlingame, CA) equipped with a TMA-1 interface (Axon Instruments). An analog-to-digital recorder (model VR-10; Instrutech, Mineola, NY) was used to create a digitized recording of the experimental data on videotape for off-line analysis of single-channel recordings. The signal for cell-attached patches was filtered to tape at 50-1,000 Hz. Cell-attached patch data were acquired at a sampling rate of 100-500 Hz for analysis. Whole cell currents were controlled by a microcomputer via an Axopatch-1D amplifier and directly loaded onto a microcomputer at 1 kHz. Data acquisition and analysis were carried out with the pClamp suite of programs (version 6; Axon Instruments, Foster City, CA) and SigmaPlot (version 5.0; SPSS, Chicago, IL).

Solutions. Normal saline solution was (in mM) 105 NaCl, 3 KCl, 2 CaCl2, 1.25 MgSO4, 1.25 KH2PO4, 5 HEPES, and 5 dextrose, pH adjusted to 7.6 with NaOH. Dissecting solution and eversion solution consisted of normal saline solution plus 1 g/100 ml of fraction V bovine serum albumin. The solution filling the cell-attached pipette was normal saline solution.

Whole cell bath solution consisted of (in mM) 105 Na gluconate, 4 KCl, 3 CaCl2, 3 MgCl2, 10 HEPES, and 5 dextrose, pH adjusted to 7.6 with NaOH. In some experiments we included 2 mM BaCl2 in this solution. To keep osmolarity constant, 1 mM CaCl2 and 1 mM MgCl2 were replaced with BaCl2.

Whole cell pipette solution consisted of (in mM) 102 K gluconate, 6 KCl, 4 MgCl2, 3 EGTA, 10 HEPES, 5 dextrose, and 2 K2ATP. The pH was adjusted to 7.4 with KOH. When potassium channel inhibitors were included in the whole cell pipette solution, the saline contained (in mM) 76 K gluconate, 6 KCl, 3 MgCl2, 15 D-gluconic acid, 20 tetraethylammonium (TEAOH), 10 CsOH, 3 EGTA, 10 HEPES, 5 dextrose, and 2 K2ATP. The pH was adjusted to 7.4 with HCl.

Insulin (bovine) was added to normal bath or whole cell bath solution, resulting in a final concentration of 1 mU/ml. Amiloride (gift from Merck, Sharp, and Dohme) was made up in whole cell bath solution to a concentration of 2 µM. Heptanol was dissolved in pure dimethyl sulfoxide (DMSO). The heptanol-DMSO solution was added directly to the whole cell bath. The apical surface was perfused with either 1 mM heptanol for 3 min or 2 mM heptanol for 5 min. The final concentration of DMSO was 0.25 or 0.5%. The osmolarity of all solutions was monitored and adjusted to 220-232 mosM. All chemicals were purchased from Sigma (St. Louis, MO) unless specified.

Statistics. Statistical analysis and graphs were completed with Sigma Plot 2001 (SPSS). Values are expressed as means ± SE. Statistical significance was determined by comparing the difference between untreated tubules and treated tubules with Student's t-test or one-tail test for the difference between independent means. A P < 0.05 was considered statistically significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Measurement of GT. We established that stable values of GT could be recorded from single cells of the initial collecting tubule. Figure 1A shows representative current traces in response to a series of 100-ms voltage steps applied from a holding potential of -80 mV. The voltage steps ranged from -100 to 0 mV in 20-mV increments and were recorded 5 min after whole cell seal formation. A GT of 7.9 nS for the cell in Fig. 1A was determined from the slope of the current-voltage plot (Fig. 1B). GT varied over time after seal formation as illustrated in Fig. 1C. GT was determined immediately after breaking into the cell and was followed for up to 17 min in a group of six cells. A decline in GT was fit with an exponential decay function with a time constant of 3 min. GT was essentially stable by 5 min (8.5 ± 0.7 nS; n = 6) and was not statistically different from values obtained at 17 min (7.3 ± 1.2 nS; n = 5). The course of initial decay in GT was not investigated, but it may reflect dialysis with the whole cell pipette solution contents or some time-dependent increase in the seal resistance. We chose to determine GT in all future experiments at least 5 min after seal formation so that all the solution changes presented to the apical or basolateral membrane occurred when currents were stable.


