Phosphatase inhibitors increase the open probability of ENaC in A6 cells

A. Becchetti1, B. Malik1, G. Yue1, P. Duchatelle1, O. Al-Khalili1, T. R. Kleyman2, and D. C. Eaton1,2

1 Center for Cell and Molecular Signaling, Department of Physiology, Emory University School of Medicine, Atlanta, Georgia 30322; and 2 Department of Medicine, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15213


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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We studied the cellular phosphatase inhibitors okadaic acid (OKA), calyculin A, and microcystin on the epithelial sodium channel (ENaC) in A6 renal cells. OKA increased the amiloride-sensitive current after ~30 min with maximal stimulation at 1-2 h. Fluctuation analysis of cell-attached patches containing a large number of ENaC yielded power spectra with corner frequencies in untreated cells almost two times as large as in cells pretreated for 30 min with OKA, implying an increase in single channel open probability (Po) that doubled after OKA. Single channel analysis showed that, in cells pretreated with OKA, Po and mean open time approximately doubled. Two other phosphatase inhibitors, calyculin A and microcystin, had similar effects on Po and mean open time. An analog of OKA, okadaone, that does not inhibit phosphatases had no effect. Pretreatment with 10 nM OKA, which blocks protein phosphatase 2A (PP2A) but not PP1 in mammalian cells, had no effect even though both phosphatases are present in A6 cells. Several proteins were differentially phosphorylated after OKA, but ENaC subunit phosphorylation did not increase. We conclude that, in A6 cells, there is an OKA-sensitive phosphatase that suppresses ENaC activity by altering the phosphorylation of a regulatory molecule associated with the channel.

epithelial sodium channel; single channels; short-circuit current; protein kinases; phosphatases


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

LUMINAL SODIUM ENTRY THROUGH amiloride-sensitive, highly-selective sodium channels (ENaC) in the apical membrane of distal nephron epithelial cells is the rate-limiting step for hormonally stimulated sodium reabsorption. This implies that a thorough understanding of the mechanisms regulating these channels is fundamental to our understanding of total body sodium homeostasis and control of blood pressure.

Previous reports by us and others have shown that ENaC in A6 distal nephron cells are regulated by several different kinases. Ling and Eaton (36) have shown that protein kinase C (PKC)-mediated phosphorylation inhibits ENaC by reducing the open probability (Po) in native cells, but when expressed in oocytes activation of PKC apparently stimulates ENaC by increasing the number of channels in the surface membrane (56, 57). Studies by Matsumoto and co-workers (40) showed that tyrosine kinase-mediated phosphorylation stimulates channel activity by increasing the number of channels per unit area of membrane. Finally, arginine vasopressin promotes protein kinase A (PKA)-mediated phosphorylation of the channel or some closely associated protein (30). In patch-clamp experiments by Marunaka and Eaton (39), arginine vasopressin, acting through PKA, increases the apical membrane density of sodium channels without affecting Po. More recently, there have been several observations of the role of serum- and glucocorticoid-dependent kinase (SGK) in regulating ENaC activity (3, 11, 17, 42, 47). The serine/threonine kinase activity of SGK appears to determine the number of ENaC in the surface membrane of transporting cells. There are also reports of the effects of phosphorylation of ENaC, particularly the COOH-terminal domains of the beta - and gamma -subunits (13, 18, 46), although there is at least one report that suggests that, at least in native cells, ENaC subunits are not constitutively phosphorylated (59) and another which suggests that direct phosphorylation of channel subunits only changes activity under special conditions (13).

Changes in protein function by phosphorylation involves not only regulation of a kinase that phosphorylates but also regulation of a phosphatase that dephosphorylates the protein. Given the substantial evidence for regulation of sodium channels by protein kinase phosphorylation, we decided to explore the phosphatase-mediated dephosphorylation step. Several different phosphatases have been described and cloned from Xenopus laevis (but not A6 cells). Two isoforms of protein phosphatase 2A (PP2A) have been reported (16), and the sequence of X. laevis PP2B (calcineurin), 2C, and 1 are all available in GenBank (accession nos. AB037146, AF019569, and L17039, respectively).

In this study, we investigated the effect of phosphatase antagonists on amiloride-sensitive, highly selective sodium channels (ENaC) in A6 cells. Okadaic acid, a monocarboxylic polyether isolated from marine sponges, is known to inhibit several types of phosphatases (28). It is a very potent blocker of PP1 and PP2A, two of the major serine and threonine phosphatases present in mammalian cytoplasm (14). At least in mammalian cells, PP2A is completely inhibited by 1 nM okadaic acid, whereas the half-maximal inhibitory concentration for PP1 is ~15 nM. On the other hand, inhibition of PP2B is measurable only at concentrations >1 µM, and other phosphatases (including 2C) or kinases are not affected by okadaic acid (14). In addition, we examined the effect of two other phosphatase inhibitors, calyculin A and microcystin. Both are strong inhibitors of PP1 and PP2A, with little effect on other phosphatases (14). In mammalian cells, calyculin blocks both phosphatases with similar potency, but microcystin is more than 40 times as effective at blocking PP2A (IC50 = 0.04 nM) than PP1 (IC50 = 1.7 nM).

We show that pretreatment of A6 cells with 100 nM okadaic acid, 20 nM calyculin A, or 20 nM microcystin all produce an increase in amiloride-sensitive, short-circuit current (Isc). In addition, our results suggest that pretreatment of aldosterone-stimulated A6 cells with the phosphatase inhibitors affects the Po through an increase in the mean open time for single sodium channels in cell-attached patches, with a time course similar to the okadaic acid-induced increase in amiloride-sensitive Isc. On the other hand, the inhibitors do not seem to affect the number of channels present in the apical membrane. This study implies the existence of a phosphatase whose activity tonically reduces the Po of apical sodium channels, without affecting channel density. This is an interesting result because, based on previous examinations of PKC and PKA (36, 39), dephosphorylation of a serine/threonine phosphorylation site would be expected to increase Po (36) or reduce channel density (39), respectively. Thus it would appear that there is an additional kinase activity, not mediated by PKC or PKA, that is important in regulating sodium channel activity. This has been suggested in reports from several other laboratories (13, 56, 57).


    MATERIALS AND METHODS
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A6 cell culture preparation. For single channel experiments, A6 cells from the American Type Culture Collection (Rockville, MD) in passage 68 were prepared as described previously (36). Experiments were carried out on passages 70-80, with no discernible variation between cells from different passages. For transepithelial measurements, A6 cells (subclone 2F3 from Krahenbuhl and Rossier) from passages 96-97 were prepared following methods we have used previously (34, 35, 37). Cells were maintained in plastic tissue culture flasks (Corning) at 26°C in a humidified incubator with 4% CO2 in air. The culture medium was a mixture of Coon's F-12 medium (3 parts) and Leibovitz's L-15 medium (7 parts) modified for amphibian cells with 104 mM NaCl-25 mM NaHCO3, pH 7.4, with a final osmolarity of 240 mosmol/kgH2O. Besides these components, 10% (vol/vol) FBS (Irvine Scientific), 1% streptomycin, and 0.6% penicillin (Hazleton Biologics) were added. Cells grown on plastic tissue culture dishes were detached when confluent by exposing them to divalent-free (calcium and magnesium) medium containing 0.05% trypsin and 0.6 mM EDTA (Irvine Scientific). The cells were then rinsed, centrifuged, repeatedly resuspended, and finally replated. When used for patch-clamp experiments, A6 cells were replated at confluent density on collagen-coated CM permeable filters (Millipore) attached to the bottom of small Lucite disks, and the disks were suspended in 35-mm petri plates, as previously described (36). This sided preparation forms a polarized monolayer with the apical surface oriented upward and net sodium transport moving from the apical to basolateral surface. The bathing medium was supplemented with 1.5 µM aldosterone and 10% FBS. Every 2 days, the cells were fed with fresh medium, and patch-clamp experiments were performed 10 days after replating.

