Prostaglandin F2
stimulates CFTR activity by PKA- and PKC-dependent
phosphorylation
Karin A.
Yurko-Mauro1 and
William W.
Reenstra2
Departments of 1 Clinical
Science and 2 Pediatrics, Alfred
I. duPont Hospital for Children, Thomas Jefferson University,
Wilmington, Delaware 19803
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ABSTRACT |
The cystic fibrosis transmembrane conductance regulator (CFTR)
can be activated by protein kinase A (PKA)- or protein kinase C
(PKC)-dependent phosphorylation. To understand how activation of both
kinases affects CFTR activity, transfected NIH/3T3 cells were
stimulated with forskolin (FSK), phorbol myristate acetate (PMA), or
prostaglandin F2
(PGF). PGF
stimulates inositol trisphosphate and cAMP production in NIH/3T3 cells.
As measured by I
efflux,
maximal CFTR activity with PGF and FSK was equivalent and fivefold
greater than that with PMA. Both PGF and PMA had additive effects on
FSK-dependent CFTR activity. PMA did not increase cellular cAMP, and
maximal PGF-dependent CFTR activity occurred with ~20% of the
cellular cAMP observed with FSK-dependent activation. Staurosporine,
but not H-89, inhibited CFTR activation and in vivo phosphorylation at
low PGF concentrations. In contrast, at high PGF concentrations, CFTR
activation and in vivo phosphorylation were inhibited by H-89. As
judged by protease digestion, the sites of in vivo CFTR phosphorylation
with FSK and PMA differed. For PGF, the data were most consistent with
in vivo CFTR phosphorylation by PKA and PKC. Our data suggest that
activation of PKC can enhance PKA-dependent CFTR activation.
protein phosphorylation; iodide efflux; phorbol ester; adenosine
3',5'-cyclic monophosphate; NIH/3T3 cells; chloride
channels
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INTRODUCTION |
CYSTIC FIBROSIS is caused by mutations in the cystic
fibrosis transmembrane conductance regulator (CFTR) protein, a
Cl
channel that is
activated by ATP hydrolysis and protein kinase A (PKA)-dependent
phosphorylation (1, 5, 22). PKA-dependent activation involves
phosphorylation of multiple sites within a highly charged intracellular
domain, the regulatory or R-domain (5). Mutational analysis suggests
that the addition of negative charge induces a conformational change in
the R-domain that is necessary for channel activation (9, 21). However,
these studies have failed to identify specific phosphorylation sites that are required for CFTR activation and suggest that channel activation is due to a nonlocalized decrease in the net charge of the
R-domain. In vivo phosphorylation studies have identified five serines
(S660, S700, S737, S795, and S813) as sites of PKA-dependent phosphorylation (5, 19). Mutations at these sites have been shown to
alter dose-response curves for agonist-dependent CFTR activation but do
not block channel activation (27). In contrast, several kinetic studies
have demonstrated functional differences among, as yet unidentified,
PKA-dependent phosphorylation sites (12, 13). Together, these data
suggest that PKA-dependent CFTR regulation may be more complex than a
two-state model.
In addition to PKA-dependent activation, CFTR can also be activated by
PKA-independent mechanisms. Several studies have demonstrated protein
kinase C (PKC)-dependent CFTR activation and modulation of
PKA-dependent activity. In excised patches from transfected Chinese
hamster ovary cells, PKC increases the open probability of
PKA-activated channels (24). In excised patches from stably transfected
NIH/3T3 (NIH-CFTR) cells, PKC activates CFTR to ~20% of the activity
obtained with PKA (3). In transfected C127 cells, phorbol myristate
acetate (PMA) increases CFTR activity but to levels that are less than
those obtained with cAMP-dependent agonists (8). CFTR activation by
both PKA and PKC has been observed in cardiac myocytes and in
pancreatic acinar cells, but in both systems the effects of CFTR
activation with PKA plus PKC has not been described (7, 16). It has
also been reported that a basal PKC-dependent phosphorylation is
required for PKA-dependent CFTR activity (15). PKC-dependent
phosphorylation has been shown to occur at serines 686, 700, and 790 (10, 19). It can be assumed, but has not been demonstrated, that
phosphorylation at these sites is required for PKC-dependent
alterations to CFTR activity. These results suggest that PKC may
modulate PKA-dependent CFTR activation. This modulatory effect may be
important under in vivo conditions in which agonists that stimulate
both pathways are likely to be present.
