Differential regulation of single CFTR channels by PP2C, PP2A,
and other phosphatases
Jiexin
Luo1,
Mary D.
Pato2,
John R.
Riordan3, and
John W.
Hanrahan1
1 Department of Physiology,
McGill University, Montreal, Quebec H3G 1Y6;
2 Department of Biochemistry,
University of Saskatchewan, Saskatoon, Saskatchewan, Canada S7N
0W0; and 3 S. C. Johnson
Medical Research Center, Mayo Clinic Scottsdale, Scottsdale, Arizona
85259
 |
ABSTRACT |
Cystic fibrosis
transmembrane conductance regulator (CFTR)
Cl
channel activity
declines rapidly when excised from transfected Chinese hamster ovary
(CHO) or human airway cells because of membrane-associated phosphatase
activity. In the present study, we found that CFTR channels usually
remained active in patches excised from baby hamster kidney (BHK) cells
overexpressing CFTR. Those patches with stable channel activity were
used to investigate the regulation of CFTR by exogenous protein
phosphatases (PP). Adding PP2A, PP2C, or alkaline phosphatase to
excised patches reduced CFTR channel activity by >90% but did not
abolish it completely. PP2B caused weak deactivation, whereas PP1 had
no detectable effect on open probability
(Po).
Interestingly, the time course of deactivation by PP2C was identical to
that of the spontaneous rundown observed in some patches after
excision. PP2C and PP2A had distinct effects on channel gating;
Po declined
during exposure to exogenous PP2C (and during spontaneous rundown, when
it was observed) without any change in mean burst duration. By
contrast, deactivation by exogenous PP2A was associated with a dramatic
shortening of burst duration similar to that reported previously in
patches from cardiac cells during deactivation of CFTR by endogenous
phosphatases. Rundown of CFTR-mediated current across intact T84
epithelial cell monolayers was insensitive to toxic levels of the PP2A
inhibitor calyculin A. These results demonstrate that exogenous PP2C is a potent regulator of CFTR activity, that its effects on single-channel gating are distinct from those of PP2A but similar to those of endogenous phosphatases in CHO, BHK, and T84 epithelial cells, and that
multiple protein phosphatases may be required for complete deactivation
of CFTR channels.
cystic fibrosis; protein phosphatase; channel rundown; cystic
fibrosis transmembrane conductance regulator
 |
INTRODUCTION |
REGULATION of the cystic fibrosis transmembrane
conductance regulator (CFTR)
Cl
channel by protein
kinases has been intensively studied during the past several years (4,
7, 12, 13, 16, 17, 35). Phosphorylation by protein kinase A (PKA)
increases open probability (Po) (35),
bursting rate (20, 23), apparent ATP affinity (20, 23), and ATP
hydrolysis rate (20). Phosphorylation by protein kinase C (PKC) is also
required for CFTR to respond to PKA (17). These actions of kinases are
antagonized by protein phosphatases, which probably vary among
different cell types. CFTR channel activity declines rapidly when
patches are excised from stimulated Chinese hamster ovary (CHO) or
airway epithelial cells into bath solution lacking PKA [~10 s
at 37°C (35); 100 s at 22°C (2)]. This spontaneous
decline in channel activity, or rundown, is slower or absent
when channels are studied in patches from guinea pig myocytes (15),
transfected fibroblasts (4), or Hi-5 insect cells (39).
Characterization and molecular identification of the phosphatases
regulating CFTR has become a priority because they are potential
therapeutic targets in the treatment of cystic fibrosis (2).
Serine and threonine phosphatases are functionally classified into two
types, protein phosphatase 1 (PP1) and protein phosphatase 2 (PP2). The
latter is subclassified into protein phosphatase 2A (PP2A), protein
phosphatase 2B (PP2B), and protein phosphatase 2C (PP2C) (see Refs. 24,
34). PP1 preferentially dephosphorylates the
-subunit of
phosphorylase kinase and is sensitive to inhibitors 1 and 2, whereas PP2
preferentially acts on the
-subunit of phosphorylase kinase and is
insensitive to these inhibitors. PP2A activity does not require
particular ions or cofactors, in contrast to PP2B, which requires
Ca2+ and calmodulin, and PP2C,
which requires relatively high levels of
Mg2+
(EC50 ~1.5 mM; Ref. 8). PP1 and
PP2A are both sensitive to okadaic acid and calyculin A but can be
distinguished by using appropriate concentrations of these inhibitors
(24, 34). PP2B can be identified by its sensitivity to inhibitors such
as deltamethyrin, cyclosporin, or FK-506 (10, 34). No specific
inhibitors of PP2C are available. Rundown and CFTR dephosphorylation
are both inhibited by phenylimidazothiazoles (2, 3), but at much higher
concentrations than are needed to inhibit alkaline phosphatase.
CFTR channel rundown in excised patches is relatively insensitive to
okadaic acid (2, 35), suggesting regulation by a robust
membrane-associated protein phosphatase other than PP1 and PP2A. These
results do not exclude regulation of CFTR by PP2A on the cell, however,
since any cytosolic PP2A would be lost from patches after excision.
Indeed, Reddy and Quinton (29) showed that okadaic acid
(10
8 M) inhibits
deactivation of CFTR currents in permeabilized sweat ducts. In guinea
pig cardiac myocytes, ~40% of the deactivation after forskolin
washout was blockable by okadaic acid or microcystin (15). Exogenous
PP1 and PP2B are not effective in regulating CFTR currents in excised
patches from fibroblasts (4), although there is evidence that CFTR
activity can be stimulated in these cells by the PP2B inhibitors
cyclosporin A or deltamethyrin, suggesting that PP2B can regulate CFTR
(11). By default, PP2C has been proposed as a CFTR phosphatase because
it is insensitive to okadaic acid and microcystin (15, 35). PP2C
dephosphorylates CFTR and deactivates macroscopic CFTR
Cl
currents (38). Deactivation by PP2C and other
phosphatases has not been studied at the single-channel level.