Single channel recordings and data analysis. Before the forming of patches, the apical cell surface was washed carefully several times with our standard extracellular solution containing (in mM) 95 NaCl, 3.4 KCl, 0.8 CaCl2, 0.8 MgCl2, and 10 HEPES (Sigma), and pH was adjusted to 7.4 with a small amount of NaOH. Patch pipettes contained the same solution. Experiments were only performed at room temperature (22-23°C) within 45-60 min of removing the A6 cells from the incubator. Patch pipettes with a tip diameter <= 1 µm were fabricated from WPI TW150 glass (New Haven, CT) and fire-polished following the procedure of Hamill et al. (27). Single channel currents from cell-attached patches were measured with an Axopatch 1-B current-voltage (I-V) clamp amplifier (Axon Instruments, Burlingame, CA), low-pass filtered at 1 KHz, recorded on a digital video recorder (Sony), and then digitized at two times the corner frequency (fc) using a Scientific Solutions analog-digital converter and IBM PC computer equipped with Axotape software (Axon Instruments). The data were subsequently transferred to a Vax computer (Digital Equipment) for single channel analysis.

Data records were low-pass filtered at 100 Hz using a software Gaussian filter. Events were detected by setting the threshold level at 50% of the estimated single channel current amplitude. We used programs that closely follow the strategy of Colquhoun and Sigworth (15), as previously described (36, 54).

We often used the product of the number of channels (N) times the open probability (NPo) as a measure of channel activity within a patch. This product can be calculated from single channel records without making any assumptions about the total number of channels in a patch or the Po of a single channel
NP<SUB>o</SUB> = <LIM><OP>∑</OP><LL><IT>i</IT> = 0</LL><UL>N<SUB>A</SUB></UL></LIM> <FR><NU><IT>i t<SUB>i</SUB></IT></NU><DE><IT>T</IT></DE></FR>
where T is the total recording time, NA is the apparent number of channels within the patch determined as the highest observable current level, i, is the number of channels open, and ti is the time during which i channels are open. If channels open independently of one another and the exact number of channels in a patch is known, then the Po of a single channel can be calculated by dividing NPo by the number of channels in a patch.

The mean open time (topen) of N channels can be calculated as follows
t<SUB>open</SUB> = <FR><NU><IT>T NP</IT><SUB>o</SUB></NU><DE><IT>n/</IT>2</DE></FR> = <FR><NU><LIM><OP>∑</OP><LL><IT>i = </IT>1</LL><UL><IT>N</IT><SUB>A</SUB></UL></LIM><IT>it<SUB>i</SUB></IT></NU><DE><IT>n/</IT>2</DE></FR>
where n is the total number of transitions between states during the total recording period T, and the other parameters are the same as in Eq. 1. This measure provides an easy way to distinguish whether experimental manipulations (i.e., okadaic acid) modify Po by affecting the channel's open states or closed states. However, this measure should not be confused with the residency time in a specific kinetic state (often referred to as the mean time for a particular state). The mean open time is merely a numerical average of all open times regardless of the kinetic state.

Fluctuation analysis. Noise analysis of sodium channels in intact epithelia generally requires the addition of a channel blocker like amiloride or CDPC (6-chloro-3,5-diamino-pyrazine-2-carboxamide) (see, e.g., Refs. 5, 9, 22, 29) to induce an fc. Fluctuation measurements on single ENaCs reveal a spontaneous corner without the need for addition of pharmacological blockers. This is possible, not because of special properties of single channel patches but rather because of limitations of recording noise fluctuations in intact epithelia. As one of us has discussed previously (20), there are two problems of recording spontaneous ENaC fluctuations from intact epithelial tissue. The first is the exceptionally low fc of ENaC (because of the long mean open and closed times, the spontaneous fc is expected to be ~0.01-0.1 Hz). This means that the preparation has to be stable with no change in transport for a very long period of time (15-30 min), which is often difficult to achieve. The second problem exacerbates the first: whole tissue preparations have a 1/f component to the noise in addition to the 1/f2 Lorentzian component. Because this component is large at low frequencies, more samples must be acquired at low frequency to resolve the Lorentzian, making the stability problem more serious. Single channel recordings have no 1/f component and can be stable for relatively long periods. Also, records with similar products of N times Po can be appended to achieve longer records.

To determine the low-frequency corner from our single channel records, a fast-Fourier transform was applied to 28 min of digitized recordings from eight patches obtained on untreated A6 cell monolayers. The same procedure was applied to 16 min of recordings from four patches obtained on cells pretreated with okadaic acid for 30 min and from 19 min of recording from four patches on cells pretreated for 1 h. The low-frequency plateau [S(0)] and the fc were determined by fitting the power spectra to the formula for a single Lorentzian function
<IT>S</IT> (<IT>f</IT>)<IT>=</IT><FR><NU><IT>S</IT>(0)</NU><DE>[1<IT>+</IT>(<IT>f/ f<SUB>c</SUB></IT>)<SUP>2</SUP>]</DE></FR>
where S(f) is the power at frequency f. Eaton et al. (21) have previously reviewed the application of single channel fluctuation analysis techniques to renal ENaCs, but briefly the methods we applied depend on several facts. First, the relative variance, sigma <UP><SUB><IT>n</IT></SUB><SUP>2</SUP></UP>, of n channels is
&sfgr;<SUP>2</SUP><SUB>n</SUB>=nP<SUB>o</SUB> (1−P<SUB>o</SUB>)
where average Po is the open probability of the n channels. The variance in the patch current is
&sfgr;<SUP>2</SUP><SUB>i</SUB>=i<SUP>2</SUP>nP<SUB>o</SUB> (1−P<SUB>o</SUB>)
where i is the current of a single channel. The mean patch current, I, is
<I>=inP<SUB>o</SUB>
Combining these expressions gives
&sfgr;<SUP>2</SUP><SUB>i</SUB>=<I>i (1−P<SUB>o</SUB>)
The power spectra are related to the current variance, since the integral of the power spectrum Lorentzian function is equal to the variance
&sfgr;<SUP>2</SUP><SUB>i</SUB>=<FR><NU>S(0)&pgr; <IT>f</IT><SUB>c</SUB></NU><DE>2</DE></FR>
where S(0) and fc are the low-frequency plateau and fc defined above. If the unit current and the mean current are known or can be calculated, then combining the last two expressions allows one to calculate Po as
P<SUB>o</SUB> = 1 − <FR><NU><IT>S</IT>(0)<IT> &pgr; f</IT><SUB>c</SUB></NU><DE>2 < <IT>I>i</IT></DE></FR>
The number of channels in the patch, N, can be calculated since
N=<FR><NU><I></NU><DE>i P<SUB>o</SUB></DE></FR>
and finally, the individual forward and reverse rate constants can be calculated as
k<SUB>−1</SUB> = <FR><NU><IT>S</IT>(0)<IT> &pgr;</IT><SUP>2</SUP><IT> f</IT><SUP>2</SUP><SUB>c</SUB></NU><DE><<IT>I>i</IT></DE></FR>
and
k<SUB>I</SUB> = <FR><NU><IT>S</IT>(0)<IT> &pgr;</IT><SUP>2</SUP><IT> f</IT><SUP>2</SUP><SUB><IT>c</IT></SUB></NU><DE><IT><I>i</IT></DE></FR><IT>−</IT><FR><NU>2<IT>&pgr;</IT></NU><DE><IT>f</IT><SUB>c</SUB></DE></FR>
One difficulty with using power spectrum analysis is that it always requires additional information over and above what can be obtained from a single channel measurement. In our case, we used the unit current and the mean current, both of which are relatively easy to obtain from the single channel measurements if there is at least one event with all channels closed within the record (virtually always true if the patch is maintained for a long enough period of time). In a tight-seal preparation with no applied potential, there is little if any measurable leakage current. Therefore, mean current and current variance can be obtained by setting the lowest absolute value to zero and averaging all of the data points. In other types of fluctuation measurements, other methods are used to provide additional information (e.g., altering the concentration of an agent that produces blocker-induced fluctuations in proportion to the blocker concentration). Another problem is one of measurement. Most of the values are derived from several measured quantities, which means that, oftentimes, the error in the final determination of these values is quite large.

