Differential effects of phorbol ester (PMA) on blocker-sensitive ENaCs of frog skin and A6 epithelia

Willem J. Els, Xuehong Liu, and Sandy I. Helman

Department of Molecular and Integrative Physiology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Activation of protein kinase C with phorbol 12-myristate 13-acetate (PMA) caused complex transient perturbations of amiloride-sensitive short-circuit Na+ currents (INa) in A6 epithelia and frog skins that were tissue and concentration dependent. A noninvasive channel blocker pulse method of noise analysis (18) was used to investigate how PMA caused time-dependent changes of apical membrane epithelial Na+ channel (ENaC) single-channel currents, channel open probabilities (Po), and channel densities (NT). In A6 epithelia, 5 and 50 nM PMA caused within 7 min concentration-dependent sustained decreases of Po (~55% below control, 50 nM) and rapid compensatory transient increases of NT within 7 min (~220% above control, 50 nM), resulting in either small transient increases of INa at 5 nM PMA or small biphasic decreases of INa at 50 nM PMA. In contrast to A6 epithelia, 50 and 500 nM PMA in frog skin caused after a delay of at least 10 min transient increases of NT to ~60-70% above control at 30-60 min. Unlike A6 epithelia, Po was increased ~15% above control within 7 min and remained within ±10-15% of control for the duration of the 2-h experiments. Despite differences in the time courses of secondary inhibition of transport in A6 epithelia and frog skin, the delayed downregulation of transport was due to time-dependent decreases of NT from their preelevated levels in both tissues. Whereas Po is decreased within minutes in A6 epithelia as measured by noise analysis or by patch clamp (8), the discrepancy in regulation of NT in A6 epithelia as measured by noise analysis and patch clamp is most likely explained by the inability of on-cell patches formed before treatment of tissues with PMA to respond to regulation of their channel densities.

sodium channels; protein kinase C; epithelial transport; noise analysis; electrophysiology; sodium channel blockers; sodium transport; tissue culture; cortical collecting ducts; kidney

    INTRODUCTION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

THE AMILORIDE-SENSITIVE Na+ channels in apical membranes of tight epithelia (ENaCs) are the main site for the regulation of transepithelial Na+ reabsorption in many vertebrates. The properties of ENaCs may be altered by a variety of biochemical reactions. Of these, phosphorylation represents an important regulatory mechanism in expression of Na+ channel activity by kinases. Subunits of the channel and/or related proteins may be phosphorylated by cAMP-dependent protein kinase (PKA) as well as by protein kinase C (PKC) (4, 26). Hence, activation of either of these signaling pathways may be involved in regulation of the gating kinetics of the channel, activation of membrane bound channels, and/or recruitment of channels by vesicle trafficking of channels to the apical membranes.

PKC is an integral component of the cell signaling machinery in epithelia (25, 29). PKC is a Ca2+- and phospholipid-dependent protein kinase that is physiologically activated by the second messenger diacylglycerol or pharmacologically by phorbol esters (2, 25). PKC has been implicated in a very broad spectrum of functions in the kidney and other salt-absorbing epithelia and is known to modulate water and Na+ transport across renal epithelia (5, 33). Although the cellular actions of PKC have not been completely elucidated, results from several studies have contributed to the hypothesis that the regulatory effects on Na+ transport by an ever-increasing number of hormones and other agents, such as prostaglandins and atrial natriuretic peptides in the collecting ducts, may be mediated in part by activation of PKC (Ref. 8 and references therein).

Defining the mechanisms of action of PKC on ENaCs has in part been complicated by the fact that its activation causes complex and apparently paradoxical responses of the macroscopic rates of Na+ transport. In epithelia of frog skin, activation of PKC causes stimulation of Na+ transport (6, 7, 23, 37), whereas in cultured renal A6 cells activation of PKC causes inhibition of transport (30, 36). Patch-clamp experiments have demonstrated that, in certain epithelia of renal origin, activation of PKC may cause a prompt inhibition of Na+ channel activity (or open-channel density) (13, 20, 21). It has been suggested that inhibition of channel activity is caused by PKC-mediated decreases of open probability (see Refs. 4 and 20 and references therein). However, as also alluded to above, activation of PKC with phorbol ester results in complex changes of Na+ transport that vary among tissues, including stimulation of transport that cannot be explained by decreases of channel open probability (Po). We were, accordingly, interested to know how apical membrane Na+ channels respond to perturbation of the PKC signal transduction pathway, especially when the transport responses are transitory and occur over periods of minutes to several hours. We were interested in knowing also why cell cultured A6 epithelia behave differently from frog skins in response to activation of PKC by phorbol ester in terms of the time-dependent changes of single-channel currents (iNa), Po, and channel densities (NT) that, together, account for time-dependent changes of the macroscopic rates of Na+ transport.