The goal of this study was to characterize the effects of PP1, PP2A,
PP2B, and PP2C on single CFTR channels exposed to comparable levels of
phosphatase activity. For comparison, the kinetics of CFTR channels
were also examined during deactivation by endogenous phosphatases.
Finally, we investigated the possible role of PP2A in intact T84
epithelial monolayers by examining the ability of calyculin A to
inhibit deactivation of short-circuit current
(Isc) after
washout of forskolin. The results suggest that CFTR deactivation is
mediated primarily by a PP2C-like phosphatase in CHO, baby hamster
kidney (BHK), and T84 epithelial cells, although PP2A and PP2B both
cause partial deactivation in vitro. These results have been reported
in preliminary form (21, 22).
 |
MATERIALS AND METHODS |
Cell culture.
BHK cells stably expressing wild-type CFTR were plated at low density
on glass coverslips 3-5 days before use in patch-clamp experiments. The T84 line was obtained from American Type Culture Collection (Rockville, MD) and studied between
passages 77 and 115. T84 cells were plated at a
density of 400,000/cm2 on porous
supports (Millipore, Toronto, ON, Canada), which had been coated with a
gel of type I collagen. The growth medium was a 50:50 mixture of DMEM
and Ham's F-12 medium and was supplemented with 15 mM HEPES, fetal
bovine serum (5%), penicillin (100 IU/ml), and streptomycin (100 µg/ml). Monolayers were studied 8-12 days after plating, when
transepithelial resistance had reached ~1,500
· cm2.
Phosphatases.
Recombinant human PP1
catalytic subunit was purchased from
Calbiochem (La Jolla, CA). PP2AI (smooth muscle phosphatase I) and PP2C
(smooth muscle phosphatase II) were prepared from turkey gizzard smooth
muscle as described previously (26, 27). PP2A was further purified by
sequential chromatography on DEAE-Sephacel,
-aminooctyl-Sepharose,
and an affinity column of thiophosphorylated 20,000 Mr myosin light
chains coupled to Sepharose 4B. The PP2C fraction from a Sephacryl
S-300 column was chromatographed on DEAE-Sephacel and on the affinity
column mentioned above. PP2C bound to the column in the presence of
Mg2+ and was selectively eluted
using EDTA. Bovine brain PP2B was purchased from Boehringer Mannheim
(Laval, QC, Canada). Purity of the protein phosphatase preparations was
assessed by SDS-PAGE (19) in a 12.5% Microslab gel and then stained
with Coomassie blue. For comparison with the four protein phosphatases,
some patch-clamp experiments were also carried out using bovine
intestinal alkaline phosphatase type VII-S (Sigma, St. Louis, MO). The
activity of this enzyme was 2,000-3,000 U/mg enzyme [where 1 unit hydrolyzes 1.0 µmol of
p-nitrophenyl phosphate (PNP)/min at
37°C]. Alkaline phosphatase was used at a final concentration
of 80 U/ml.
Enzyme activities were determined by measuring release of
[32P]orthophosphate
from phosphorylated myosin light chains as described previously (28).
Assays were carried out under four conditions: 1) high-salt solution used during
patch-clamp experiments, 2) low-salt
solution (same as 1 but lacking 150 mM
NaCl), 3) standard Tris-dithiothreitol (DTT) solution, and
4) Tris only solution (same as
3 but lacking DTT). These and other
solutions are listed in Table 1. The
substrate used in all phosphatase assays was prepared as described
previously (33) and had a specific activity of 3,736 cpm/pmol. After
adding phosphatase to the reaction mixture, aliquots were taken at
timed intervals, and the reaction was terminated by addition to 100 µl of 17.5% TCA and 100 µl of 6 mg/ml BSA. The resulting solutions
were chilled and centrifuged at 15,000 rpm for 1 min. Aliquots of the
supernatant (200 µl) were counted by liquid scintillation (Beckman LS
7800) to determine the amount of
32P released.
Patch-clamp studies.
BHK cells were placed in a recording chamber (200 µl vol), containing
(in mM) 150 NaCl, 2 MgCl2, and 10 TES (pH 7.4). In most experiments, this solution also contained 0.5 mM
MgATP and 100 nM PKA catalytic subunit (prepared in laboratory of Dr.
M. P. Walsh, University of Calgary). Patch-clamp experiments were
carried out at room temperature (22°C). Pipettes were pulled in two
stages (PP-83, Narishige Instrumentation Laboratory, Tokyo, Japan) and had resistances of 4-6 M
when filled with 150 mM NaCl solution. The bath was grounded through an agar bridge having the same ionic composition as the pipette solution. Single-channel currents were recorded from both cell-attached and excised patches; the pipette potential was held at +30 mV. Single-channel currents were amplified (Axopatch 1B, Axon Instrument, Foster City, CA), recorded on
videocassette tape by a pulse-coded modulation-type recording adapter
(DR384, Neurodata Instrument, New York), and low-pass filtered during playback using an eight-pole Bessel filter (900 LPF, Frequency Devices,
Haverhill, MA). Final records were sampled at 0.5 kHz and analyzed
using a laboratory microcomputer system and DRSCAN, a pCLAMP-compatible
program developed in this laboratory for analyzing long records.
Channel cycle time was used to calculate the mean burst duration
(
o) as
|
(1)
|
where
N is the number of channels,
Po is the
probability of each channel being in the open state (i.e., above a
threshold set at half open-channel amplitude), T
is the duration of the segment analyzed, and
b is the number of bursts observed
during the segment (14). This method of calculating
o is useful
because it does not require determination of
N, relying instead on
NPo, which can be
measured regardless of the number of channels as
|
(2)
|
where
ti is the time
spent above a threshold "i" set at 0.5, 1.5, 2.5, ... times the single-channel current amplitude. To evaluate changes
in channel gating,
o was
calculated for each 5-s segment in the record. Because each of the
~200
o values is essentially a sample estimate
(
) of the mean (µ) and SD
(
), the sampling distribution of
o has the form Z = (
µ)/(
/
), which is closely
approximated by a Gaussian distribution when there are
n > 30 observations.