Pretreatment of A6 cells with phosphatase inhibitors. For pretreatment experiments, phosphatase inhibitors (LC Services or Calbiochem) were added to both the basolateral and apical bathing medium, achieving a final concentration of about 10 times the IC50 for the phosphatase of interest. Cells were then incubated for 30, 60, or 120 min at 26°C in a humidified incubator with 4% CO2 in air before cell-attached patch experiments were performed.

Acute perfusion of A6 cells with okadaic acid. Cell-attached patches were obtained on untreated A6 cell monolayers, and baseline channel activity was measured. After the patches were established (5 min), the apical solution was replaced with standard extracellular solution containing 100 nM okadaic acid. Perfusions were accomplished using a 23-gauge hypodermic needle positioned in the apical bath chamber of the Lucite disk and connected to a peristaltic pump (Buchler) via thin silicon tubes. A perfusion rate of 1 ml/min was sufficient to fill the cell chamber in ~60-90 s without disturbing the gigaohm patch seal. The pump was then stopped. For acute perfusion experiments, we always chose patches that initially contained a low number of channels per patch (N <=  4). In this way, even a short recording time (4-5 min) before the beginning of perfusion was sufficient to obtain a good estimate of the number of channels in the patch. Marunaka and Eaton (39) have previously reviewed the conditions under which highly selective sodium channel number and Po can be accurately determined using patch-clamp methods.

Isc recording. For transepithelial electrical parameters, A6 cells were seeded on type I collagen-coated nitrocellulose filters (Costar) at a density of 1 × 106 cells/cm2. Mature monolayers were selected for study 7-20 days after seeding. A6 cells were placed in serum-free medium for 40 h immediately before study with the serum-free medium supplemented with 300 nM aldosterone for the last 16 h (overnight). Measurements were made on confluent A6 monolayers using an Ussing chamber specifically designed to hold the nitrocellulose filters (4.7 cm2). The apical and basolateral bathing solutions had the same composition as the standard extracellular solution used for patch-clamp experiments. Baseline Isc and potential difference measurements were initially made, and then okadaic acid was added to the extracellular bath. Amiloride (10 µM) was added to the apical compartment to measure the amiloride-sensitive component of Isc. Exposure of A6 cells to vehicle alone produced no effect on baseline transepithelial parameters.

Okadaic acid-induced phosphorylation of cellular proteins and sodium channels. Okadaic acid should promote protein phosphorylation and might promote phosphorylation of sodium channel subunits. To examine these questions, we examined total cellular phosphorylation and phosphorylation of the alpha -, beta -, and gamma -subunits of the ENaC. A6 cells were deprived of serum overnight. The cells were washed two times with phosphate-free DMEM adjusted to amphibian osmolarity and supplemented with 5 mM glutamine and 1.5 µM aldosterone. After being washed, the cells were incubated in phosphate-free DMEM for 2 h before replacement of the media with fresh phosphate-free DMEM and incubation for another 2 h. Finally, the media was replaced with new phosphate-free DMEM containing a total of 5 mCi NaH233PO4 or NaH232PO4 overnight. The next morning, okadaic acid at a final concentration of 100 nM along with phosphate-free DMEM plus 33PO4 or 32PO4 was added to one group of cells while only medium was added to the other, and the cells were incubated overnight. 33P was used in experiments to examine all cellular proteins with the hope that its lower energy emissions would produce narrower bands when imaged: a hope that was not fully realized (see Fig. 11). The next day, the cells were chilled on ice and scraped in a solution containing (in mM) 100 KCl, 1 EGTA, and 10 KH2PO4 adjusted to pH 7.4 with KOH and containing protease inhibitors (100 µM antipain, 100 µM leupeptin, 100 µM N-tosyl-L-lysyl chloromethyl ketone, 100 µM N-tosyl-L-phenyl chloromethyl ketone, and 1 mM phenylmethylsulfonyl fluoride). The cells were centrifuged and lysed in 400 µl of 50 mM Tris buffer by douncing 40 times. The cells were centrifuged for 10 min in a tabletop centrifuge, and the precipitate was dounced again in 400 µl of 50 mM Tris buffer and centrifuged for 10 min. The combined supernatant was precipitated with 10× volume of cold acetone for 15 min at 4°C and then centrifuged in a tabletop centrifuge for 10 min. The precipitate was dissolved in 5× TBS. Part of the lysate was reserved to examine total cellular phosphorylation. Preimmune serum (20 µl) was added to the remaining lysate and incubated at room temperature for 1 h. Protein A beads (220 µl) were added for 30 min and centrifuged two times for 10 min at 13,000 revolutions/min. Polyclonal anti-alpha -, anti-beta , or anti-gamma X. laevis ENaC (20 µl) antibodies were added to one-third of the lysate and incubated overnight. The antibodies have been characterized in previous work (32, 33, 35, 38, 41, 44, 51). The next day, 400 µl of protein A beads were added and incubated for 30 min before centrifuging two times for 10 min at 13,000 revolutions/min. The precipitate was washed three times with 150 mM NaCl, 5 mM EGTA, 1% (vol/vol) Triton X-100, and 50 mM Tris · HCl (pH 7.4) and washed two times with 50 mM EGTA, 0.4% (wt/vol) sodium deoxycholate, 1% (vol/vol) Nonidet P-40, and 10 mM Tris · Cl (pH 7.4). The precipitate was boiled with sample buffer for 10 min and resolved on a 7.5% SDS-PAGE, and the phosphorylation was determined by quantitation of the gel on a PhosphorImager.