In addressing these questions, we made use of a noninvasive pulse inhibition method of blocker-induced noise analysis that allowed us to determine, within relatively short intervals of time and over several hours, modulations of iNa, NT, and Po that underlie the changes in rates of Na+ transport (18). It turned out that this approach was especially useful following activation of PKC with phorbol 12-myristate 13-acetate (PMA) when the responses of the short-circuit currents (Isc) were most complex and reflected complex time- and concentration-dependent compensatory changes of the iNa, Po, and NT.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Tissues. The A6 cells used in the present studies were obtained as a gift from Dr. Nancy Wills (Univ. of Texas Medical Branch, Galveston, TX). When grown to confluency on permeabilized supports in the absence of exogenous mineralocorticoid stimulation, the tissues exhibited baseline rates of Na+ transport comparable in value to those of frog skin (see RESULTS). After the cells were subcultured in tissue culture flasks, they were seeded onto Millicell HA tissue culture inserts (30 mm, 0.45 µm; Millipore, Bedford, MA). The culture medium was a mixture of DMEM (84-5022EC, GIBCO, Grand Island, NY) supplemented with 4 mM HEPES, 10% fetal bovine serum (HyClone, Logan, UT), and penicillin (25 U/ml) and streptomycin (25 µg/ml) (17-719R, BioWhittaker, Walkersville, MD). The cells were grown, and tissues were maintained at 28°C in a humidified incubator containing 1% CO2.

Isolated epithelia of frog skin, devoid of glands and connective tissue of the corium, were prepared from abdominal skins of R. pipiens pipiens (Kons Scientific, Germantown, WI) by collagenase digestion according to methods previously described (12).

Electrical measurements. The methods of study with blocker-induced noise analysis were identical to previous reports from our laboratory except for modification of the original blocker pulse protocol (17), which permits determination of iNa, Po, and NT during transient periods of Na+ transport (18). The theory and methods have been described elsewhere in detail (18), and the salient points are described briefly herein. After transfer of the A6 epithelia or isolated epithelia of frog skin to chambers designed for noise analysis, the tissues were short-circuited continuously with a low-noise voltage clamp for at least 1-2 h to allow the Isc to stabilize; 10 µM of the weak Na+ channel blocker, 6-chloro-3,5-diaminopyrazine-2-carboxamide (CDPC; Aldrich Chemical, Milwaukee, WI), was added to the apical perfusion solution, and the Isc was allowed to autoregulate back to the original baseline values of Isc (~30 min). After subtraction of 100 µM amiloride-insensitive currents measured at the ends of the experiments (IAmilsc), the blocker-sensitive currents in the presence of 10 µM CDPC (I10Na) were measured continuously. At intervals of 20 min during control periods and at ~7-min and 20-min intervals thereafter following exposure of the tissues to PMA, the CDPC concentration was increased from 10 to 30 µM for pulse intervals of 3 min (see Fig. 2). The Isc at 30 µM CDPC (I30Na) were measured ~30-40 s after the CDPC concentration was increased, at which times channels were redistributed at equilibrium between closed, open, and blocked states following blocker inhibition of the open channels (17, 18). The resulting fractional inhibitions of the Isc (I30/10Na triple-bond  I30Na/I10Na) were used in calculation of the Po as described previously; in Eq. 1, KB is the equilibrium constant for blocker interaction with the open state of the channel, B2 and B1 refer to blocker concentrations B, which for our studies are 30 and 10 µM (B30 and B10), respectively
<IT>P</IT><SUB>o</SUB> = <FENCE><FR><NU>1 − <IT>I</IT><SUP><IT> B</IT><SUB>2</SUB>/<IT>B</IT><SUB>1</SUB></SUP><SUB>Na</SUB></NU><DE><IT>B</IT><SUB>2</SUB> <IT>I</IT><SUP><IT> B</IT><SUB>2</SUB>/<IT>B</IT><SUB>1</SUB></SUP><SUB>Na</SUB> − <IT>B</IT><SUB>1</SUB></DE></FR></FENCE> <IT>K</IT><SUB><IT>B</IT></SUB> (1)
Blocker-induced current noise was measured at 10 µM CDPC just before the CDPC concentration was increased from 10 to 30 µM and at 30 µM CDPC following redistribution of the channels and measurements of the fractional inhibitions of blocker-sensitive Na+ currents (INa). Corner frequencies of the Lorentzians at 10 and 30 µM CDPC (f10c and f30c) and low-frequency plateaus (S100 and S300) were determined by nonlinear curve fitting of the spectra. Blocker on and off rates, kob and kbo, respectively, were determined from the slopes and intercepts of rate concentration plots (2pi fc = kobB + kbo). KB was calculated from the quotient kbo/kob at each time point of measurement and, hence, as a function of time during control and experimental periods.