To study the effects of the combined activation of PKA and PKC on CFTR
activity, we have examined stimulation by prostaglandin F2
(PGF). PGF-dependent
stimulation was compared with stimulation with forskolin (FSK) and PMA,
activators of PKA and PKC, respectively. In NIH/3T3 cells, PGF
increases intracellular Ca2+ and
inositol trisphosphate (11, 18, 26). It has also been reported, and
confirmed in this study, that PGF increases intracellular cAMP in
NIH/3T3 cells (11). Thus, in NIH-CFTR cells, PGF provides a model for
evaluating the combined effects of PKA and PKC on CFTR activity. PGF
caused maximal stimulation of CFTR activity at a level of cellular cAMP
that was 25% of that required for maximal stimulation with FSK. PMA
increased CFTR activity to ~20% of the maximal activity seen with
FSK without increasing cellular cAMP. Both PMA and PGF had additive
effects on FSK-dependent CFTR activity. PMA- and PGF-dependent CFTR
activity and phosphorylation were inhibited by PKC antagonists.
Phosphopeptide maps of in vivo-phosphorylated CFTR allowed PKA- and
PKC-dependent phosphorylation to be distinguished and
suggested PGF-dependent activation was due to phosphorylation by
both kinases. Our results demonstrate that the combined activation of
both signal transduction pathways allows maximal CFTR activation at
lower cAMP levels than those required for activation solely by PKA.
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METHODS |
Cell culture.
NIH-CFTR and NIH-mock cells, obtained from Dr. Richard Mulligan, were
grown as described previously (2, 14). Briefly, cells were cultured at
37°C with 5% CO2 in DMEM
supplemented with 100 µg/ml penicillin, 50 µg/ml streptomycin, 50 µg/ml gentamicin, and 10% newborn calf serum. For
I
efflux studies, cells
were grown on four-well, 15-mm plates. For in vivo phosphorylation and
cAMP measurements, cells were grown on 35-mm plates; for peptide
mapping, cells were grown on 60-mm plates.
I
efflux.
CFTR activity was assayed by measuring the rate of
I
efflux. Measurements were
performed as previously described (14, 25). Cells, at 37°C, were
incubated for 30 min with efflux buffer (in mM: 141 NaCl, 3 KCl, 2 KH2PO4,
0.9 MgCl2, 1.7 CaCl2, 10 HEPES, and 10 glucose,
pH 7.4) containing 5 µCi/ml carrier-free
125I (sodium salt). For studies at
50 mM extracellular K+, 45 mM NaCl
was replaced with KCl. The loss of intracellular 125I was determined by replacing
the bathing solution with efflux buffer every 60 s for 10 min. Agonist
or vehicle was present at all times after 4 min. When present,
N-[2-p-bromocinnamyl(amino)ethyl]-5-isoquinolinesulfonamide (H-89) was added at the start of the
125I uptake and staurosporine was
added at the start of the measured efflux. Intracellular
125I
was calculated at each time point and rates of
I
efflux
(r) were determined from
r = ln(125It1/125It2)/(t1
t2), where
125It1
and
125It2
are intracellular 125I at
successive time points t1 and
t2. Rates of
I
efflux were time
dependent, and all comparisons were based on maximal values for the
time-dependent rates (peak rates). Dose-response curves were obtained
by fitting data for peak rates to
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where
V0 and
Vmax are the
rates at 0 and infinite agonist concentration, [Ag] is the
concentration of agonist, and
Ka is the agonist
concentration that gives a half-maximal change. Nonlinear least squares
regression was performed with software from Jandel Scientific.
Measurement of cellular cAMP.
Cellular cAMP was determined by radioimmunoassay using a commercial kit
(Amersham). Samples were prepared as recommended by the manufacturer.
Briefly, confluent NIH-CFTR cells were preincubated for 10 min at
37°C with phosphate-free efflux buffer (in mM: 141 NaCl, 5 KCl, 0.9 MgCl2, 1.7 CaCl2, 10 HEPES, and 25 glucose,
pH 7.4). Cells were stimulated with agonist for 2 min, and the reaction was terminated by removing the buffer and adding 500 µl of 70% ethanol at 4°C. Cells were collected and centrifuged at 2,000 g for 15 min, and the supernatant was
recovered. Cells were reextracted with ethanol. The supernatants were
combined and evaporated overnight at 70°C. The residue was
dissolved in 600 µl of 50 mM sodium acetate, pH 5.8, and cAMP
concentrations were determined by radioimmunoassay. cAMP levels were
normalized to total cell protein determined by bicinchoninic acid (BCA)
assay (23).
In vivo CFTR labeling with
32Pi.