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Fig. 1.   Measurement of cell currents and total cell conductance (GT) in principal cells of the amphibian collecting tubule. A: whole cell current traces recorded in 20-mV increments between 0 and -100 mV. The currents were recorded 5 min after breaking into the cell. B: current-voltage plot of the data in A. The line was drawn by linear regression analysis; the slope, GT, is 7.9 nS, and the x-axis intercept is the resting membrane voltage of -74 mV. C: time course of GT after breaking into the cell. The decline in GT was fit to a single exponential decay with a time constant of 3 min. Bars indicate SE.

Two types of cells are present in the initial collecting tubule, principal cells and intercalated cells (18, 28). The principal cell is involved in sodium transport, and the intercalated cell is involved in hydrogen ion regulation (18, 28, 29). Amiloride is a rapid inhibitor of ENaC, and its effect on GT has been used as a measure of the sodium conductance (17, 18, 32, 41). Figure 2A shows a current-voltage relationship from a cell before and after exposure of the apical membrane to 2 µM amiloride for 1 min. The decrease in GT caused by amiloride (2 µM) is presumably due to the inhibition of apical ENaC. The presence of an amiloride-sensitive GT is taken to indicate that the cell being studied was a principal cell. Amiloride (2 µM) caused a hyperpolarization of the resting membrane voltage, as expected. Figure 2B shows current-voltage plots before and after exposure to amiloride (2 µM) from another cell. This particular cell did not exhibit an amiloride-sensitive conductance and was therefore considered an intercalated cell. In this study, we exposed 23 cells to 2 µM amiloride; 4 (17%) of the cells lacked an amiloride-sensitive conductance and were discarded from the data pool.


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Fig. 2.   Representative current-voltage plots of the effect of amiloride on amiloride-sensitive (A) and amiloride-insensitive (B) cells of the amphibian collecting tubule. After a control voltage step, 2 µM amiloride was placed on the apical membrane for 1 min before another voltage step was completed. , GT before application of amiloride; open circle , amiloride-insensitive conductance after amiloride application. The changes in slope and hyperpolarization seen in A indicate that this cell is a principal cell.

Measurement of GNa. We used two different approaches in this study to estimate sodium conductance. In the first, the total and amiloride-insensitive conductances were determined from the slope of the current-voltage plot as shown in Fig. 2. The amiloride-sensitive change in GT (Delta GT) was determined by subtracting the amiloride-insensitive slope conductance from the GT. Using this method of subtracting the two slopes assumes that amiloride blocks only the sodium conductance. Figure 3A shows a linear increase in Delta GT, with time, after exposure of the apical membrane to amiloride (2 µM). The mean values of Delta GT were significantly higher at 5 (P < 0.05) and 8 (P < 0.005) min after application of amiloride (2 µM) compared with the 1 min value. Because the difference in slope conductances is not a true GNa in our model system, we refer to this value as Delta GT.


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Fig. 3.   Two methods to determine the sodium conductance in principal cells of the amphibian collecting tubule after 2 µM amiloride exposure. The same principal cells were used in both methods. , Change in GT (Delta GT) determined by subtracting the 2 slope conductances; open circle , GNa computed by Eq. 1 (see MATERIALS AND METHODS). A: data from principal cells in which the potassium conductances were not inhibited. B: data from principal cells in which the potassium conductances were inhibited by perfusing the apical and basolateral membranes with a solution that contained BaCl2. The interior of the cells was dialyzed with a whole cell pipette solution that contained tetraethylammonium (TEAOH) and CsOH. Bars indicate SE; nos. of experiments at each point are in parentheses. *P < 0.05; #P < 0.005.

The second method that we used to estimate the amiloride-sensitive conductance was to measure the change in current caused by amiloride when the cell was clamped to a membrane voltage of -100 mV. Because this voltage is close to the equilibrium potential for potassium, the observed current is presumably due to sodium (35). (The chloride concentrations of our whole cell saline solutions were lowered to ~16 mM.) Figure 3A shows GNa determined by Eq. 1 (see MATERIALS AND METHODS). GNa averaged 1.3 nS and was stable with time. This value was identical to Delta GT when extrapolated to time 0 (Fig. 3A). The stability of the amiloride-sensitive sodium current at -100 mV led us to choose this method to compute GNa.