Single-channel currents (iNa triple-bond  i10Na) were calculated in the usual way from the quotient [S100(2pi f10c)2]/[4I10NakobB10]. Open-channel densities (No) at 10 µM CDPC (N10o) were calculated as I10Na/i10Na, and No in the absence of blocker was calculated as N10o[1 + (PoB10/KB)]. The total pool of channels in closed and open states in the absence of blocker, NT, was calculated as No/Po.

Solutions and reagents. A6 epithelia, while short circuited in the chambers, were perfused continuously and symmetrically with the culture medium minus the fetal bovine serum and antibiotics, thereby assuring that the tissues were studied under essentially the same conditions under which they were cultured. Isolated epithelia of frog skin were perfused with a Ringer solution containing (in mM) 100 NaCl, 2.4 KHCO3, and 2.0 CaCl2 at a pH of ~8.1. Stock solutions of PMA and 4alpha -phorbol (Sigma Chemical, St. Louis, MO) were prepared by dissolving in ethanol and were stored at -20°C.

Statistical analysis. Data are expressed as means ± SE. Statistical analyses were performed with SigmaStat (Jandel Scientific Software, San Rafael, CA) using paired or unpaired t-tests when appropriate. P < 0.05 was considered significant.

    RESULTS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

The phorbol ester PMA is a potent activator of PKC in eukaryotic cells (2, 25). Like previous investigators (6, 20, 23, 36), we found that the responses of short-circuited tissues to PMA were mostly absent or markedly smaller and variable when applied from the basolateral surface of the tissues. In preliminary studies, we first examined the effects of PMA at various concentrations falling within the range commonly used for similar experiments. We elected to perform our experiments at two different concentrations, a low concentration at which we could still consistently observe an effect on the Isc and a 10-fold higher concentration, close to the maximum concentrations often used in other studies. Hence, with A6 epithelia, we tested the effects of PMA within the range of 1-100 nM and performed our experiments with 5 and 50 nM PMA. In experiments with isolated epithelia of frog skin, we used higher concentrations of 50 and 500 nM PMA, which were similar to concentrations used in other experiments on native epithelia to produce minimum and maximum effects on Na+ transport rates (31, 32). To test the specificity of PMA, we also determined that application of 2 µM of the inactive phorbol ester 4alpha -phorbol to the apical surfaces of cells had no significant effect on the Isc (data not shown). Addition of the solvent alone had no significant effect on the transport rate.

Because prolonged stimulation with PMA may lead to downregulation of PKC (2) and because we wanted to test for reversibility of the effects of PMA, we arbitrarily elected to remove PMA from the apical solutions at a time when the Isc had decreased to 85% of control values (Figs. 1 and 2). This was done regardless of the time it took to reach this value, which for A6 epithelia treated with 50 nM PMA was as short as 10 min or up to ~60 min in those A6 epithelia and frog skins exhibiting transient increases of transport (see Figs. 1 and 2).


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Fig. 1.   Short-circuit current (Isc) responses to apical membrane exposure of phorbol ester in A6 epithelia treated with either 5 nM (A) or 50 nM (B) phorbol 12-myristate 13-acetate (PMA). Shown are strip chart records in which Isc values have been normalized to zero time control values. PMA was removed from the apical solution when Isc had fallen to 85% of the zero time control value, regardless of the time at which this occurred. Isc responses during the first few minutes could appear delayed (as in A) before onset of the transient increase of Isc. In some experiments, the Isc fell initially before rising again toward and above the zero time control value. In other experiments (as shown in B), the Isc fell within minutes to an apparent plateau before the secondary decrease of Isc.


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Fig. 2.   Normalized strip chart recordings of Isc responses to apical membrane exposure of phorbol ester in frog skins treated with either 50 nM (A) or 500 nM (B) PMA. In contrast to A6 epithelia, all tissues exhibited small transient increases of Isc. At the intervals shown, apical solution 6-chloro-3,5-diaminopyrazine-2-carboxamide (CDPC) concentration was pulse increased from 10 to 30 µM for 3 min. Isc (µA/cm2) was recorded digitally at 10-s intervals as indicated in C from which the fractional inhibition of current was determined ~30-40 s after elevation of the CDPC concentration and before the delayed onset of the autoregulatory increase of transport. Current noise was measured for 2 min (60 by 2-s frames) just before CDPC concentration was increased to 30 µM and at 30 µM during the last 2 min of the pulse interval. It may be noted that corner frequencies are independent of Isc and so can be measured during the delayed autoregulatory increase of transport.