PCR cloning of A6 cell phosphatases. Phosphatase clones were amplified by PCR from A6 cDNA prepared by using a Marathon cDNA amplification kit (Clontech). The PCR primers were based on consenus sequences for X. laevis phosphatases obtained from GenBank (accession nos. L17039, X62114, and Z50852). For PP1 phosphatases, we used AAGAGAATGAGATTCGAGGGC for the forward primer and ACGGGTCTGCTTGCGTTAGG for the reverse primer. For PP2A phosphatases, we used the degenerate sequences GTGGATCGAGCAGCTGAAYGA (Y = T,C) for the forward primer and CCAAAGTAAGSCCATTAGCATGG (S = C,G) for the reverse primer. The PCR reaction conditions were 30 s at 94°C and 30 s at 60°C followed by a 2-min extension at 68°C for 30 cycles. Products of the expected size were subcloned into pGEM vector, and the nucleotide sequence of the PCR product was determined by Sanger dideoxynucleotide sequence analysis.

Statistics. Unless otherwise stated, data are presented as means ± SE. Paired or unpaired Student's t-test or ANOVA with a Tukey posttest was used as appropriate to test for significance.


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A6 cells contain PP1 and PP2A isoforms. Using PCR primers to conserved regions of PP1 and PP2A that should amplify any of the respective phosphatase isoforms, we were able to isolate and clone three phosphatase isoforms from A6 cells: one for PP1 (PP1gamma 1) and two for PP2A (PP2Aalpha and PP2Abeta ). The sequence of these clones was identical, at an amino acid level, to the previously reported sequences for phosphatases from other X. laevis tissues (16, 31, 55). We did not attempt to clone other phosphatases, since none of the agents we used will inhibit other categories of phosphatase.

Pretreatment with okadaic acid increases Isc. The amiloride-sensitive component of Isc was measured hourly (for up to 4 h) from aldosterone-stimulated A6 cell monolayers in either the absence or presence of 100 nM okadaic acid (5 monolayers for each condition) in the extracellular bath (Fig. 1). Exposure to okadaic acid increased the transepithelial Isc, beginning ~1 h after addition of the phosphatase inhibitor and peaking to ~1.5 times baseline Isc at ~2 h after addition. In contrast, the stimulatory effect of okadaic acid was quite small in A6 cells grown in the absence of aldosterone (data not shown). Lower doses of okadaic acid had no effect on transepithelial electrical parameters (data not shown).


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Fig. 1.   Okadaic acid increases short-circuit sodium current (Isc) in A6 cells. Amiloride-sensitive Isc was recorded from A6 cells monolayers in either the absence (, n = 6) or in the presence of 100 nM okadaic acid (black-down-triangle , n = 6). Isc is expressed as a fraction of the initial baseline Isc value in untreated cells (a value of 8.3 ± 0.91 µA/cm2). All values are means ± SE.

As noted earlier, Isc measurements alone cannot, in general, differentiate among the relative contributions of single channel conductance, Po, or channel density to changes in total apical sodium current. Therefore, we proceeded with cell-attached patch-clamp experiments to obtain more information about the mechanism underlying the effects of okadaic acid at the level of single sodium channels. In apical, cell-attached patches on aldosterone-stimulated A6 cells, we examined only the amiloride-sensitive, 4-pS, highly selective sodium channel that is responsible for arginine vasopressin- and aldosterone-stimulated sodium reabsorption in both A6 and mammalian (rat, rabbit) distal nephron cells (24).

For our analysis, we tried to examine patches that contained only one channel. However, there were many patches that have more than one channel. For some of these (N <=  4), we could calculate NPo and make good estimates of N and Po using conventional methods of analyzing channel properties; but, in other cases (N > 4), the number of channels in patches was too large for such methods to be applied with accuracy. Therefore, we applied fluctuation methods to these patches. Fluctuation measurements have the advantage of allowing an examination of a relatively large number of channels but has the disadvantage of requiring additional information about channel properties, in our case the unit conductance and mean patch current.

Pretreatment with okadaic acid decreases the fc of power spectra. We applied fluctuation analysis methods to the single channel records of cell-attached patches that contained a large number of channels per patch (N > 4). Figure 2 shows typical current records from two such patches, one untreated and one pretreated for 1 h with 100 nM okadaic acid. These and other similar records were used to generate power spectra and amplitude histograms from the same current records. The amplitude histograms indicate that both patches contain eight or more channels. Figure 2, bottom, shows the power spectra calculated from the same two current records. The power spectra from a number of similar patches (all with no potential applied to the pipette) were used to calculate the kinetic characteristics for a large number of sodium channels (see MATERIALS AND METHODS). We found that the fc of power spectra derived from control cells (0.35 ± 0.051; n = 8) were significantly higher than those from okadaic acid-pretreated cells (0.22 ± 0.0070; n = 4 after 30 min, and 0.27 ± 0.0090; n = 4 after 1 h), suggesting that phosphatase inhibition produced a change in the Po of single sodium channels. Consideration of the mean current and unit current indicates that the Po increases (Table 1). On the other hand, although the estimate of apical membrane channel density from power spectra was not very precise, our analysis suggested that, if channel number was increased by okadaic acid, the increase must have been small (<25%).


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Fig. 2.   Okadaic acid reduces the corner frequency of power spectra obtained from cell-attached patches with large numbers of channels. Left: data from a patch on an untreated cell. Right: a similar patch except that the cell was pretreated for 1 h with 100 nM okadaic acid. Downward transitions are inward current associated with channel openings. Top: current records in which it is difficult to observe discrete channel transitions. c, Closed state. Middle: amplitude histograms produced from the current records indicating that there are at least 8 channels in each patch. In the histogram from an okadaic acid-treated patch, there is a shift leftward toward occupancy of states with a larger number of channels open. Bottom: power spectra for the patches. The corner frequency of the okadaic acid-treated patch is shifted slightly to a lower frequency.


                              
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Table 1.   Summary of the effect of phosphatase inhibitor pretreatment on sodium channels of A6 cells

Pretreatment with okadaic acid increases NPo for sodium channels. To further investigate the effects of okadaic acid on individual sodium channel kinetics, analysis of single channel recordings was also performed on the cell-attached patches that contained a small number of channels per patch (N <=  4). A representative example of baseline sodium channel activity in a cell-attached patch from an untreated A6 cell is shown in Fig. 3, top left. Current traces represent ~1.5 min of continuous recording at resting membrane potential, with downward transitions representing inward sodium current. The amplitude histogram generated from this recording shows that this patch membrane contained three active sodium channels, whose Po was ~0.36 (Fig. 3, bottom left). This experiment can be compared with a representative cell-attached recording from an A6 cell after 1 h of pretreatment with 100 nM okadaic acid (Fig. 3, top right). Notice that the closed level, representing zero channel activity in the patch, was very rarely reached after phosphatase inhibition. From the amplitude histogram, this recording also contained at least three active channels, whose Po was ~0.58 (Fig. 3, bottom right). Table 1 summarizes our measurements from patches with multiple channels on untreated cells and cells pretreated with okadaic acid. Okadaic acid pretreatment for 1 h increased mean NPo (1.7 ± 0.46; n = 15) compared with controls (0.74 ± 0.12; n = 25).


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Fig. 3.   Effect of okadaic acid on a patch containing a small number of sodium channels. Left: sodium channel activity from a cell-attached patch formed on an untreated cell. Right: activity from a cell-attached patch formed on a cell after 1-h pretreatment with 100 nM okadaic acid. Top: current traces at resting membrane potential (-VP = 0 mV) from cell-attached patches containing at least three channels. Horizontal lines to left of each trace show closed level (c) with inward current downward. Currents were software filtered at 100 Hz. Bottom: amplitude histograms obtained from recordings at the top.