Calculation of Po
and mean interburst duration do require estimates of
N.
Po was determined
by dividing NPo
by N, estimated as the largest number
of channels open simultaneously during long recordings when the
channels were stimulated (i.e., when
Po was high,
before addition of phosphatase). This method tends to underestimate N and therefore overestimate
Po; however, the
error is small for wild-type CFTR channels (<10%) (23). If the
method for calculating
o were
sensitive to N, any error should be
similar for all phosphatases and therefore would not explain their
different effects on burst duration. Finally, data were obtained from
the same patches before and after adding phosphatases; therefore
correction for any underestimate of N
would, if anything, tend to increase differences between phosphatase
effects. We did not attempt to calculate the mean interburst
duration,
c = [(N
N · Po) · T)/(n
1)]. Interburst duration is an absolute measurement that
requires an accurate determination of
N. This could not be obtained by
adding AMP-PNP at the end of each recording (23), since channels were
not locked open after phosphatase exposure.
Transepithelial experiments.
T84 cell monolayers were studied in water-jacketed Ussing-type chambers
(Vangard International, Neptune, NJ) that had been fitted with Teflon
adaptors for holding cell culture inserts. The control solution
contained (mM) 115 NaCl, 2.5 K2HPO4,
1.5 CaCl2, 1 MgSO4, 10 glucose, and 25 NaHCO3 and was gassed with 95%
O2-5%
CO2 at 37°C. Forskolin (10 µM) was used to stimulate Isc, which, in
this preparation, is carried entirely by net
Cl
secretion. All chemicals
were from Sigma except calyculin A, which was from Calbiochem (San
Diego, CA). Monolayers were short-circuited using a conventional
voltage clamp (DVC-1000, WPI Instruments, Sarasota, FL), which applied
brief voltage pulses every 2 min to monitor transepithelial resistance.
Isc values were
transferred to a computer spreadsheet program for calculations and
graphics.
Statistics.
Values are presented as means ± SE. Significance was assessed at
the 95% confidence level using the Student's
t-test. Histograms of
o were
fitted with Gaussian curves by least squares using Origin software
(version 4.1, Microcal Software, Northampton, MA).
 |
RESULTS |
Characterization of protein phosphatase preparations.
Figure 1 shows an SDS-polyacrylamide gel of
the four protein phosphatase preparations used in patch-clamp
experiments, stained with Coomassie blue. The PP1 and PP2B preparations
did not contain detectable contamination. The PP2A had 8% contaminant
with Mr of
~90,000, whereas the PP2C profile shows two impurities
(Mr ~30,000),
constituting <3% of the protein. Western blot analysis with specific
antibodies revealed that the PP2A preparation did not contain
detectable PP2C or vice versa (data not shown). This was confirmed by
functional assays; okadaic acid (0.01 nM) inhibited 50% of the PP2A
activity but had no effect on the PP2C preparation. Representative
activity curves of PP2C using phosphorylated myosin light chains as
substrate under different conditions are shown in Fig.
2. Orthophosphate release was linear during
the first 75 s under all conditions and during the first 2 min under
the high-salt conditions used in patch-clamp experiments. Phosphatase activities calculated from the initial linear release rates are summarized in Table 2. All four enzymes
were inhibited by the patch-clamp solution containing 150 mM NaCl. This
was most pronounced for PP1 (72%) and PP2A (66%). PP2B activity was
negligible in the absence of Ca2+
and calmodulin. The activity of PP1 was relatively labile, declining two- to threefold when diluted with BSA and stored on ice for several
hours.

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Fig. 1.
SDS-PAGE of protein phosphatases (PP) used in this study. PP2B (60 and
16 kDa; lane 1), catalytic subunit
of PP1 (38 kDa; lane 2), PP2A (60, 55, and 36 kDa; lane 4), and PP2C
(43 kDa; lane 5) were applied to a
12.5% Laemmli gel. Molecular masses are from Bio-Rad molecular mass
standards, in lane 3, in order of
descending size: myosin, -galactosidase, phosphorylase, serum
albumin, ovalbumin, carbonic anhydrase, trypsin inhibitor, and RNase.
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Fig. 2.
Activity of PP2C assayed under 4 different conditions. Time course of
32PO4
release from 1 µM radiolabeled myosin light chains at 30°C.
Phosphatase activity was measured in solution
1 (high-salt; ) containing 150 mM NaCl, which was
used for patch-clamp recording, same solution lacking NaCl
(solution 2, low-salt; ),
conventional Tris-buffered solution containing dithiothreitol (DTT,
solution 3; ), and same
Tris-buffered solution lacking DTT ( ). See Table 1 for composition
of solutions.
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Two channel behaviors suggest that phosphatase activity is unevenly
distributed.
CFTR channel activity was usually observed on BHK cells after
incubation with 20 µM forskolin for 5-10 min. Channel activity usually persisted for >5 min after patches were excised into bath solution containing 0.5-1 mM MgATP (Fig.
3,
A-C); however, activity did
decline rapidly in about one-fourth of the excised patches (Fig. 3,
D-F). These results contrast
with our previous observations with CHO and airway epithelial cells, in
which CFTR channel activity always declined rapidly (2, 35). We
tentatively attribute the variable rundown to an uneven distribution of
phosphatase activity in the plasma membrane of BHK cells (perhaps
exacerbated by a very high level of CFTR expression), since other
factors such as time after plating and recording conditions were kept constant. Channel activity declined to 5-10% of the starting
value within 100 s in those patches that exhibited rundown and could be
restimulated by adding PKA as in previous studies of CHO, T84, and
airway epithelial cells (2, 35, 36).

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Fig. 3.