Effects of PMA on blocker-sensitive macroscopic rates of Na+ (INa). PMA induced transient responses of the macroscopic rates of Na+ transport that were both concentration and tissue dependent. An appreciation of the complexity of the responses of Isc most commonly produced by stimulation of PKC is provided by the illustrations shown in Fig. 1 for experiments done with A6 epithelia. For comparison, the Isc data were normalized to zero time control values to reflect the time-dependent changes as a percentage of control. At zero time, PMA was added to the apical solution. As shown in Fig. 1, PMA typically produced different concentration-dependent responses in A6 cells. After a delay of a few minutes, 5 nM PMA most often caused a rapid small initial stimulation of the Isc, which lasted for ~20 min (Fig. 1A). Occasionally, stimulation was preceded by a very small inhibition of the Isc. Maximum stimulation was usually reached within ~10-15 min. Irrespective of the time-dependent changes of Isc seen during the initial 10-20 min, the earlier transients were always followed by a slow secondary inhibition of the Isc to values substantially lower than controls within 1-2 h. At a higher concentration, 50 nM PMA consistently induced a biphasic inhibition of Na+ transport (Fig. 1B). Typically, the Isc would decrease within the first few minutes to an apparent plateau 10-20% below control levels within ~10 min. As with the lower concentration of PMA, this was followed by a slower secondary downregulation of the Isc over the next 60-90 min.

In contrast to A6 epithelia, epithelia of frog skin always responded to PMA at 5 nM (data not shown) or higher concentrations (50 and 500 nM), typically with an initial stimulation of the Isc (Fig. 2). PMA at 50 nM or a much higher concentration of 500 nM PMA produced a small transient stimulation of the Isc that persisted for 30-60 min. As with A6 epithelia, stimulation was followed by a sustained and delayed secondary inhibition of transport.

Figure 2 also shows the Isc responses to pulse elevations of CDPC concentration. Current noise spectra were measured at 10 µM CDPC and during the last 2 min of the 3-min pulse interval at 30 µM CDPC. With the measurements of fractional inhibition of current and those derived from the paired Lorentzians at each interval of measurement, we calculated iNa, No, Po, and NT as indicated in MATERIALS AND METHODS. It may be noted, as indicated in Fig. 2C, that the fractional inhibitions of current (I30/10Na) were derived from measurements of the Isc just before and after 30-40 s following elevation of CDPC concentration and before the delayed autoregulatory increase of transport that occurs in response to inhibition of apical membrane Na+ entry (1, 17, 18).

Mean Isc at times just before elevation of CDPC concentration are summarized in Fig. 3 for A6 epithelia and frog skins treated with PMA. It was clear from all experiments within the range of concentrations used in these studies that the effects of PMA on the Isc were not reversible. Despite the time of removal of PMA from the apical solution, the Isc continued to decrease and remained inhibited for several hours (data not shown).


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Fig. 3.   Summary of mean blocker-sensitive Isc (INa) before and after treatment with PMA of A6 epithelia (A) and frog skins (B). Data points correspond to the times at which CDPC concentration was elevated from 10 to 30 µM and represent the currents just before measurement of the fractional inhibition of transport by CDPC. Numbers in parentheses indicate no. of experiments.

Baseline values of INa, iNa, No, Po, and NT. The zero time control values just before treating the tissues with PMA are summarized in Table 1. The iNa averaged between 0.27 and 0.38 pA and Po averaged between 0.29 and 0.35 for intact A6 epithelia and frog skins, indicating as in previous studies from our own laboratory (3, 9, 10, 17, 18) and those of others (see Refs. 4, 8 and references therein) that ENaCs of these and other epithelia exhibit similar characteristics. At mean macroscopic rates of Na+ entry into the cells between 8.73 and 17.26 µA/cm2, No varied between 26.0 and 73.6 channels/100 µm2 and total functional channel density (NT) varied between 77 and 268 channels/100 µm2.

                              
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Table 1.   Zero time control values

Blocker rate coefficients. Zero time CDPC blocker on rate (kob) averaged 6.78 ± 0.31 (19) and 4.71 ± 0.21 (18) rad · s-1 · µM-1, whereas the kbo averaged 245 ± 5.9 (19) and 176 ± 5.6 (18) rad/s for A6 epithelia and frog skins, respectively. Mean KB averaged near 37 µM in both tissues (Fig. 4C), although both kob and kbo were significantly less in frog skins than in A6 epithelia (Fig. 4, A and B). It was readily apparent during the control periods that neither kob nor kbo was completely stable with time (Fig. 4, A and B). Due to time-dependent increases of kbo and decreases of kob, KB continued to increase at the same rates during the control periods and thereafter following treatment of the tissues at the lower concentrations of PMA. However, clear evidence of PMA-related effects on the rate coefficients were observed at the higher concentrations of 50 nM (A6 epithelia) and 500 nM PMA (frog skins). In frog skins, this was most evident 80-90 min after exposure of the tissues to PMA, despite removal of the PMA from the apical solution at a much earlier time.