The time-dependent increase in apparent sodium conductance would be consistent with amiloride directly or indirectly blocking a conductance other than sodium, perhaps potassium. To test whether amiloride caused a secondary decrease in a potassium conductance, we exposed the apical and basolateral membranes to a whole cell bath solution that contained BaCl2 while dialyzing the interior of the cell with a pipette solution that contained TEAOH and CsOH (32). In Fig. 3B the amiloride-sensitive Delta GT and GNa were calculated as in Fig. 3A. The potassium-inhibiting solutions abolished the linear increase in the amiloride-sensitive Delta GT that was observed in Fig. 3A. GNa remained stable with time. Our data are consistent with the notion that the increase in Delta GT shown in Fig. 3A is due to a secondary affect of amiloride on a potassium conductance.

Cell coupling. Connexins allow electrical communication to occur between cells and have been reported to be present in the kidney (4, 19). If current passes between cells via connexins, our method of measuring GNa would provide an overestimate. Importantly, efforts to evaluate experimental perturbations that change the GNa could be complicated by any changes in the resistance of the connexin and not the channel under observation. Therefore, to accurately measure single-cell conductances, the cells must be electrically isolated. No information is available regarding the presence of connexins or electrical coupling of principal cells in the collecting tubule of Ambystoma.

One way we addressed this question was by measuring the capacitance of the cell. Theoretically, coupled cells would present multiexponential decay of current, whereas electrically isolated cells would express a single exponential decay transient (13, 17). The measurement of cell capacitance was determined by fitting the transient current response from a +20-mV step to a single exponential decay (Fig. 4). The fit line in Fig. 4 had a time constant of 0.566 ms. In the experiments in which the capacitance could be determined in untreated tubules, the average capacitance was 63 ± 5 pF (n = 6; Table 1). In 15% of the cells studied, the transient decay was too rapid and resulted in a poor fit. Because the line was best fitted to a single exponential, not a multiexponential, the cells were not well coupled, if coupled at all (13).


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Fig. 4.   Measurement of cell capacitance. A representative transient current decay observed when the cell voltage clamp was changed by 20 mV is shown. Current values were obtained by subtracting the current at 1-ms intervals from the maximum current at time zero. The line was drawn as a best fit to a single exponential. The time constant of decay was 0.566 ms, corresponding to a capacitance of 65 pF. See Eq. 4 for the calculation of capacitance from these data.


                              
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Table 1.   Whole cell parameters in principal cells of Ambystoma collecting tubule

Another way we evaluated electrical coupling between cells was to apply compounds that cause connexins to disconnect. Heptanol has been shown to uncouple connexins in cardiac myocytes within a few minutes of its application (1). When cardiac myocytes were uncoupled, there was a decrease in the junctional current that resulted in a decrease in GT. In a series of seven cells we showed a lack of an effect of heptanol (Fig. 5). Measurements of GT were made every minute while the apical surface of the everted initial collecting tubule was perfused with 1 mM heptanol for 3 min or 2 mM heptanol for 5 min. GT was determined from the slope of the linear regression plot from -100 to 0 mV. There was no apparent effect on GT of dose of heptanol or the amount of time the cell was followed. There was no statistical difference in GT before (8.3 ± 2.6 nS; n = 7) or after heptanol (8.0 ± 2.3 nS; n = 7).


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Fig. 5.   Lack of effect of heptanol on GT in principal cells of the amphibian collecting tubule. Each symbol represents a separate cell followed before and after the addition of heptanol. GT was computed from the slope of the linear regression analysis from voltage steps ranging from -100 to 0 mV in 20-mV steps.

