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
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ABSTRACT |
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
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INTRODUCTION |
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
- and
-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).
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MATERIALS AND METHODS |
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
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
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
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, 
,
of n channels is
where average Po is the open probability
of the n channels. The variance in the patch current
is
where i is the current of a single channel. The mean
patch current, I, is
Combining these expressions gives
The power spectra are related to the current variance, since the
integral of the power spectrum Lorentzian function is equal to the
variance
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
The number of channels in the patch, N, can be
calculated since
and finally, the individual forward and reverse rate constants
can be calculated as
and
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
-,
-, and
-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-
-,
anti-
, or anti-
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.
 |
RESULTS |
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
(PP1
1) and two for PP2A (PP2A
and PP2A
). 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 ( , 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.
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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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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.
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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.
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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).
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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.
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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 (
)
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 ( ). 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.
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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).
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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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Okadaic acid alters the phosphorylation of some cellular proteins
but does not alter the phosphorylation of
-,
-, or
-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,
,
, and
. 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
- and
-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 - and -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
- and -subunits run at 95 and 98 kDa, respectively, which is
similar to that reported by us and others in previous work. The
-subunit consists of two bands at 78 and 85 kDa, which we presume
corresponds to core glycosylated and unglycosylated forms of , as
previously reported by others.
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 |
DISCUSSION |
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
- and
-subunits but not the
-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
-,
-,
or
-subunits in response to okadaic acid. Based on the previous
results in Madin-Darby canine kidney (MDCK) cells, the lack of
phosphorylation of the
-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
- or
-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.
 |
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