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Fig. 4.   Summary of blocker on rate (kob; A) and blocker off rate (kbo; B) coefficients and KB (= kbo/kob; C) before and after apical membrane exposure of A6 epithelia and frog skins to PMA. Some standard error bars have been omitted for clarity. Zero time values were estimated by extrapolation of the control data to the time of addition of PMA to the apical solution.

PMA-related changes of iNa and No. PMA caused very small decreases (~8-9%) of iNa within 7 min in A6 epithelia (Fig. 5A) and no change of iNa in frog skins (Fig. 5B). At 67 min, iNa had decreased slowly and substantially to 78.0 ± 6.9% (50 nM) and 67.6 ± 6.0% (500 nM) of control in frog skins, unlike the responses in A6 in which the iNa remained near control values. Normalized time-dependent changes of iNa are summarized in Fig. 9.


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Fig. 5.   Summary of changes of single-channel currents (iNa) caused by PMA in A6 epithelia (A) and frog skins (B). Zero time values were estimated by extrapolation of the control data.

It was evident during the control periods that iNa was not stable in A6 epithelia or in the group of frog skins treated with 50 nM PMA. The apparent relative stability of the INa in these groups of tissues during the control periods (see Fig. 3) was due to time-dependent compensatory increases of iNa and decreases of the No (No = INa/iNa) (Fig. 6). Despite the falling control baseline values of No, or its relative absence in the group of frog skins treated with 500 nM PMA, PMA caused similar time-dependent quantitative and qualitative changes of No at 50 and 500 nM PMA. These changes of No in frog skin paralleled those of the INa (see Fig. 3).


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Fig. 6.   Summary of changes of open-channel densities (No) caused by PMA in A6 epithelia (A) and frog skins (B). Data were normalized to zero time values determined by extrapolation of the control data.

It is important to note in experiments of this kind that stability of the Isc during control periods does not necessarily mean that iNa, NT, and Po are stable with time. Despite the relative constancy of the Isc, it was clearly evident that substantial time-dependent decreases of No were accompanied by increases of iNa (and to a lesser extent increases of Po), leading to the appearance of relatively stable baselines of transport before treatment of the tissues with PMA.

After PMA treatment of A6 epithelia, 5 nM PMA caused a small transient increase of No (Fig. 6A), whereas No decreased continuously after treatment with 50 nM PMA. Because No is determined by the product PoNT (or NPo in patch-clamp experiments), it was of particular interest to know the comparative effects of PMA on Po and NT in frog skin and A6 epithelia, independent of the time-dependent changes of Po and NT that were occurring during the control periods.

PMA has differential effects on channel Po in A6 epithelia and frog skins. Despite similar zero time control values of Po that averaged near 0.3 in A6 epithelia and frog skins, the responses of Po to PMA were markedly different between tissues (Fig. 7). In A6 epithelia, PMA caused a rapid, concentration-dependent decrease of Po within 7 min to 76.0 ± 0.03 and 44.6 ± 0.04% of the zero time control in tissues treated with 5 and 50 nM PMA, respectively. The initial decreases of Po were transient, especially at 50 nM PMA with recovery to sustained values of Po below control for the duration of the experiments.


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Fig. 7.   Summary of changes of channel open probabilities (Po) caused by PMA in A6 epithelia (A) and frog skins (B). Zero time values were estimated by extrapolation of the control data.

In sharp contrast with A6 epithelia, PMA treatment of frog skins caused relatively small increases of Po within 7 min (with 50 nM PMA: 116.6 ± 3.3%; with 500 nM PMA: 113.6 ± 5.6% of control) (Fig. 7B). Thereafter, the Po continued to decrease for 40-60 min to values less than or near the zero time control values. During control periods, Po was relatively stable with time, tending to increase slightly on average but increasing significantly only in the group of frog skins treated with 50 nM PMA.

PMA-related changes of NT. Despite stable or falling baseline values of NT during control periods (Fig. 8), PMA caused transient and consistent increases of NT. Within 7 min, NT was increased significantly in A6 epithelia (with 5 nM: 135.4 ± 5.4%; with 50 nM: 220.1 ± 37.2% of the zero time control) but returned to control within 27 min when these tissues were treated with 50 nM PMA and within 47 min when treated with 5 nM PMA (Fig. 8A). Increases of NT were delayed by at least 7 min in frog skin, increasing thereafter (with 50 nM: 175.5 ± 27% at 47 min; with 500 nM: 163.0 ± 22.6% at 27 min) but falling again slowly toward and below the original zero time control values (Fig. 8B). Although PMA caused transient increases of NT in A6 epithelia and frog skin, the time-dependent changes were different between tissues both in magnitude of response as well as the times at which the channels were activated. The changes of NT were transient and especially long lived in frog skins compared with A6 epithelia.