In vivo CFTR labeling was performed as previously described
(20). Cells were incubated for 90 min in phosphate-free efflux buffer
containing 0.4 mCi/ml
[32P]orthophosphate
(3,000 Ci/mmol). Cells were washed with phosphate-free efflux buffer
and stimulated with agonist for 2 min. When present, H-89 was added to
the 32P buffer for the final 30 min and was present during treatment with agonist; staurosporine was
added with agonist. After stimulation, cells were lysed with 4°C
RIPA buffer (100 mM NaCl, 50 mM NaF, 0.1% SDS, 1% sodium
deoxycholate, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, 0.1 mM PMSF, 0.1 mg/ml aprotinin, 1 mM orthovanadate, and 50 mM
Tris · HCl, pH 7.5). Lysates were cleared by
centrifugation (100,000 g for 20 min),
and CFTR immunoprecipitated from 1.7 mg/ml supernatant with 2 µg/ml
CFTR monoclonal antibody (MAb; Genzyme, anti-COOH-terminal). CFTR was
resolved by SDS-PAGE, visualized by autoradiography, and quantified by
scintillation counting of excised bands. For two-dimensional (2-D)
phosphopeptide mapping, cells were incubated for 150 min with
phosphate-free buffer containing 2 mCi/ml
[32P]orthophosphate
(3,000 Ci/mmol), and cleared lysates (3.8 mg/ml) were incubated with
7.5 µg/ml anti-CFTR MAb. Protein was determined by BCA assay.
Staphylococcus aureus V8 protease digestion.
V8 Protease digestion was performed as described previously (6,
20). Immunoprecipitated CFTR was excised from gels and placed into the
wells of 20% acrylamide gels with 2.5 units of Staphylococcus aureus V8 protease
(Sigma). SDS sample buffer supplemented with 1 mM EDTA was added, and
samples were run into the stacking gel and digested for 2 h. The
peptides were then separated by electrophoresis, and labeled peptides
were visualized by autoradiography.
Tryptic digestion and 2-D peptide mapping.
2-D phosphopeptide mapping of in vivo-labeled CFTR was
performed as described previously (4). After separation by SDS-PAGE, gel slices containing 32P-labeled
CFTR were rehydrated and suspended in 50 mM
NH4HCO3 (pH 7.4) containing 0.1% SDS and 5%
-mercaptoethanol. BSA (20 µg) was added to the clarified supernatant, and protein was
precipitated with 17% TCA. Pellets were washed with 100% ethanol,
suspended in 50 mM
NH4HCO3,
and digested for 16 h at 37°C with 25 units N-tosyl-L-phenylalanine
chloromethyl ketone (TPCK)-trypsin (Worthington). An additional 25 units of TPCK-trypsin were added, and the digestion continued for 2 h.
Samples were diluted with water and lyophilized to dryness four times.
Peptides were then spotted onto 20 × 20-cm TLC plates (EM
Science), separated by electrophoresis for 30 min at 1,000 V in 100 mM
(NH4)2CO3,
pH 8.9, and followed by ascending chromatography in
n-butanol-pyridine-water-acetic acid,
15:10:12:3 (vol/vol/vol/vol). Phosphopeptides were detected with the
Storm 860 PhosphorImager (Molecular Dynamics) and identified by
comparison with previously published studies (5) and maps provided by Dr. Jonathan Cohn.
Materials.
PGF was obtained from Sigma and dissolved in 100% ethanol; the
final ethanol concentration in all buffers was
0.1%. FSK, PMA, H-89,
and staurosporine were obtained from Alexis. Stock solutions were
dissolved in DMSO so that the final concentration of DMSO in all
buffers was
0.2%. Radiochemicals were obtained from DuPont NEN.
Statistics.
Calculated values are presented as means ± SE. Comparisons were
made among paired data by Student's
t-test, with
P < 0.05 taken as significant.
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RESULTS |
CFTR activation by FSK and PGF.
Kinase-dependent CFTR activation was investigated by assaying
I
efflux and
in vivo CFTR phosphorylation in NIH-CFTR cells. As shown in
Fig. 1, the addition of agonist caused a
time-dependent increase in the rate of
I
efflux. A transient
increase in the rate of I
efflux with a peak rate 2-3 min after agonist addition was
observed with FSK and with PGF. With PMA, a more gradual increase in
the rate of I
efflux was
observed. Mean values for peak rates of
I
efflux were 0.64 ± 0.04 min
1 [no. of
experiments (n) = 15] for 1 µM FSK, 0.25 ± 0.01 min
1
(n = 16) for 200 nM PMA, 0.54 ± 0.04 min
1
(n = 16) for 1 µM PGF, and 0.27 ± 0.02 min
1
(n = 16) for 0.01 µM PGF. Mean rates
in the absence of agonist were 0.12 ± 0.01 min
1
(n = 15). Two tests were performed to
determine if PGF-dependent I
efflux was due to
stimulation of CFTR or another anion channel. 1)
I
efflux was assayed in
mock-transfected NIH/3T3 (NIH-mock) cells. In NIH-mock cells, the
average rate of I
efflux
was 0.16 min
1
(n = 2) in the absence of agonist.