Effect of insulin. Table 1 compares measurements of whole cell parameters from untreated initial collecting tubules and tubules pretreated with insulin (1 mU/ml) by perfusion of the basolateral membrane for 10-60 min. The mean capacitance in cells was not different in the two groups, indicating that membrane surface area is similar in both groups. After a 1-min application of 2 µM amiloride to the apical membrane, there is an expected hyperpolarization that is significant in both the control and insulin-pretreated tubules (P < 0.01). The normalized GNa is 70% higher in the insulin-pretreated tubules compared with the untreated tubules (2.0 ± 0.4 nS vs. 1.2 ± 0.3 nS; P < 0.05). Thus insulin does increase the sodium conductance and presumably the sodium transport across the apical membrane in the principal cells of the Ambystoma collecting tubule.

Cell-attached patch clamp. To investigate the single-channel mechanism responsible for the increase in the normalized GNa, we used the cell-attached patch-clamp technique on the apical membrane of the everted initial collecting tubule. Three changes in ENaC activity could account for the increase in GNa: 1) an increase in the number of active channels, 2) an increase in open probability, and 3) an increase in conductance.

Others, when studying the effect of vasopressin on ENaC, suggested that the presence of the patch-clamp pipette might have prevented the incorporation of new channels into the membrane under the patch pipette (26). Because this limitation could also apply to our tubules, we chose to evaluate the effect of insulin by comparing control tubules to insulin-treated tubules (1 mU/ml). Seals were formed on the apical membrane of principal cells after the basolateral surface was perfused for 10-60 min with an appropriate saline solution.

Table 2 shows the results of the above experiments. There was no statistical difference between the control and insulin-pretreated (1 mU/ml) tubules with regard to single-channel conductance or pipette reversal potential. The data in Table 2 reporting NPo were collected from cell-attached patch-clamp experiments in which the patch could be held for a long period of time (see MATERIALS AND METHODS). The average NPo for insulin-pretreated (1 mU/ml) cells was 2.6 ± 0.9 (n = 6), a value 70% greater than that seen in untreated cells (1.5 ± 0.7, n = 6). The apparent change in NPo was not statistically significant. It is of interest that our whole cell sodium conductance also increased by 70% in insulin-treated cells (see Table 1). The failure to see a significant increase in NPo is presumably related to the fact that there is a wide variability in the number of active levels per patch. However, the open probability of ENaC was significantly higher (P <=  0.01), from 0.21 ±0.04 to 0.46 ±0.07 (n = 6), in patches on insulin-treated cells (Table 2).

                              
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Table 2.   Cell-attached parameters in principal cells of Ambystoma collecting tubule

Although the number of active levels observed in patches that contained ENaC did not appear to change, the possibility arose that the frequency of patches that contain any ENaC could have increased or decreased. Therefore, we computed the relative abundance of levels in each group of patches. Relative abundance was calculated as the fraction of successful seals that contained sodium channels multiplied by the average number of levels per patch. Insulin did not appear to alter the relative abundance of ENaC (Table 2).

In recordings taken from cells exposed to insulin, the increase in open probability is apparent to the eye, especially when the number of active levels in the patch is small. Figure 6 shows representative current traces in a control cell (Fig. 6A) and a cell pretreated with insulin (Fig. 6B). The pipette holding potential is +40 mV in both conditions. A dashed line indicates the closed state. In Fig. 6A two active levels are observed with an NPo of 0.42, whereas in Fig. 6B three active levels are observed with an NPo of 1.0. It should be noted that in the second trace of Fig. 6B, the first level is open throughout the entire trace.


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Fig. 6.   Representative current traces of epithelial sodium channels (ENaC) in apical membrane patches of principal cells from control and insulin-treated tubules. Current traces recorded from a pipette holding potential of +40 mV in an untreated cell (A) and a pretreated insulin cell (B). In each case, traces were selected so that both the closed state and the maximum number of levels observed are visible. The dashed line indicates the closed state. Inward current is downward.

The data presented in Table 2 and Fig. 6 indicate that insulin enhanced sodium reabsorption, at least in part, by increasing the fraction of time the channels spent in the open state. Because of the variability in the number of active levels per patch, an increase in the number of active channels in insulin-treated patches cannot be excluded.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

There are two cardinal observations of this study. First, we demonstrate the ability to measure ionic conductances in single principal cells of everted Ambystoma initial collecting tubules. Second, this is the only study in native renal tissue to demonstrate an effect of insulin on the amiloride-sensitive ENaC located on the apical membrane.