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Fig. 8.   Summary of changes of channel densities (NT) caused by PMA in A6 epithelia (A) and frog skins (B). Zero time values were estimated by extrapolation of the control data.

To better appreciate the time-dependent interrelationships among iNa, Po, and NT that give rise to the macroscopic changes of Na+ transport, we have plotted, as shown in Fig. 9, the normalized values of iNa, Po, NT, and INa following treatment of the tissues with PMA.


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Fig. 9.   Fractional time-dependent changes of iNa, Po, and NT contributing to the macroscopic changes of Na+ transport (INa) in A6 epithelia (A: 5 nM PMA; B: 50 nM PMA) and frog skins (C: 50 nM PMA; D: 500 nM PMA). Data were normalized to zero time control values.

Regardless of tissue or the test concentration of PMA, the changes of transport rate are complicated and cannot be attributed to one factor alone. In A6 epithelia, although INa changed very little over 30 min, it became clear that within 7 min Po and NT changed in a compensatory and a concentration-dependent manner with relatively small decreases of iNa (Fig. 9, A and B). The secondary and delayed inhibition of transport could be attributed to decreases of NT at times when the Po was essentially constant, although depressed from control values.

The consistent initial stimulation of transport in frog skin could be attributed not only to a delayed increase of NT but also to an earlier initial stimulation (within 7 min) of Po (Fig. 9, C and D). Whereas Po appeared to remain near its control value, the secondary decreases of transport could be attributed to decreases of NT and to decreases of iNa with the long-term decreases of transport due principally to decreases of NT. If, indeed, PMA acts explicitly through activation of PKC in both A6 epithelia and frog skins, then clear differences emerge in our comparative studies of these tissues in understanding how phosphorylation by this kinase is involved in regulation of apical membrane Na+ entry into the cells of these tissues.

A comparison of results summarized in Fig. 9, C and D, indicates that near-maximum effects on transport in frog skin are elicited at 50 nM PMA, since the results with 500 nM PMA are essentially the same as those at 50 nM PMA.

    DISCUSSION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

There is considerable interest in understanding how phosphorylation of channels and associated proteins is involved in the regulation of Na+ transport in tight epithelia. Activation of PKC in renal epithelia has generally been associated with inhibition of Na+ transport (13, 15, 21, 24, 32, 35). Understanding the mechanism(s) involved is, however, complicated because of the time-dependent differences in responses of the macroscopic rates of Na+ transport among different tissues and within the same tissues. Inhibition is often preceded by stimulation of transport as measured by Isc, with the inhibitory phase of transport delayed by several tens of minutes as is consistently the case for frog skin and in A6 epithelia treated with low (5 nM) concentrations of PMA. From the point of view of macroscopic rates of transport, the initial changes of transport (within ±10-20%) caused by PMA activation of PKC are in particular not very impressive quantitatively. Such observations may in part explain the emphasis on the action of PKC as an inhibitor of Na+ transport, since the secondary changes of transport are clearly greater than the small stimulations, inhibitions, and oscillatory-like behavior of transport that precedes the consistent but delayed inhibition of transport. Because the early changes in transport are relatively small and easily dismissed in favor of the secondary downregulation of transport, we chose to examine the time courses of change of the iNa, Po, and NT during early and late phases of PKC activation that underlie the changes of macroscopic rates of transport in two tissues that have served as model tissues for distal tubular cortical collecting ducts (CCDs) of the kidney. We were particularly interested in the early phase, with the premise that the primary effects of PKC activation would be manifest within the first few minutes of tissue treatment with PMA.

Comparative effects of PMA on NT and Po. As found in both frog skin and A6 epithelia during the initial response, PMA caused an increase in density of the functional pool of channels (NT), although in frog skin the increase of NT was delayed by at least 10 min relative to the earlier onset of responses in A6 epithelia (Fig. 8). These increases of NT were responsible for the early time or initial stimulation of transport as observed in the macroscopic Isc records. The reason(s) for the delay in response in frog skin of NT to PMA is unknown and appears unrelated to diffusional or access delays to PKC, since the responses were essentially the same at 10-fold differences of concentration (50 and 500 nM PMA). Because the responses are the same at these concentrations of PMA, the mean 60-80% increases of NT at 30-60 min posttreatment are considered to be the maximal stimulation of NT in frog skin that can be elicited by PMA activation of PKC.