Two types of channel activity in patches excised from baby hamster
kidney (BHK) cells. A-C: examples
of patches in which cystic fibrosis transmembrane conductance regulator
(CFTR) channels remained active after excision (typical of
~3/4 of BHK patches). A:
patch recording obtained before and after excision at arrow. Upward
deflections indicate channel openings in this and all subsequent traces
(holding potential +30 mV). After stimulation of cells by forskolin (10 µM), patches were excised into protein kinase A (PKA)-free bath
solution containing MgATP (0.5 mM). B
and C: mean number of channels open
(NPo) and mean
burst duration
( o),
respectively, calculated during 5-s segments from 3 patches.
D-F: examples of patches in which
channel activity rapidly declined after excision (typical of
~1/4 of BHK patches). D:
patch recording showing spontaneous rundown after excision at arrow.
E and
F:
NPo rapidly
declined with no change in
o; data
pooled from 3 patches.
|
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Burst duration during spontaneous rundown in excised patches.
To assess alterations in single-channel kinetics during rundown, we
calculated that
o values
during each 5-s interval after
Po had declined
by at least 90% and compared them with values obtained from patches
having stable channel activity (Fig. 3, C and
F). The distributions of open burst
durations in both groups of patches could be fitted with Gaussian
functions having similar means (n = 7 patches with rundown, n = 5 patches
without rundown; P = 0.24; see Fig.
4). Thus, when membrane-associated
phosphatase activity was present in a particular patch, it reduced CFTR
channel activity by >90% without significantly altering
o.

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Fig. 4.
Histograms of
o in patches
with and without rundown.
o was
calculated during each 5-s interval as described in
MATERIALS AND METHODS.
A: distribution of
o in a patch
with stable activity, i.e., without rundown. B: distribution
of o in a
patch displaying rundown of CFTR channel activity. Both distributions
were fitted by 2 Gaussian curves. Overall mean and SD areas derived
from fits of each Gaussian.
|
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Effect of calyculin A on CFTR channels from BHK cells.
Channel activity declined slowly in 20 of 84 patches excised in the
presence of low (50-100 nM) PKA activity and 0.5 mM MgATP. The
contribution of membrane-associated PP1/PP2A was assessed in some of
the patches that displayed rundown by addition of the potent PP2A
inhibitor calyculin A while rundown was in progress. Calyculin A
(100-1,000 nM) did not block further rundown in 10 of 12 patches,
nor did it enable the low PKA activity present in the bath solution to
restimulate channels in any of the patches (n = 10). Figure
5 shows a representative experiment in
which CFTR channel activity was high in the cell-attached configuration during forskolin stimulation (indicated by letter
a) and declined spontaneously when
excised into bath solution containing 100 nM PKA (compare with
b and
c). Rundown continued after the
addition of 0.1 and 1.0 µM calyculin A
(d and
e, respectively) but was reversed by
raising the PKA concentration from 0.1 to 0.3 µM
(f). These results confirm that a
fraction of membrane patches from BHK cells contain robust phosphatase
activity resembling that in CHO cells (35). This predominant
phosphatase is insensitive to calyculin A and independent of
Ca2+ and calmodulin and therefore
unlikely to be PP1, PP2A, or PP2B. However, because calyculin A did
slow rundown in 3 of 10 patches, excised membrane patches from BHK
cells can apparently contain some PP1 or PP2A activity that mediates a
small fraction of the deactivation.

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Fig. 5.
Lack of effect of calyculin A on channel rundown in an excised patch.
CFTR channel activity was stimulated by preincubating cells with
forskolin (20 µM). After recording channel activity in cell-attached
configuration (see segment marked
a), patch was excised at 1st arrow
into bath solution containing low PKA activity (100 nM). Rundown was
still observed under these conditions (compare
b and
c) and continued after addition of
100 nM calyculin A (d) and 1,000 nM
calyculin A (e). Decline in channel
activity was caused by dephosphorylation, since it was partially
restored by raising concentration of PKA to 350 nM
(f).
|
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Effect of phosphatases on NPo.
Patches that had stable CFTR activity after 5-min exposure to low PKA
(50-100 nM) were used to test the effects of exogenous phosphatases (solutions 1-3,
Table 1). PP1 (final activity 1.6 pmol/min in 1 ml of bath buffer) did
not affect Po
(P > 0.38), consistent with a
previous study (4). By contrast, addition of PP2A (0.78 pmol/min) under
the same conditions caused a slow decline in
Po from 0.46 to
0.12 within 6.2 ± 0.6 min (P < 0.005; Fig.
6A).
PP2B (0.4-0.8 pmol/min) caused some decrease in
Po, but this just
reached statistical significance (P = 0.04) after a 10-min exposure in solution
2 containing Ca2+
and calmodulin. PP2C was the most potent phosphatase tested, causing
Po to decline
rapidly from 0.48 to 0.11 within 3.4 ± 0.8 min (final activity,
0.16-0.32 pmol phosphate transferred/min; P < 0.0004; Fig.
6B). Alkaline phosphatase at much
higher levels (80 µmol phosphate transferred/min when assayed using
PNP as substrate) caused slow deactivation
(P < 0.002; Fig.
6C) as reported previously (2, 35).
The effects of exogenous phosphatases on
Po are summarized
in Fig. 7.

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Fig. 6.
Patch recordings of PP2A, PP2C, and alkaline phosphatase effects on
CFTR activity in excised patches. After stable channel activity had
been recorded in excised patches for several minutes, exogenous
phosphatases were added at arrows. A:
PP2A caused 95% deactivation of CFTR channel in ~5 min;
B: PP2C reduced channel activity by a
similar amount within ~1 min; C:
deactivation by alkaline phosphatase began after a delay of several
minutes and required ~10 min for activity to decline to low levels.
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Fig. 7.
Summary of PP1, PP2A, PP2B, PP2C, and alkaline phosphatase effects on
open probability
(Po) in excised
patches. Patches were excised from BHK cells into bath solution
containing 100 nM PKA at time 0.
Po was computed
for each 1-min segment. For clarity, values plotted represent
Po determined
during preceding minute (i.e., up to and including time point).