We previously reported (14, 43-45) that the collecting tubules of both rats and salamanders can be everted and perfused in vitro. One important advantage of the everted tubule as a biological preparation is that it provides access to the apical membrane for electrophysiological studies while permitting independent control of the saline solution to which the apical and basolateral membranes are exposed.

Two types of cells are present in the initial collecting tubule, principal cells and intercalated cells. The principal cells have characteristically been described as polygonal in shape and are involved in sodium transport, whereas the intercalated cells are rounded or elongated in shape and are involved in hydrogen ion transport (18, 29). In rabbit and rat collecting duct, the distribution of the amiloride-sensitive ENaC is limited to principal cells, not intercalated cells.

Initially, we made no attempt to identify the type of cell being studied. The pooled resting membrane potential of these cells averaged -76 ± 1 mV (n = 18). This is a value very close to the potassium equilibrium potential (-84 mV) calculated from the concentrations of potassium in our bath and pipette solutions. Therefore, the predominant conductance was for potassium, suggesting viability of the cells studied. The GT of these same cells was stable and averaged 8.2 ± 1.2 nS (n = 18) >= 5 min after seal formation, another observation consistent with healthy cells. This value is similar to that reported for rat cortical collecting duct principal cells (17) and cultured mouse cortical collecting duct cells (23).

We used the amiloride sensitivity of GT as a feature to distinguish principal cells from intercalated cells. Nearly 83% (19 of 23 cells) of the cells in which whole cell patches were treated with amiloride had an amiloride-sensitive conductance. This value was comparable to a 75% incidence of principal cells reported in the initial collecting tubule of a similar salamander, Amphiuma means (38). In previous studies that used cell-attached patch-clamp methods, nearly 80% of successful seals expressed ENaC (44, 45).

The whole cell patch-clamp technique allows the investigator to electrically and chemically control the cell. The cells must be electrically isolated for this experimental technique to yield data that represent the functions of a single cell and not that of a syncytium. In a syncytium, gap junction proteins (connexins) establish cytoplasmic continuity between adjacent cells for electrical and metabolic communication (4). Nine different connexins, at the mRNA or protein level, have been identified in the kidney (19). Freeze-fracture electron and scanning microscopy show gap junctions to be present in the proximal tubule segment but not in the distal tubule segment in reptiles (33), amphibians (22, 38) and mammals (24). In the rat cortical collecting duct, the lack of movement of a fluorescent dye into neighboring cells after it was injected into a principal cell suggests that the cells of the mammalian distal nephron are not coupled (17).

We used two approaches to test for the presence of cell coupling in our tissue. First, we determined the capacitance of the principal cells under investigation. Others have noted that the presence of cell coupling should present capacitive discharge profiles that fit a multiple exponential decay (13). The capacitive discharge of our cells fit a single exponential decay, suggesting that the cells were not electrically coupled.

In the rat distal nephron other groups calculated the cell capacitance from known morphometric estimates of surface area of principal cells, which compares favorably to measured capacitance (17). [It is widely accepted that biological membranes have a specific capacitance of 1 µF/cm2 (13).] Although morphometric estimates of cell surface area are lacking for our species, they do exist for Amphiuma (38), a species very similar to Ambystoma. From their values of surface area of the amphibian principal cell, we calculate the capacitance to be 57 pF/cell. Our value of capacitance in untreated tubules is 63 ± 5 pF/cell. The similarity between these values argues that there is little, if any, electrical coupling between principal cells in our preparation.

The second method to test for electrical coupling is to apply an agent known to uncouple connexins, heptanol, while monitoring the current (1). If cells are coupled, then heptanol should decrease the total cell conductance. We observed no change in total cell conductance when heptanol was added to the apical surface. This evidence also supports the notion that the cells of the amphibian initial collecting tubule are not well coupled.