At the lower concentrations of 5 and 50 nM PMA, increases of NT in A6 epithelia occurred not only with a shorter delay (see Figs. 1 and 8) but with concentration-dependent increases of NT of near 40 and 120% above control (Fig. 9) at 5 and 50 nM PMA, respectively. These increases of NT in A6 epithelia, like those in frog skin, were not sustained but returned relatively slowly toward and below zero time control values. Examination of the data indicated clearly that the secondary downregulation of transport in both tissues could be attributed completely to decreases of NT albeit from elevated levels.

Such complex behavior of NT cannot be explained solely on the basis of PMA activation of PKC acting at a single site. PMA undoubtedly enters the cells and may exert its influences through PKC at a variety of sites that may be involved in regulation of the apical membrane bound pool of channels directly involved in mediating Na+ entry into the cells. There is presently no molecular basis known to us for critical or detailed speculation that permits a more in-depth understanding of either the direct and/or feedback mechanisms underlying the time-dependent changes of NT in either A6 epithelia or frog skin. If we assume that the initial increase of NT is the direct result of phosphorylation by PKC, then an increase of the apical membrane pool size of NT can occur through trafficking of channels from the cytosol to the apical membrane and/or through recruitment of inactive (nonfunctional) channels within the apical membrane to an active, functional state. Our data do not permit such a distinction to be made, as both mechanisms are possible, and thus will require further inquiry.

Regardless of the time of exposure of the tissues to PMA, its effects on transport and on the time course of change of NT and Po were not reversible by omission of PMA from the bathing solution. NT fell progressively with time after the initial increases of NT, thereby accounting for the secondary inhibition of transport as noted above. According to Heasley and Johnson (16), PMA activation of PKC is not readily reversible for several hours. In the face of irreversible effects within the time frame of our experiments, it is presently impossible to know which PKC-mediated effects are responsible for the secondary downregulation of transport.

Activation by PMA of NT can occur with or without changes of Po as was clearly evident comparatively between A6 epithelia and frog skin. In A6 epithelia, the initial increases of NT were accompanied by concentration-dependent decreases of Po, whereas in frog skin rather small initial increases of Po were followed by small oscillatory changes of Po that remained on average within about ±10-15% of the zero time control values. In this regard, frog skins behave quite differently from A6 epithelia in terms of PKC regulation of ENaC Po. We have considered the possibility that frog skins and A6 epithelia differ in the availability of Ca2+ for activation of PKC associated with regulation of Po. However, in more recent experiments with ionomycin, elevation of cytosolic Ca2+ with this ionophore caused an initial increase of Po in frog skin but a decrease of Po in A6 epithelia (19), similar to those reported here for PMA activation of PKC. Thus, regardless of whether PKC is activated by PMA or by increases of cytosolic Ca2+, regulation of Po by PKC is fundamentally different between frog skins and A6 epithelia that at least in part could be due to different isoforms of PKC in these tissues (5, 25). Such differences in behavior and regulation between two tissues that have and continue to be used as models of distal nephrons of the kidney underscore the need for caution in extrapolation of the findings in a particular tissue to understanding the behavior and regulation of transport in other tissues. Apparently, some behaviors may be conserved, whereas others like Po are not.

Our understanding of regulation of transport in CCDs in the rat and rabbit is no less confusing. On the one hand, Rouch et al. (31) reported that neither Ca2+ nor PKC activation changed Na+ transport in rat CCDs as measured by lumen-to-bath Na+ fluxes in isolated perfused tubules. Phorbol ester activation of PKC studied in preformed apical membrane on-cell patches of rat CCDs caused a mean 67% decrease of NPo or No within ~6 min (13), with an equally large expected inhibition of transport that would certainly be detectable in flux studies of Na+ transport like those carried out by Rouch et al. (31). Decreases of NPo (44.1%) have also been observed in patch-clamp records of rabbit CCDs within seconds of treating preformed on-cell patches of apical membranes with phorbol ester (21). Left unresolved in the patch-clamp studies of the rat and rabbit CCDs is the question of whether decreases of NPo are due to decreases of N and/or decreases of Po or perhaps compensatory changes of N and Po that give rise to net decreases of NPo. Regardless of change of Po and/or NT, decreases of NPo do not explain the absence of change of transport as measured in intact tubules. This problem is not unique to CCDs as will be indicated below.