Po of CFTR
channels in most control patches increased gradually and then
stabilized (A;
n = 5). After recording of channel
activity for 5 min, PP1 (C;
n = 5), PP2A
(D; n = 4), PP2B (E;
n = 5), PP2C
(F; n = 50), or alkaline phosphatase (G;
n = 4) was added to bath. PP1 did not
cause induced deactivation under these conditions. PP2A, PP2C, and
alkaline phosphatase deactivated CFTR channels with different time
courses. PP2B caused some deactivation that reached statistical
significance after 10 min. For comparison with spontaneous rundown
occurring in ~25% patches, excisions of patches with rundown are
aligned in B with time at which
exogenous phosphatases were added in other panels to patches having
stable Po. Fast
decline after PP2A (time constant = 5.03 min) or PP2C ( = 0.69 min) was fitted by single exponential functions (dotted
lines in D and
F, respectively). Spontaneous decline
in Po in patches
with rundown was very similar to that induced by addition of exogenous
PP2C ( = 0.69 min).
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The time of excision (for those patches displaying spontaneous rundown;
Fig. 7B) and phosphatase addition
(for those patches with stable channel activity after excision) are
shown aligned in Fig. 7, so that time courses of deactivation by
endogenous and exogenous phosphatases can be compared. Maximal
deactivation by PP2C was reached within 3.4 ± 0.8 min and was well
fitted by a single exponential function having
= 0.69 min (Fig.
7F). This decline was similar to
that observed during spontaneous rundown after excision (
= 0.66 min, Fig. 7B) and was
faster than the deactivation induced by PP2A (
= 5.03 min, Fig.
7D). By contrast, the decline
induced by alkaline phosphatase began after a considerable delay (>6
min) despite higher unit activity.
Effect of PP2A and PP2C on burst duration.
To study phosphatase effects on channel gating, we used patches from
forskolin-stimulated BHK cells having stable channel activity after
excision. Phosphatases were added after recording channel activity for
several minutes (Fig. 8). Representative control data under these conditions are shown in Fig.
3A, and the mean control values are
given in Fig. 4A. Forskolin
stimulation before excision was the only stimulus used to activate CFTR
in these experiments, since exposure of excised patches to both
phosphatase and PKA could complicate interpretation of gating effects.
Both PP2A (Fig. 8A) and PP2C (Fig.
8B) caused deactivation of CFTR channels under these conditions, although the effect of PP2A was slower, in qualitative agreement with the results obtained when low PKA
activity was also present (compare Fig. 7).

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Fig. 8.
Effects of exogenous PP2A and PP2C on CFTR channel activity.
NPo was
calculated before and after addition of PP2A
(A) or PP2C
(B) to patches that had stable
channel activity for at least 1 min after excision from
forskolin-stimulated cells.
(s) was calculated during 2 phases: rapidly declining initial phase
(i) and deactivated phase, during
which NPo was
reduced by >90% (ii). Note
decrease in
o between
phases i and
ii after addition of PP2A
(A). By contrast,
NPo was similar
during phases i and
ii after PP2C addition
(B).
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To examine rundown more precisely, the response to PP2A and PP2C was
divided into two phases: phase i) a
rapidly declining phase immediately after addition of PP2A or PP2C and
phase ii) a stable phase, which was
achieved after channel activity had declined by >90% (see Fig. 8,
A and
B).
o was
analyzed in 5-s segments during each phase. Deactivation by PP2A was
accompanied by a striking decline in burst duration between
phases i and
ii (Fig.
8A; compare
o in
phases i and
ii). No burst lasting >0.8 s was
observed during 30 min of recording with PP2A present (3 patches, 10 min each). By contrast, PP2C induced rapid deactivation without
altering burst duration significantly
(n = 3; Fig.
8B). The distribution of
o in the
presence of PP2C (Fig. 9B)
yielded a
o
during phase ii that closely resembled the one observed during spontaneous rundown (see Fig.
4B). The overall
o was
shorter during exposure to PP2A, reflecting the loss of a population of
long bursts (Fig. 9A). By contrast, long bursts were frequently observed during phase
ii when channels were exposed to PP2C. The
o values in
the presence of PP2A and PP2C are summarized in Fig.
10.

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Fig. 9.
Distributions of
o after
exposure to PP2A or PP2C. Histograms of
o
calculated during each 5-s interval during 2 phases shown in Fig. 8.
Data were pooled from at least 3 patches under each condition.
Distributions from phase i immediately
after PP2A or PP2C addition were both fitted by two Gaussian curves.
A: after prolonged exposure to PP2A
(phase ii), slow
o component
disappeared and bursts in fast component became shorter. By contrast,
prolonged exposure to PP2C did not eliminate slow component or shorten
o
(phase ii of
B).
|
|

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Fig. 10.
o after
channel rundown and phosphatase exposure. Summary of
o calculated
in control patches (no rundown), after activity had declined
spontaneously to <10% of starting value, and immediately after
addition of PP2A or PP2C (phase i)
or after prolonged exposure to PP2A and PP2C when activity had declined
to <10% of starting value (phase
ii).
|
|
The above results indicate that both exogenous PP2A and PP2C can
deactivate CFTR channels, but the rates of deactivation and effects on
single-channel kinetics are different. PP2A causes slow deactivation
and shortening of bursts, whereas PP2C causes rapid deactivation with
no change in burst duration. The rate of decline in channel activity
during exposure to PP2C closely matches that during spontaneous
rundown, further suggesting that PP2C is primarily responsible for
rundown in patches excised from BHK cells.
Effect of
Mg2+ on CFTR
rundown in excised patches from CHO cells.
Unlike other protein phosphatases, PP2C activity strongly depends on
free Mg2+ concentration
([Mg2+]). We examined
the effect of lowering total
[Mg2+] from 2 to 0.5 mM on the rundown of CFTR channels in patches excised from CHO cells
(Fig. 11). This represents a decrease in free [Mg2+] from 1.45 to 0.36 mM, which is expected to reduce PP2C activity by 60-70%
according to previous biochemical studies (8). CHO cells were used for
these experiments because all patches excised from CHO cells displayed
rapid rundown under control conditions. Channel activity was increased
approximately fourfold during the first 5 min of exposure to 0 mM
Mg2+, consistent with the
Mg2+-dependent phosphatase
activity.