The apparent viability and stability of the everted initial collecting tubule preparation made it ideal for studying the regulation of conductance of various ions in individual principal cells. When we measured the sodium conductance by subtracting the GT measured in the presence of amiloride from that in the absence of amiloride, as shown in Fig. 3A, the resultant measurements of conductance were not stable. A linear increase in the conductance persisted for at least 8 min after the application of amiloride. The Delta GT rose from 1.6 ± 0.3 to 3.5 ± 0.5 nS. This observation surprised us because it is generally accepted that the time course of amiloride blockage of sodium channels is very rapid and others have observed a time-dependent decrease in the amiloride-sensitive sodium conductance (32). The intercept between the control and amiloride lines (Fig. 2A) should represent the equilibrium potential for sodium. We expected a more positive value for this intercept (approximately +55 mV). In addition, for reasons we cannot explain, we observed a great deal of variability in the intercept value (data not shown). However, the intercept value continued to shift in the negative direction with time after the amiloride application (data not shown). This shift would be consistent with an amiloride-dependent decrease in a potassium conductance. Because the value in Fig. 2A is near 0 mV after exposure of the apical membrane to amiloride for 1 min, we presume that the secondary decrease in a potassium conductance has already begun. We believe the contribution of the nonselective cation channel would be small, with a relative abundance of 0.07 levels/seal under control conditions (45). Other investigators were unable to show the effect of 2 µM amiloride on these nonselective cation channels (45).

A time-dependent reduction in the activity of small-conductance potassium channels located in the basolateral membrane of the rat cortical collecting duct when amiloride is applied to the apical membrane has already been reported by Lu et al. (25). The decrease in potassium channel activity continued for several minutes after the application of amiloride (25), a time course similar to the amiloride-induced GT observed in this study. We were able to abolish the time-dependent increase in the amiloride-sensitive Delta GT (Fig. 3) when potassium channel blockers were present. These data are consistent with the failure of other investigators to observe secondary effects of amiloride in the presence of potassium channel blockers (32). The data suggest that the time-dependent increase in the amiloride-sensitive Delta GT observed in Fig. 3A was due to a secondary effect of amiloride on a potassium conductance. It should be noted that, even in the presence of potassium channel blockers, the equilibrium value for sodium determined from the intercept of the amiloride and control lines was highly variable (data not shown).

We decided to estimate GNa under conditions in which there is little or no potassium conductance. By clamping the cell at -100 mV, the potassium equilibrium potential, there should be little or no current due to potassium. At this voltage, the change in current flowing across the membrane is due to sodium because there is no driving force for potassium movement (35). As shown in Fig. 3A, we obtained the same initial conductance as before, 1.3 nS; however, GNa was now stable with time.

Although subtraction of the amiloride-insensitive conductance from the total cell conductance provides a reasonable estimate of GNa, especially if one measures the amiloride-insensitive conductance within 1 min of applying the drug, the presence of a time-dependent change in potassium conductance certainly reduces its accuracy. We believe that estimates of GNa determined by measuring the amiloride-inhibitable current with the cell clamped at -100 mV should yield more accurate values because they are stable with time and do not appear to be complicated by time-dependent changes in other ionic currents. We conclude that the best way to measure GNa is to measure the amiloride-sensitive current at or near the potassium equilibrium potential.

That a similar value of GNa can be calculated from our cell-attached patch-clamp data lends support to the notion that the GNa presented here is valid. From the data of Stanton et al. (38), we estimate the surface area of the apical membrane of principal cells of a similar salamander, Amphiuma means, to be 349 µm2. Given that the average number of active ENaC levels is 5.8 levels/patch (Table 2) and the estimate of the surface area of a single patch is 1.4 µm2 (44), the total number of active levels for the apical membrane should be 1,446 levels/cell. The whole cell sodium conductance is given by the following equation
G<SUB>Na</SUB> = <IT>g</IT><SUB>Na</SUB><IT> · N · P</IT><SUB>o</SUB>
where gNa is single-channel sodium conductance. Our values for gNa and Po were 4.6 pS and 0.21 (Table 2), respectively. The product of these values yields a GNa of 1.4 nS, a value similar to the normalized GNa of 1.2 nS.

It is also possible to calculate the rate of sodium transport of an initial collecting tubule from the single-cell amiloride-sensitive sodium current measured at -100 mV. The normalized sodium current of 120 pA (Table 1), when divided by Faraday's constant, yields a sodium flow rate of 124 × 10-17 mol · s-1 · cell-1. Using 60 s/min and assuming 300 principal cells per millimeter of tubule length, we compute a transport rate of 22.4 pmol · mm-1 · min-1. Stoner (39) reported a measured sodium transport rate for this tubule segment of 21.2 pmol · mm-1 · min-1. The agreement between these values suggests that the methods used to measure GNa are valid.