Pretreatment with okadaic acid increases Po without affecting channel density. For the patches containing a small number of channels (N <=  4) and a Po >0.1 and <0.9, N could be estimated with good accuracy, even for short recording periods (4-5 min), making it possible to calculate Po (see MATERIALS AND METHODS). The mean Po value measured from control patches was 0.29 ± 0.027 (n = 25), whereas the Po after 1 h of pretreatment with okadaic acid increased by ~40% (0.38 ± 0.053; n = 15). Therefore, this increase in sodium channel Po appeared to be sufficient to fully account for the 40-50% increase in amiloride-sensitive Isc previously observed after okadaic acid exposure. Consistent with our fluctuation analysis results, the number of sodium channels per patch from cells pretreated with okadaic acid (3.7 ± 0.93; n = 15) was not significantly different from that measured in the controls (3.1 ± 0.27; n = 25).

Pretreatment with okadaic acid increases the mean time that channels are open. A change in Po could be because of a change in either the mean open or closed time of apical sodium channels. Table 1 shows that 1 h of pretreatment with okadaic acid increased the mean time open (see MATERIALS AND METHODS) of sodium channels from 92 ± 18 to 170 ± 28 ms. The average open time for multiple channels is not equivalent to the mean residency time in any specific channel state (which requires information about the correct kinetic model for channel transitions) but is the average open time for all open states. The increase in average open time (~70%) produced by okadaic acid seemed sufficient to account for the observed changes in Po. This result suggested that the activity of an okadaic acid-sensitive phosphatase decreases the fraction of time during which the sodium channels are open.

However, to obtain a more accurate measure of the effect of okadaic acid on channel kinetics, we examined the properties of patches that contained only one sodium channel. These patches allowed us to determine mean open and closed times from the interval histograms for open and closed durations, respectively. Data from two such patches are shown in Fig. 4, top, where representative currents from an untreated patch (left) and from a patch treated with 100 nM okadaic acid for 1 h (right) are shown. The amplitude histograms (Fig. 4, bottom) show that there is only one channel in the patch. The interval histograms for the single channel events shown in Fig. 4 are shown in Fig. 5 [untreated (left) and okadaic acid treated (right); open (top) and closed (bottom) intervals]. For these histograms, we ignored a population of fast events (<100 ms) and measured only the duration of the state with a mean duration of the order of seconds. Table 1 summarizes our measurements from patches with single channels on untreated cells and cells pretreated with okadaic acid. Okadaic acid pretreatment for 1 h increased mean Po (0.54 ± 0.034; n = 5) compared with controls (0.31 ± 0.038; n = 11) ~1.7-fold, consistent with all of our previous results. Okadaic acid pretreatment for 1 h increased mean open time from 1.4 ± 0.12 to 2.7 ± 0.27 s, whereas mean closed time did not change significantly (2.3 ± 0.35 to 2.3 ± 0.16 s).


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Fig. 4.   Effect of okadaic acid on a cell-attached patch containing a single sodium channel. Left: sodium channel activity from a patch on an untreated cell. Right: activity from a cell-attached patch formed on a cell after 1 h of pretreatment with 100 nM okadaic acid. Top: current traces at resting membrane potential (-Vp = 0 mV) from a cell-attached patch containing only one channel. Horizontal lines to left of each trace show closed level (c) with inward current downward. Data were software filtered at 100 Hz. Bottom: amplitude histograms obtained from recordings at the top.



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Fig. 5.   Okadaic acid increases the mean open time of sodium channels. Interval histograms were constructed from 10 min of data from single channel patches. A: open interval histograms. B: closed interval histograms. Left: cell not pretreated. Right: cell pretreated for 1 h with 100 nM okadaic acid. Only the mean open time increases after treatment with okadaic acid.

Relative frequency of observation of patches with differing number of channels is not altered by okadaic acid. We used a variety of analysis methods to determine that the Po of channels was increased by okadaic acid, but we wished to assure ourselves that okadaic acid (or the other phosphatase inhibitors) was not altering channel density (N). Therefore, we tabulated the number of channels per patch for all the patches with channels used in this study (48 untreated, 33 treated with okadaic acid, and 10 treated with other phosphatase inhibitors). The probability of observing different numbers of channels in a patch is shown in Fig. 6 for untreated and phosphatase inhibitor-treated cells. There is no statistically significant difference between the distributions, although we cannot rule out a very small increase in N after inhibitor treatment. This distribution does not take into consideration the fact that 34% of the seals we formed had no detectable channels within the patch (and also did not appear to depend on treatment).


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Fig. 6.   Relative frequency of observation of patches with differing number of channels. The no. of channels per patch was tabulated for all patches with channels used in this study (48 untreated, 33 treated with okadaic acid, and 10 treated with other phosphatase inhibitors). The probability of observing different numbers of channels in a patch is shown for untreated (filled bars) and phosphatase inhibitor-treated (open bars) cells. There is no statistically significant difference between the distributions, although we cannot rule out a very small increase in the no. of channels per patch after inhibitor treatment. This distribution does not take into consideration the fact that 34% of the seals we formed had no detectable channels within the patch (and also did not appear to depend upon treatment).

Two other phosphatase inhibitors alter channel activity like okadaic acid. When cells were pretreated with either 20 nM calyculin A or microcystin, channel Po increased, with most if not all of the increase because of an increase in channel open time. Calyculin A pretreatment for 1 h increased mean Po (0.47 ± 0.0049; n = 5) compared with controls (0.36 ± 0.019; n = 5) ~1.4-fold, similar to the results with okadaic acid. Calyculin A pretreatment also increased mean open time from 1.3 ± 0.075 to 2.3 ± 0.068 s, whereas mean closed time did not change significantly (2.3 ± 0.16 to 2.6 ± 0.072 s). Microcystin pretreatment for 1 h also increased mean Po from 0.37 ± 0.013 to 0.49 ± 0.0064 (n = 5), again an ~1.4-fold increase. Microcystin pretreatment also increased mean open time from 1.3 ± 0.029 to 2.5 ± 0.053 s, whereas mean closed time did not change significantly (2.3 ± 0.16 to 2.6 ± 0.069 s). These results are summarized in Table 1.

An inactive analog of okadaic acid, okadaone, has no effect on channel Po. Okadaic acid has occasionally been reported to have effects unrelated to its inhibition of phosphatases. These effects can be controlled for by examining the effect of a structurally related compound, okadaone, that does not inhibit phosphatases (14). Therefore, in one batch of cells we examined single channels in untreated, okadaic acid-treated, and okadaone-treated patches. In five patches in each condition, okadaic acid-treated channels consistently had a Po higher (0.41 ± 0.028) than untreated channels (0.23 ± 0.027, P < 0.05) and okadaone-treated channels (0.19 ± 0.016, P < 0.05), and there was no significant difference in Po between untreated channels and okadaone-treated channels (Fig. 7). Therefore, we conclude that the effect of okadaic acid is specifically related to its ability to inhibit PPs.