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Fig. 11.
Effect of reducing free Mg2+
concentration ([Mg2+])
on rundown of CFTR channel rundown in patches excised from Chinese
hamster ovary cells.
NPo calculated
during 10-s intervals after excision, normalized to value measured in
cell-attached configuration. A:
normalized NPo
under control conditions with 2 mM
Mg2+ in bath solution.
B: same experiments carried out with
0.5 mM free Mg2+. Note that
channel activity was prolonged by reducing
[Mg2+].
|
|
Effect of calyculin A on CFTR rundown in T84 cell monolayers.
If soluble phosphatases are lost from excised patches, studying rundown
might underestimate the importance of cytosolic PP2A in intact cells.
We therefore examined the effect of calyculin A on
Isc across intact
(unpermeabilized and undialyzed) T84 monolayers. Previous studies have
shown that Isc in
this preparation provides a measure of net
Cl
secretion under various
conditions (9) and that Cl
secretion is mediated by CFTR channels (6, 37). Calyculin A (20 nM-100 nM) had no effect on
Isc when it was
added to both sides (Fig. 12). Subsequent
addition of 10 µM forskolin to the serosal side increased
Isc from 3 to 43 µA/cm2 within ~8 min. The rate
and magnitude of the forskolin stimulation were not affected by
calyculin A (Fig. 12).
Isc declined
exponentially back to the baseline level when forskolin was washed out.
The rate of this decline was also unaffected by 20 nM calyculin A (Fig.
12A);
Isc after
forskolin washout was well fitted by a single exponential having the
same time constant in the absence (
= 3.6 ± 0.2 min)
or presence (
= 3.5 ± 0.1 min) of calyculin A. Complete
deactivation of
Isc was observed
after forskolin removal despite the presence of 20 nM calyculin A. Similar results were obtained using T84 monolayers at three different
passages.

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Fig. 12.
Effect of calyculin A on deactivation of short-circuit current
(Isc) across
T84 cell monolayers after forskolin washout.
A: adding calyculin A (20 nM;
n = 12) had no effect on baseline
Isc, stimulation
elicited by forskolin (10 µM), or rate of decay of
Isc after washout
of calyculin compared with control monolayers
(n = 24).
B: adding 100 µM calyculin A at same
time as forskolin was withdrawn did not affect rate of deactivation
(n = 13).
|
|
When forskolin was removed and 100 nM calyculin A was added
simultaneously,
Isc increased
slightly (8%) and then declined rapidly to the same low level as in
control monolayers. Calyculin A (100 nM) had no effect on the rate or
extent of deactivation at concentrations that were 10- to 500-fold
higher than the IC50 for PP2A
(0.5-2 nM) (34). Activation by forskolin was inhibited somewhat at
1 µM (data not shown), presumably a nonspecific effect of high
calyculin A. Nevertheless, even at this toxic concentration of
calyculin A, the decline in
Isc after
forskolin washout was similar to that of control monolayers. These data
in intact (unpermeabilized and undialyzed) T84 cells do not exclude an
accessory role for PP2A but indicate that most CFTR deactivation in T84
cells is mediated by a phosphatase other than PP1 or PP2A, as
previously suggested for CHO (35) and human airway epithelial cells
(2).
 |
DISCUSSION |
The goal of this study was to compare the ability of PP2C and other
protein phosphatases to downregulate CFTR and assess their relative
importance in deactivating CFTR channels in the epithelial cell line
T84. By adding exogenous phosphatases to BHK patches having stable
channel activity, we found that PP2A and PP2C both deactivated CFTR
channels which had been activated in vivo (by forskolin) or in vitro
(by PKA catalytic subunit). The protein phosphatase preparations were
assayed biochemically with a common substrate and patch-clamp solution,
so that comparable levels of PP activity could be used when recording
channel activity.
Evidence that PP2C is the primary phosphatase regulating CFTR in BHK
and T84 cells.
Several results in this study suggest that endogenous PP2C regulates
CFTR in the cells studied: 1)
deactivation induced by exogenous PP2C occurred at the same rate as
that mediated by endogenous, membrane-associated phosphatase;
2) burst duration did not change during deactivation by exogenous PP2C or during spontaneous rundown after excision, whereas exogenous PP2A caused burst shortening; 3) rundown in excised patches and
deactivation after forskolin washout from cell monolayers were both
insensitive to calyculin A; and 4)
spontaneous rundown was
Mg2+-dependent within the
millimolar range, consistent with the known properties of PP2C. Recent
studies (38, 40) indicate that PP2C is expressed in most tissues,
including T84 cells. Our conclusions regarding the role PP2C in
regulating epithelial CFTR are in agreement with recent results of
Travis et al. (38), who found that okadaic acid did not inhibit
deactivation of Cl
current
across permeabilized human airway and T84 cell monolayers. They also
found that macroscopic (CFTR-mediated) current in excised patches was
reduced by ~80% after exposure to recombinant PP2C
. PP2A may play
a larger role in regulating CFTR in cardiac (15) and sweat duct
epithelial cells (29). Distinct effects of PP2A and PP2C on the gating
of single CFTR channels have not been described previously.
Properties of protein phosphatase preparations.
Analysis of the phosphatases by SDS-PAGE indicated that the PP2A and
PP2C preparations contained some impurities (8 and <3%, respectively). However, Western blot analysis with specific antibodies did not reveal any cross-contamination, i.e., no PP2A in the PP2C preparation or PP2C contamination in the PP2A preparation. Moreover, okadaic acid did not affect phosphatase activity of the PP2C
preparation but abolished that of the PP2A preparation, and the PP2C
preparation exhibited no activity in the absence of
Mg2+. We conclude that the PP2A
and PP2C preparations used were functionally homogeneous. PP2A from a
commercial supplier had ~1,000-fold lower activity than the PP2A
preparation used here when it was assayed under the same conditions.