Results from noise and patch clamp in A6 epithelia. According to Eaton et al. (8), "patch-clamp experiments in A6 cells show that the primary effect of activating apical membrane-bound PKC with phorbol esters and synthetic diacylglycerols is to reduce open probability of Na+ channels with little change in the number of channels." Within 5-15 min, Po was decreased markedly to near zero by 100 nM PMA treatment of preformed apical membrane on-cell patches (20). Such large changes of Po would cause large decreases of transport within minutes if NT is unchanged. To our knowledge of the literature and according to our own observations, large decreases of transport have never been observed within the time frame of minutes in A6 epithelia. Thus, according to analysis with patch clamp, equally large increases of NT corresponding to the magnitudes of decrease of Po must occur, but are not observed in patches, to account for the relatively small stimulation or inhibition of transport observed at these early times of response to PMA (see also below). It is, therefore, of particular interest to ask why data derived by patch clamp do not account for the macroscopic transport responses of CCDs and A6 epithelia.

With regard to regulation of Po by PMA activation of PKC, there is indeed agreement between our own studies with noise analysis and those of patch clamp by Ling and Eaton (20). PMA activation of PKC at least in A6 epithelia causes a prompt and substantial decrease of Po within several minutes.

Our observations with noise analysis with regard to regulation of NT are, however, not the same as those observed by patch clamp (20). Because we have measured transport directly in the same tissues in which we have evaluated not only Po but also iNa and NT, we are forced to conclude that in the face of substantial decreases of Po there must be relatively large increases of NT to account for the early stimulation or relatively small changes of transport measured during the early initial phase of the response to PMA activation of PKC.

Why do noise analysis and patch clamp give incompatible results with regard to PKC regulation of NT in the face of agreement on regulation of Po? We do not believe the problem rests with noise analysis, since unexplained disparities arise when the quantitative behavior of macroscopic rates of transport as measured by unquestionably reliable measurements of Isc or Na+ fluxes are taken into account in evaluation of mechanism. In this regard, attention to an important patch-clamp protocol dependency related specifically to studies of regulation of channels is warranted.

Marunaka and Eaton (22) showed that observations of large increases of NT caused by hormonal treatment (antidiuretic hormone, cAMP, forskolin) of A6 epithelia were completely prevented in on-cell patches formed before hormonal treatment. Patches formed after hormonal treatment showed large, severalfold increases of NT similar in magnitude to those observed by noise analysis (9), thereby accounting for the sustained stimulation of transport in nonpatched cells. Accordingly, the hormonal response, namely increase of NT, was completely prevented in patches formed before hormonal treatment of the tissues. This important protocol dependency has also been observed by others in studies of Cl- channels in pancreatic ducts (14) and in studies of Ca2+ channels in phorbol ester-treated Aplysia neurons (34). Consequently, in the face of such evidence, we favor the view that patch formation alters or interferes with the mechanism(s) responsible for increases of NT not only mediated by cAMP but also mediated by activation of PKC. Definitive experiments will be required to test this particular thesis in studies of regulation of functional NT and to evaluate to what extent, if any, patch formation alters or interferes with regulatory responses of Po in response to hormones and manipulations of other signaling pathways.

PKC and PKA as activators of NT. On quantitative grounds, PKC is a relatively poor activator of NT relative to cAMP/PKA activation of NT; in both frog skin and A6 epithelia NT is increased to sustained values 5- to 10-fold above controls as measured by noise analysis in frog skin (9) or in A6 by noise analysis and patch clamp (22, 27, 28). It may be noted that baseline values of Po and NT are essentially the same when measured by noise analysis and patch clamp of A6 epithelia, as evidenced in the results of the present experiments (see Table 1) and our previous experiments. Because the responses to maximal activation of PKC are so much smaller than can be achieved with cAMP activation of PKA, we cannot rule out the possibility, at least in part, that the changes of NT observed in our experiments with PMA are due to PKC activation of the same site(s) phosphorylated by PKA.

    ACKNOWLEDGEMENTS

We are most grateful to A. L. Helman for maintenance of our tissue culture facility, growth of the A6 epithelia, and assistance in the preparation of this manuscript. We are also grateful to Dr. Nancy Wills for providing the A6 cells used in the present study.

    FOOTNOTES

This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-30824 to S. I. Helman.

X. Liu is a doctoral student in the Dept. of Molecular and Integrative Physiology (University of Illinois at Urbana-Champaign). W. J. Els was on leave from the University of Cape Town Medical School while the experiments were carried out in Urbana.

A preliminary report of this work has been presented (11).

Present address of W. J. Els: Dept. of Anatomy and Cell Biology, University of Cape Town Medical School, Cape Town, South Africa.

Address for reprint requests: S. I. Helman, Dept. of Molecular and Integrative Physiology, Univ. of Illinois at Urbana-Champaign, 524 Burrill Hall, 407 South Goodwin, Urbana, Illinois 61801.

Received 15 September 1997; accepted in final form 23 March 1998.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

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