The second achievement of this study was to characterize the modulation of ENaC after short-term application of insulin in native renal tissue. It was also the first attempt to study the effect of insulin by measuring both whole cell GNa and then the single ENaC activity. In whole cell patch-clamp experiments, normalized GNa was 1.2 ± 0.3 nS in untreated tubules and 2.0 ± 0.4 nS in tubules pretreated with insulin for 10-60 min. Assuming that the increase in normalized GNa reflects a proportional increase in sodium transport, this represented an increase of 70% in sodium reabsorption at the single-cell level. Increases in net sodium transport of similar magnitude have been reported for the effect of insulin on frog skin (9, 36), toad bladder (5), and A6 cells (2, 6, 15, 27, 34).

Using cell attached patch-clamp methodology, we previously characterized, at the single-channel level, the 5-pS ENaC (42, 44). As reported for the rat collecting tubule (17, 30, 31), this was the only sodium channel we observed under control conditions. For a near doubling of the amiloride-sensitive current to occur after exposure of cells to insulin, there had to be an increase in the single-channel conductance, the number of active channels in the patch, the open probability, or a combination of the three. Cell-attached patch-clamp techniques were used to provide more detail regarding which of these changes in ENaC activity were responsible for the increase in sodium reabsorption.

There has been general agreement that insulin does not cause a significant change in single-channel conductance of ENaC (2, 6, 15, 27). Several investigators, using noise analysis on A6 cells, showed that insulin increased the number of active apical membrane channels (2, 6, 15). In contrast, Marunaka et al. (27), using cell-attached patch-clamp techniques on A6 cells, reported an increase in the open probability of ENaC in the presence of insulin. Our results in native renal tissue confirmed the observations of Marunaka et al. (27) and showed a statistically significant (P <=  0.01) increase in the open probability of ENaC in the presence of insulin. This increase was of similar magnitude to that observed for the whole cell sodium conductance. These observations, together with the lack of any measurable effect of insulin on the relative abundance of ENaC or single-channel conductance, suggest that the primary mechanism by which insulin increases GNa is to increase the fraction of time individual ENaC stay in the open state.

The physiological significance of the effect of insulin on sodium reabsorption has not been well defined. Some have suggested that the physiological importance of this hormonal effect resides in the need to increase sodium reabsorption to compensate for increased delivery of sodium to the distal nephron (20). Presumably, increased delivery of sodium is due to increases in glomerular filtration rate associated with eating a large meal. There is little experimental evidence to support this hypothesis. Another possibility is the need to secrete potassium associated with food intake (M. Halperin, personal communication). Elevated activity of ENaC would increase transepithelial voltage, enhancing potassium secretion. There is little experimental evidence to support either hypothesis.

On the other hand, many have suggested that the pathophysiological significance of the effect of insulin on sodium transport could be that hyperinsulinemia would lead to inappropriate sodium reabsorption, volume expansion, and hypertension, a common malady associated with people in a hyperinsulinemic state (3, 6, 10). The direct link between hyperinsulinemia and hypertension remains to be established.


    ACKNOWLEDGEMENTS

The authors are grateful to Drs. Peter Holohan, Mitchell Halperin, and Edward Solessio for careful reading of this manuscript and many helpful suggestions. Susan Viggiano gave excellent advice both in the planning of protocols and in manuscript preparation, and Susan Sutterer provided precise and meticulous technical assistance.


    FOOTNOTES

This work was supported by National Science Foundation Grant IBN9812314 and American Heart Association Predoctoral Fellowship Grant 0110110T.

Address for reprint requests and other correspondence: L. C. Stoner, Dept. of Neuroscience and Physiology, State Univ. of New York Upstate Medical Univ., 766 Irving Ave., Syracuse, New York 13210 (E-mail: ytallini{at}hotmail.com).

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.

June 26, 2002;10.1152/ajpcell.00606.2001

Received 20 December 2001; accepted in final form 12 June 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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Am J Physiol Cell Physiol 283(4):C1171-C1181
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