Commercially prepared PP2B was more satisfactory, having low
phosphatase activity in the absence of Ca2+ and calmodulin that was
stimulated 30- to 40-fold by addition of these cofactors.
Dephosphorylation of myosin light chains by PP1 and PP2C was linear
during the first 1-2 min under four assay conditions (high salt,
low salt, Tris-DTT, and Tris only). Protein phosphatase activities were
determined as picomoles of phosphate released from phosphorylated
myosin light chains per minute per microliter at 30°C (Table 2).
All four protein phosphatases were partially inhibited by the standard
high-salt (150 mM NaCl) solution used during patch-clamp experiments.
This inhibition was greatest for PP1 (72%) and least for PP2C (20%).
Although freeze-thaw cycles were avoided with routine handling of the
phosphatases, in control experiments, PP1 activity declined fourfold
within 3 h even when kept on ice.
Phosphatases were assayed in patch-clamp solutions and using the same
substrate, so that comparable phosphatase activities could be tested in
patch-clamp experiments. However, activities could only be
approximately matched, since they depend somewhat on the particular
substrates used in the assays. We used myosin light chains as the
substrate for this standardization rather than phospho-CFTR because of
the difficulty in preparing sufficient quantities of purified,
full-length CFTR. When another phosphatase substrate, phosphorylase
kinase, is used instead of myosin light chains, PP2A is about twofold
more effective than PP2C. Thus more PP2A would have been indicated in
this study if the phosphatase activities had been matched using
phosphorylase kinase. However, during patch-clamp experiments, we
tested PP2A activities that were 2.4- to 4.8-fold higher than those of
PP2C and we still observed slower deactivation; therefore, PP2C is
indeed more efficacious. This issue of substrate specificity is of less
concern when PP2C is compared with PP1 and PP2B, which are five- and
threefold less potent in dephosphorylating myosin light chains,
respectively, since compensating for the use of myosin light chains in
assays would only strengthen our conclusion that PP1 and PP2B are less effective in deactivating CFTR.
Effect of PP2A on burst duration.
Exogenous PP2A caused a remarkable shortening of burst duration when
added to BHK patches. Burst shortening did not occur during the
spontaneous deactivation induced by endogenous phosphatase activity or
after addition of exogenous PP2C, but has been reported previously for
cardiac CFTR where it is a major mechanism by which Po is
downregulated (16). The different results would be reconciled if
cardiac cells had more membrane-associated PP2A activity than the cells
used here. Higher PP2A activity would explain both burst shortening and
the more pronounced inhibition of rundown in cardiac cells by okadaic
acid (15).
[Mg2+] and
relative importance of PP2C and PP2A.
In contrast to the present results, deactivation of macroscopic CFTR
conductance in isolated, permeabilized sweat ducts is prevented by
okadaic acid (29). This could reflect higher PP2A and/or lower
PP2C expression in sweat duct epithelium, or it may be due to the
experimental conditions used. Sweat ducts were treated with
-toxin
to permeabilize the basolateral membrane to small solutes such as ATP,
cAMP, and phosphatase inhibitors. The free [Mg2+] of the bath
solution (exposed to permeabilized membrane) was strongly buffered to
~0.1 mM by high concentrations of ATP, gluconate, and EGTA. Most PP2C
would have been inactive if intracellular [Mg2+] approached this
same low level, which may explain the absence of an okadaic
acid-insensitive component. In a whole cell patch-clamp study of
cardiac cells, in which >50% of the deactivation was okadaic
acid-insensitive, Hwang et al. (15) used pipette solution containing 0.92 mM
[Mg2+]. In our studies
of excised patches, okadaic acid-insensitive rundown was routinely
observed with solutions containing 1-3 mM [Mg2+]. Thus the
relative contribution of PP2C reported for different preparations
correlates with
[Mg2+], although other
explanations for the differences are also possible. Cytoplasmic free
[Mg2+] has not been
measured in the cells used here, but recent estimates for the bulk
cytoplasm of other cells are in the 0.5-1.1 mM range (e.g., for
review, see Ref. 32).
Calyculin A did not affect spontaneous rundown of CFTR channel activity
when added to most patches from BHK cells, although average
Po was higher in
3 of 10 BHK patches continuously exposed to calyculin A, suggesting
that some patches have membrane-associated PP2A (or PP1) activity. More
significantly, in intact T84 epithelial cells, which presumably have
physiological [Mg2+]
and normal phosphatase activities, calyculin A did not inhibit deactivation of CFTR after forskolin was removed. This extends previous
studies of CFTR deactivation in this cell line and suggests that CFTR
channels are downregulated primarily by a calyculin A-insensitive
phosphatase in T84 cells (see also Ref. 38).
Other phosphatases: PP1, PP2B, and alkaline phosphatase.
PP1 is a ubiquitous protein phosphatase that is known to regulate other
ion channels. Exogenous PP1 (1.6-3.2 U/ml) failed to deactivate
CFTR Cl
channels when added
to patches from BHK cells. This result agrees with that of Berger et
al. (4), who found that PP1 (5 U/ml) did not deactivate CFTR when
patches were excised from NIH/3T3 cells and is also consistent with the
insensitivity of
Isc deactivation in T84 monolayers exposed to calyculin A (present study) and okadaic acid (38). Insensitivity to calyculin A in T84 cells excludes a role
for other isoforms of PP1 (PP1
or PP1
), since they would also
have been inhibited.