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Fig. 7.   An inactive analog of okadaic acid, okadaone, does not alter the open probability (Po) of sodium channels. Single channels were examined in patches from untreated, okadaic acid-treated, and okadaone-treated cells. In five patches in each condition, okadaic acid-treated channels consistently had a Po higher (0.41 ± 0.028) than untreated channels (0.23 ± 0.026, P < 0.05) and okadaone-treated channels (0.19 ± 0.016, P < 0.05), and there was no significant difference in Po between untreated channels and okadaone-treated channels.

Incubation with 10 nM okadaic acid or 0.4 nM microcystin does not affect sodium channel activity. To distinguish whether the okadaic acid effect was directed on PP1 or PP2A, we pretreated A6 cells for 1 h with 10 nM okadaic acid or 0.4 nM microcystin (remembering that the IC50 of okadaic acid is 50-300 nM for PP1 and 0.1-2 nM for PP2A, and that of microcystin is 1.7 nM for PP1 and 0.04 nM for PP2A; see Ref. 14). This treatment did not produce any significant effect on sodium channel activity, suggesting that the inhibitor's effects on sodium channels are mediated by inhibition of a type 1 phosphatase. The mean sodium channel Po after incubation with okadaic acid was 0.17 ± 0.050 (n = 6) compared with the control mean Po of 0.18 ± 0.089 (n = 4). For microcystin treatment of a different batch of cells, the mean Po was 0.29 ± 0.034 (n = 5) compared with the control mean Po of 0.30 ± 0.049 (n = 5). Although these results appear to imply a type 1 phosphatase, some care should be used in interpreting these results, since the pharmacology is based on the responses in mammalian cells and the response of X. laevis phosphatases may be different.

Pretreatment with okadaic acid does not alter the I-V relationship. Figure 8 shows the I-V relationship for sodium channels from cell-attached patches obtained by plotting the single channel current amplitudes measured at various negative applied pipette potentials, -Vp (negative values = hyperpolarization, positive values = depolarization). Even without knowing the actual apical membrane potential, -Vp values ranging from -80 to +60 mV should cover the physiological range of transmembrane potentials. The I-V curves obtained from untreated cells and cells pretreated with okadaic acid for 30 min and 1 h (black-down-triangle ) were perfectly superimposable. Importantly, they were also consistent with I-V curves previously obtained for highly selective sodium channels in A6 and mammalian distal nephron cells (24). It is thus clear that okadaic acid does not affect transepithelial sodium transport through alterations in the rectification properties, ion selectivity, or the unit conductance of apical sodium channels.


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Fig. 8.   Okadaic acid does not alter the sodium channel current-voltage (I-V) relationship. Single sodium channel current amplitudes (pA) were recorded at various pipette potentials (-Vp) from untreated cells () or cells pretreated with 100 nM okadaic acid for 30 min () or 1 h (black-down-triangle ). Data points represent means ± SE. Lines are the best least squares fits to the data, and the unit conductances (~3.8 pS) were calculated from the slope near the resting membrane potential.

Time course for okadaic acid-induced effects. From all of the data in Table 1 combined, we plotted the time course for the stimulatory effect of phosphatase inhibition on the Po of sodium channels (Fig. 9). After 30 min of okadaic acid pretreatment, the sodium channel Po was increased but was not significantly different from that measured under control conditions. Therefore, it appeared that the okadaic acid-induced effect was only detectable in A6 cells pretreated for >30 min and reached its maximal value between 1 and 2 h after phosphatase inhibitor addition. The time course for the okadaic acid-induced increases in amiloride-sensitive Isc (Fig. 1) was similar to the time course for increases in individual sodium channel Po (Fig. 9), although the Isc response appeared to be delayed relative to the single channel response. Despite the differences in the two measurement techniques and the use of two different A6 cell clones, these results appear to be consistent.


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Fig. 9.   Time course for okadaic acid-induced stimulation of Po. Po values relative to the Po at time 0 were measured from patches as a function of pretreatment time with 100 nM okadaic acid. Data points are taken by combining the values of Po for all patches represented in Table 1 and are means ± SE. At 30 min, there is an increase, but it is not statistically significant (P = 0.053). At 1 h, the okadaic acid increase is highly significant (P < 0.01).

Acute perfusion with okadaic acid affects sodium channel Po. We and other groups have previously noted the large variability in Po that exists among single, high-selectivity sodium channels from different cell-attached patches (24). Because such variability might have obscured our ability to detect a subtle effect of okadaic acid pretreatment before 1 h, we also examined individual cell-attached patches before and after acute perfusion with 100 nM okadaic acid. Immediately after a gigaohm seal was established, baseline channel activity was recorded for ~4-5 min. The phosphatase inhibitor was then perfused in the apical bath chamber for ~10 min (see MATERIALS AND METHODS). We then proceeded with a single channel recording for as long as high-resistance seals could be maintained. During the course of these experiments, all patches were held at resting membrane potential. To maximize our ability to detect a subtle, early, okadaic acid effect, these experiments were performed on patches initially containing low baseline sodium channel activity (low N, a low Po, or both). In general, the acute effect of okadaic acid was not large, although Po decreased in only two of the eight patches we examined (Fig. 10B). However, comparing channel activity before and after application of okadaic acid may be inappropriate, since we really should compare the effect of okadaic acid with the effect of no treatment. This is particularly so because highly selective sodium channels in A6 cell-attached patches often show a spontaneous decay in activity ("rundown"), which could obscure the stimulatory effect of okadaic acid. Therefore, for comparison, we examined 12 patches from the same batch of cells with initial Po comparable to those treated with okadaic acid. We recorded for an initial 4-min period, perfused the patches with A6 saline for ~10 min, and then recorded for four additional minutes to produce recordings comparable to the time course for okadaic acid-perfused patches. Figure 10A plots Po for the first (left) and second (right) 4 min of cell-attached recording and shows a 49% decrease in mean Po from 0.25 ± 0.036 to 0.16 ± 0.033 (P = 0.006; paired t-test). This frequent spontaneous loss of channel activity in the cell-attached configuration strongly suggested that we could have underestimated the stimulatory effects of okadaic acid. The mean changes in Po between the first and second 4 min of recording from untreated cells (-0.10 ± 0.030; Fig. 10A and Table 2) and between the 4-min periods before and after okadaic acid perfusion (+0.047 ± 0.06; Fig. 10B and Table 2) were significantly different (P = 0.019; t-test). However, despite acute perfusion of A6 cells with okadaic acid, channel activity still eventually decayed (data not shown). Unfortunately, we were unable to maintain stable cell-attached patches for >30-40 min, preventing a direct comparison of acute okadaic acid-induced effects with that after more prolonged pretreatment.


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Fig. 10.   Acute okadaic acid perfusion prevents increases in the Po of sodium channels above that for untreated patches. A: sodium channel Po for cell-attached patches at the resting membrane potential are calculated for the 4-min period after a gigohm seal is established and then for the subsequent 4 min after 10 min of saline perfusion. B: Po is calculated for the 4-min periods after a gigohm seal is established and the subsequent 4-min period after a 10-min perfusion with 100 nM okadaic acid. Lines connecting symbols represent data from the same cell-attached patch. In patches on untreated cells, mean Po values for the first period (0.25 ± 0.036) and the second period (0.15 ± 0.033) were significantly different by paired t-testing (P < 0.001). Mean Po values before (0.24 ± 0.059) and after (0.29 ± 0.051) okadaic acid perfusion were not significantly different by paired t-testing.