Exogenous PP2B did not fully deactivate the channel even under optimal
conditions of Ca2+ and calmodulin,
although there was a small decrease in
Po that reached
statistical significance after a 10-min exposure. The normal rundown of
CFTR channels in nominally calmodulin- and
Ca2+-free solution (35) suggests
that PP2B plays little role in deactivating CFTR. Although we found
only weak effects of PP2B in the present study, the data are at least
compatible with some regulation of CFTR by PP2B, particularly when it
is considered that regulation might not be fully recapitulated in
excised patches (e.g., if regulation is indirect or requires a
cytosolic factor or ancillary protein in addition to
Ca2+ and calmodulin). Macroscopic
CFTR currents in patches from NIH/3T3 cells were completely insensitive
to PP2B (4), although the PP2B inhibitors cyclosporin and deltamethyrin
can stimulate CFTR channel activity in cell-attached patches on the
same cells (11). Further studies to determine PP2B expression in
NIH/3T3 cells and the specificity of PP2B inhibitors under
cell-attached recording conditions may resolve those apparently
contradictory findings.
Exogenous alkaline phosphatase deactivated CFTR channels after some
delay, consistent with previous studies from this laboratory (2, 35;
but see Ref. 4). Becq et al. (1) showed that a polyclonal antibody
which recognizes the regulatory region of alkaline phosphatase inhibits
rundown, as do phenylimidazothiazoles such as bromotetramisole (2),
which are potent alkaline phosphatase inhibitors. Bromotetramisole activated wild-type and mutant CFTR channels in cell-attached patches
and inhibited rundown, but the concentration required was several
orders of magnitude higher than needed to inhibit alkaline phosphatase
in biochemical assays. Both alkaline phosphatase and PP2C are
Mg2+-dependent enzymes, and it is
possible that CFTR is downregulated by a protein phosphatase which is
distinct from alkaline phosphatases but which shares some
pharmacological and structural similarities. The effect of
bromotetramisole on PP2C activity has not yet been reported.
PP2A and PP2C act at functionally distinct phosphorylation sites.
The results in this study indicate that CFTR deactivation in several
cell lines, including T84 epithelial monolayers, is mediated primarily
by an okadaic acid- and calyculin A-insensitive phosphatase (35). These
properties and the Mg2+ dependence
of rundown point to PP2C; however, a specific inhibitor of PP2C is not
presently available. About 10% of the PP2C
in human HL-60 cells is
membrane associated (25). Approximately 33% of PP2C
activity in
BHK, CHO, T84, and Calu-3 cells is in the particulate (membrane)
fraction (40). A membrane-bound form of PP2C has also been reported in
Paramecium (18). Although PP2A appears
to mediate little, if any, deactivation in these cells under normal
conditions (i.e., with physiological levels of
Mg2+), neither PP2C nor PP2A was
able to completely deactivate CFTR channels in the present study.
Between 5 and 10% of the initial channel activity persisted in the
presence of each exogenous phosphatase. These results suggest that
multiple phosphatases may be required for complete deactivation,
analogous to previous studies (15). Alternatively, PP2C may need to be
associated with CFTR in the membrane to be fully effective. The
qualitatively different effects of PP2A and PP2C on gating imply that
they dephosphorylate functionally distinct sites on CFTR. Deactivation
by PP2A was accompanied by a reduction in
o. By
contrast, deactivation by PP2C was not associated with burst
shortening, although it caused a similar decline in
Po. The inability
of PP2C to alter burst duration was not due to weaker phosphatase
activity, since PP2C caused more rapid deactivation than did PP2A, and
raising the concentration of PP2C by fourfold still did not result in
shorter bursts (Luo, unpublished observation). As discussed above, the
PP2A and PP2C preparations used in patch-clamp experiments had
comparable phosphatase activities when assayed for their ability to
dephosphorylate myosin light chains. These results suggest that burst
and interburst durations are regulated independently, presumably by
distinct phosphorylation sites.
In this paper, we have focused on the acute regulation of CFTR activity
by dephosphorylation of PKA sites. This regulation is relatively rapid
(usually complete within ~3 min) and corresponds to the rapid rundown
that is observed immediately after excision and can be fully reversed
by PKA (35). It has recently been shown that constitutive PKC
phosphorylation is also required for CFTR to respond to PKA (17). We
found that, once channels were deactivated (due to rapid
dephosphorylation of PKA sites), their ability to respond to exogenous
PKA gradually declined unless PKC was also present in the bath.
Moreover, once CFTR channels became refractory to PKA stimulation
(attained after ~10 min without PKC), their responsiveness could be
fully restored by PKC exposure. The present experiments were not
designed to characterize dephosphorylation of the permissive PKC sites.
If membrane-associated PP2C also mediates regulation of those sites,
the relatively slow decline in responsiveness to PKA implies that PKC
sites are less efficiently dephosphorylated.
It has been proposed that CFTR possesses modulation
(P2) sites, which control
Po and are
dephosphorylated by an okadaic acid-insensitive phosphatase, and
activation (P1) sites, which
convert the channel from an inactive to an active state (15). The
effects of exogenous phosphatases in the present study support the
notion of regulation by particular sites but imply a somewhat different
scheme. Our data suggest that one PKA site or group of sites, which we
call PKAi site(s), controls
interburst duration (i.e., opening rate) and is susceptible to both
PP2A and PP2C. Another PKA site or group of sites, which we refer to as
PKAb site(s), controls burst duration (closing rate) and is susceptible to PP2A but not PP2C. The
PP2C-sensitive sites mediating rapid rundown of
Po and the PP2A-sensitive sites mediating rundown and burst shortening should be
identified and their relationship to
P1 and
P2 sites established. The
differential effects of PP2A and PP2C on burst duration described here
could provide a tool for identifying the site (or sites) that control
the rate of closing from bursts.
 |
ACKNOWLEDGEMENTS |
We thank Jie Liao for excellent technical assistance.
 |
FOOTNOTES |
J. Luo was supported by a Canadian Cystic Fibrosis Foundation
studentship. J. W. Hanrahan is a Medical Research Council (Canada) scientist. This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases grants to J. W. Hanrahan and J. R. Riordan.
Address for reprint requests: J. W. Hanrahan, Dept. of Physiology,
McGill University, 3655 Drummond St., Montreal, QC, Canada H3G 1Y6.
Received 14 October 1997; accepted in final form 29 January 1998.
 |
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