                              
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Table 2.   Acute perfusion of okadaic acid increases sodium channel activity

Okadaic acid alters the phosphorylation of some cellular proteins but does not alter the phosphorylation of alpha -, beta -, or gamma -ENaC. Because of the manner in which we performed our experiments, the effect of the phosphatase inhibitors could be due to either a change in the phosphorylation of the sodium channels themselves or the phosphorylation of another protein, which in turn regulates sodium channel activity. In an attempt to investigate these possibilities, we labeled A6 cells to equilibrium with [33P]phosphate, washed out excess phosphate, and then examined the effect of a 1-h treatment of the cells with okadaic acid on the patterns of total cellular phosphorylation. The problem with this approach is that phosphatases, unlike kinases, have relatively nonspecific targets; thus, decreasing phosphatase activity will generally increase the phosphorylation of many proteins that were originally phosphorylated by quite specific serine/threonine kinases. This problem is obvious in Fig. 11A, where a few phosphoprotein bands (imaged on a PhosphorImager) are relatively distinct in untreated cells, but after okadaic acid treatment there are too many bands to resolve. This does indicate that there are many proteins that are differentially phosphorylated by okadaic acid; however, we could identify no single protein band as a candidate for producing the okadaic acid-induced increase in sodium channel Po. We also wished to test whether sodium channel proteins themselves might be a target for okadaic acid-induced phosphorylation. Sodium channel proteins consist of three subunits, alpha , beta , and gamma . We could identify the phosphorylation of the subunits by immunoprecipitation of the subunit with antisubunit antibodies. When we immunoprecipitated the three subunits from cellular extracts of okadaic acid-treated or untreated cells labeled as above and examined the immunoprecipitate on SDS gels, we could find no evidence for differential phosphorylation of ENaC (Fig. 11B). However, the baseline phosphorylation, particularly of the beta - and gamma -subunits, was high enough that we might not have detected as much as a 50% change in the okadaic acid-induced phosphorylation.


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Fig. 11.   Okadaic acid does not alter the phosphorylation of epithelial sodium channel (ENaC) subunit proteins. We labeled A6 cells to equilibrium with [33P]- or [32P]phosphate, washed out excess phosphate, and then examined the effect of 1 h of treatment of the cells with okadaic acid on the phosphorylation of all proteins and the phosphorylation of ENaC. A: all phosphoproteins were labeled in the presence and absence of okadaic acid. This shows that okadaic acid does increase the levels of protein phosphorylation, but it also shows that examining all phosphoproteins is not very useful since, after okadaic acid, there are too many bands to resolve individual proteins. Mol wt, molecular weight. B: on the other hand, when we immunoprecipitated the three ENaC subunits from 32P-labeled cellular extracts of okadaic acid-treated or untreated cells labeled as in A, resolved the immunoprecipitate on SDS gels, and quantitated the phosphorylation on a PhosphorImager, we could find no evidence for differential phosphorylation of ENaC by okadaic acid. However, the baseline phosphorylation, particularly of the beta - and gamma -subunits, was high enough that we might not have detected as much as a 50% change in the okadaic acid-induced phosphorylation. The bands for the beta - and gamma -subunits run at 95 and 98 kDa, respectively, which is similar to that reported by us and others in previous work. The alpha -subunit consists of two bands at 78 and 85 kDa, which we presume corresponds to core glycosylated and unglycosylated forms of alpha , as previously reported by others.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We have demonstrated that there are potential targets in A6 cells for all of the phosphatase inhibitors we applied. However, despite the presence of both PP1 and PP2A, based on the pharmacology PP1 is a possible candidate to alter the activity of ENaCs.

On the basis of transepithelial current measurements alone, it is impossible to determine the specific mechanisms responsible for the okadaic acid-induced increase in sodium permeability, since total sodium current per unit area of apical membrane is the product of the unit conductance, Po, N, and electromotive driving force. In theory, modification of any of these parameters could alter apical sodium flux. We know from our previous studies that PKC-mediated phosphorylation alters Po, PKA alters N, and different tyrosine kinases may alter both N and Po (36, 39). Therefore, to correlate phosphatase activity with a specific protein kinase pathway requires an examination of how dephosphorylation could affect the parameters that alter sodium transport. To do this, we supplemented transepithelial current measurements with single channel recordings from cell-attached patches and simple measurements of cellular phosphorylation. We used both standard single channel statistical analysis of records from patches containing one or a few channels (N <=  4) and fluctuation analysis of patches containing a large number of channels (N > 4). Use of these techniques allowed us to obtain consistent information on the effects of okadaic acid and other phosphatase inhibitors, both on the number of channels per patch (N) and on intrinsic channel properties like Po and the rate constants for individual channel openings and closings.

Sodium channel activity is dependent on phosphorylation/dephosphorylation reactions. In mineralocorticoid-stimulated A6 cells, okadaic acid induces an increase in the Po of highly selective sodium channels with little, if any, effect on apical membrane channel density. This is consistent with the presence of an okadaic acid-sensitive phosphatase that counteracts a stimulatory protein kinase-mediated phosphorylation step. Indeed, several different types of phosphorylation reactions are known to modulate sodium transport in distal nephron cells. However, the literature and our previous work preclude many of these protein kinase pathways from being responsible for the phosphorylation step linked to the phosphatase blocked by okadaic acid. Our results indicate that the serine/threonine kinase associated with okadaic acid-inhibitable phosphatase must stimulate sodium channels by increasing their Po. Phosphorylation reactions mediated by PKC in native cells are inhibitory (23, 36 and see below).

PKA activation does result in stimulation of apical sodium transport (39). PKA activity is induced by arginine vasopressin and would not be expected to be stimulated under the aldosterone-stimulated conditions of this study. However, the mechanism by which PKA alters sodium channel activity is somewhat controversial, since at least one amiloride-sensitive channel from A6 cells is stimulated by PKA (43). Unfortunately, it is not clear exactly how this latter channel is related to the channel responsible for physiologically relevant sodium uptake. In any event, if inhibition of a phosphatase produces an increase in single channel activity, then the kinase producing the increase must be tonically active in A6 cells. However, our laboratory has recently shown that PKA inhibition by H-89 has no effect on baseline sodium channel activity (12), thus ruling out the presence of tonically active PKA in A6 cells grown in the presence of aldosterone. Finally, PKA has been shown to increase the apparent density of sodium channels in the apical membrane of A6 cells without significantly affecting the Po (39).

The okadaic acid concentration we used (100 nM) should exclude nonspecific inhibition of PKC or tyrosine kinase (14). Regardless, based on our previous studies, a nonspecific effect on tyrosine kinase would also be expected either to alter sodium channel density or reduce Po (40). Also, the similar effect of two other phosphatase inhibitors also supports the idea that the effect is on a serine/threonine phosphatase. In addition, the lack of effect of okadaone also argues for a specific effect of the inhibitors.

Still, our present results suggest that at least one PP is tonically active in A6 cells grown in the presence of aldosterone and is in some way linked to sodium channel activity. We would predict that this tonically active phosphatase reverses a phosphorylation step of unknown nature, which enhances sodium channel activity. Many possible target proteins exist that might modify apical sodium transport in response to phosphorylation/dephosphorylation reactions, and our examination of the phosphorylation of all cellular proteins does show differential phosphorylation in response to okadaic acid. One seemingly reasonable target for phosphorylation would be one or more of the sodium channel subunits themselves. Indeed, recent work in a model system has shown that the beta - and gamma -subunits but not the alpha -subunit are phosphorylated in response to aldosterone, insulin, and activators of PKC (46). However, in our present work, there does not appear to be any differential phosphorylation of either the alpha -, beta -, or gamma -subunits in response to okadaic acid. Based on the previous results in Madin-Darby canine kidney (MDCK) cells, the lack of phosphorylation of the alpha -subunit is not surprising, but, if the phosphatase inhibited by okadaic acid is related to any of the kinases stimulated in the MDCK cells, then we would have expected phosphorylation of the beta - or gamma -subunits. The lack of constitutive phosphorylation in native renal cells is consistent with our previous work (58, 59) in which we did not observe changes in phosphorylation even when protein kinases were stimulated. Thus we must conclude that, under the conditions used in our experiments, phosphorylation of the ENaC subunits does not play a role in the okadaic acid-stimulated increase in channel Po. This result is consistent with the observations of Volk et al. (56, 57), who showed that some staurosporine-sensitive kinase increased the activity of ENaC but did not appear to phosphorylate ENaC directly. We tentatively conclude the most likely target is some modulatory protein known to be associated with renal ENaCs.

One possibility is that okadaic acid is promoting SGK-mediated phosphorylation of the ENaC ubiquitin ligase Nedd4. Nedd4 is responsible for binding to and ubiquitin conjugation of ENaC subunits (4, 25, 45, 48-50). Ubiquitin conjugation has the following two effects: first, we have shown that ubiquitin conjugation increases the Po of conjugated channels (38); and, second, ubiquitin conjugation promotes removal of ENaC from the surface membrane, thus reducing the number of functional ENaCs. Recently, two groups have shown that increased phosphorylation of Nedd4 reduces Nedd4 activity, leading to reduced ubiquitin conjugation of ENaC and an increased lifetime of ENaC in the surface membrane (17, 47). This observation is consistent with the reduced rate of loss of ENaC activity associated with exposure of cells to okadaic acid but does not explain the increase in Po. In fact, if ubiquitin conjugation was reduced by a reduction in Nedd4 activity, one might expect a decrease in Po based on our previous patch-clamp results (38).

Of course, the mechanisms for the increase in Po and the decreased rate of loss of channel activity do not have to be the same. An alternative mechanism to explain the increase in Po involves the recent observation that a methylation reaction increases the Po of ENaC (6, 52, 53) and the enzyme responsible for methylation is activated by serine phosphorylation (1). Ascertaining if either or both of these mechanisms is important in the action of phosphatases on ENaC activity in A6 cells will require future studies in which the phosphorylation of these regulatory molecules is examined and correlated with channel activity. However, examination of okadaic acid-induced changes in the phosphorylation of modulatory proteins will not be simple, since, unlike kinases, phosphatases have relatively nonspecific targets; thus, decreasing phosphatase activity will generally increase the phosphorylation of many proteins that are phosphorylated by quite specific serine/threonine kinases. This problem was obvious in Fig. 11A, where okadaic acid increased protein phosphorylation to the extent that it was difficult to identify any individual protein that was phosphorylated. However, we have recently demonstrated that, not surprisingly, okadaic acid does increase the phosphorylation of isoprenyl-cysteine methyltransferase (1). This would be expected to increase the Po (6, 52), but determining if this is the mechanism by which okadaic produces its effect will require inhibiting the function of the methyltransferase in the presence of okadaic acid.

The I-V relationship is not affected by phosphatase inhibition. Pretreatment of A6 cells with okadaic acid did not appreciably alter the single sodium channel I-V curve. Therefore, okadaic acid had no affect on the sodium channel unit conductance or ion selectivity. However, because the short-circuit sodium current and sodium channel Po are increased by okadaic acid, one may wonder why the apical membrane potential did not appear to change with the increase in apical sodium entry (i.e., the curve of the I-V reversal potential did not shift). The most likely explanation for this observation is that the increased intracellular sodium activity after okadaic acid treatment contributed to an increase in the rate of Na+-K+-ATPase pump exchange, with a concomitant increase in transepithelial voltage, as is generally observed in tight epithelia (19, 26). Such an increase would hyperpolarize the apical membrane and counteract the depolarizing effect of the increased apical sodium permeability. Thus this is also an indication that A6 cells exhibit normal physiological responses after okadaic acid addition and gigaohm seal formation.

Sodium channels spontaneously lose activity with time. Understanding the mechanism underlying the frequently observed loss of activity of A6 cell sodium channels in the cell-attached mode could have theoretical and practical interest (for an example, see Ref. 59). Okadaic acid was an effective stimulus to sodium channel activity both acutely (minutes) and chronically (hours), thereby attenuating, but not abolishing, normal loss of ENaC activity. Because loss of ENaC activity was only delayed, the okadaic acid-sensitive phosphatase cannot be the only mechanism producing the loss in activity. Although okadaic acid transiently increased sodium channel Po, an independent and superimposed process eventually reduced the Po of most of the channels. In view of the requirement for normal structural links between the cytoskeleton and renal ENaCs for the maintenance of channel activity (7, 8, 10), the working hypothesis we would favor is that the mechanical action of the patch pipette could, in a fraction of experiments, lead to a disruption of this link and loss of channel activity. This hypothesis is now under study in our laboratory.

Our results suggest that okadaic acid can chronically and, to a lesser extent, acutely activate ENaC in apical cell-attached patches. An okadaic acid-sensitive, serine/threonine phosphatase tonically reduces the phosphorylation of a protein, which is intimately related to the control of the sodium channel complex in A6 distal nephron cells. Using transepithelial, fluctuation and single channel analysis methods, we also show that inhibition of a phosphatase that is likely to be PP1 affects channel kinetics with only a small effect on channel density. In addition, spontaneous loss of ENaC activity is delayed but finally occurs despite reduction of serine/threonine dephosphorylation reactions. Finally, ENaC subunits in A6 cells under our experimental conditions do not appear to be the targets for serine/threonine phosphorylation.


    ACKNOWLEDGEMENTS

We thank Elisabeth E. Seal and Billi Jean Duke for skillful technical assistance in the preparation of the A6 cell cultures suitable for patch-clamp work.


    FOOTNOTES

Present addresses: A. Becchetti, Dipartmento di Biotecnologie e Bioscience, via L. Emanueli 2, 20126 Milano,Italy (E-mail: andrea.becchetti{at}unimib.it); P. Duchatelle, Department de Physiologie, Université de Caen, Caen, France (E-mail: p.duchatelle{at}scvie.unicaen.fr).

This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants R01-DK-37963, DK-56305, and P01DK-50268 to D. C. Eaton, R01-DK-51391 and R01-DK-54354 to T. R. Kleyman and by core support from the Center for Cell and Molecular Signaling.

Address for reprint requests and other correspondence: D. C. Eaton, Emory Univ. School of Medicine, Dept. of Physiology, 1648 Pierce Dr., N.E., Atlanta, GA 30322 (E-mail: deaton{at}emory.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

June 26, 2002;10.1152/ajprenal.00011.2002

Received 10 January 2002; accepted in final form 3 June 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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