Peak rates in the presence of 1 µM PGF and 1 µM FSK were on average
0.18 min
1
(n = 2) and 0.17 min
1
(n = 2), respectively; neither agonist
caused a significant increase. 2) In NIH-CFTR cells,
I
efflux was assayed in the
presence of dinitrostilbene-2-2'-disulfonic acid (DNDS), a
nonspecific Cl
channel
inhibitor that does not inhibit CFTR from the extracellular side (17).
Peak rates of I
efflux in
the absence of DNDS were 0.49 ± 0.03 min
1
(n = 3) for 1 µM FSK and 0.70 ± 0.08 min
1
(n = 4) for 1 µM PGF. In the
presence of 100 µM DNDS, peak rates were 0.63 ± 0.02 min
1
(n = 3) for 1 µM FSK and 0.76 ± 0.05 min
1
(n = 4) for 1 µM PGF. Together,
these results demonstrate that the PGF-dependent increases in
I
efflux reflect CFTR
activation.

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Fig. 1.
Time courses for agonist-stimulated
I efflux in NIH-CFTR cells.
Confluent monolayers at 37°C were loaded with
125I for 30 min, extracellular
125I was removed, and
I efflux was assayed by
changing bathing medium every 60 s for 10 min. Rates were calculated as
described in METHODS. Agonists
[1.0 µM forskolin (FSK, ), 1.0 µM prostaglandin
F2 (PGF, ), 0.01 µM PGF
( ), 200 nM phorbol myristate acetate (PMA, ), or vehicle (0.2%
DMSO) ( )] were present in bathing medium where indicated by
bar. Data are means ± SE [no. of experiments
(n) 15].
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Although these results demonstrate PGF-dependent CFTR activation,
PGF-dependent activation of a K+
conductance could increase the membrane potential and thereby increase
the rate of CFTR-dependent
I
efflux. To determine if
an increase in K+ conductance
contributed to the PGF-dependent increase in
I
efflux, rates of
I
efflux were measured in
the presence of 5 and 50 mM extracellular K+. If, in the presence of 5 mM
extracellular K+,
K+ channel activation increased
I
efflux by hyperpolarizing
the membrane potential, K+ channel
activation with 50 mM extracellular
K+ would depolarize the membrane
potential and inhibit the rate of
I
efflux. At 50 mM
K+, the rates of
I
efflux as a percentage of
those with 5 mM K+ were 92 ± 11% (n = 3) with 1 µM FSK and 112 ± 3% (n = 4) with 1 µM PGF.
Because PGF-stimulated I
efflux was not reduced with 50 mM extracellular
K+, a PGF-dependent
hyperpolarization of the membrane potential is unlikely to contribute
to the observed PGF-dependent rate of I
efflux.
The dose-response relationship for PGF-dependent activation of
I
efflux was examined. As
shown in Fig. 2, a maximal rate of
I
efflux was seen at 1 µM
PGF with an EC50 of 50 ± 10 nM. Additive effects of PMA and PGF on FSK-dependent CFTR activity were
observed. As shown in Fig. 3, rates of
I
efflux with saturating
concentrations of PGF and FSK (14) were similar [0.66 ± 0.03 min
1
(n = 7) and 0.74 ± 0.04 min
1
(n = 7), respectively]. However,
in the presence of both agonists, the rate of
I
efflux, 1.06 ± 0.04 min
1
(n = 7), was significantly greater
than that with either agonist alone. A similar additive effect on
FSK-dependent I
efflux was
produced with PMA.

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Fig. 2.
Dose-response relationship for PGF-stimulated
I efflux in NIH-CFTR cells.
I efflux was assayed as
described in Fig. 1, and peak rates were plotted as a function of PGF
concentration. Data are means ± SE
(n 2). The line was fit by
nonlinear least squares regression.
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Fig. 3.
Additive effects of PGF and PMA on FSK-dependent
I efflux in NIH-CFTR cells.
I efflux was assayed as
described in Fig. 1, and peak rates for the indicated agonists are
plotted as means ± SE (n 7).
Rates in the presence of 1.0 µM PGF plus 1.0 µM FSK were
significantly greater than those for 1.0 µM PGF or 1.0 µM FSK
alone, ** P 0.01. Rates in
the presence of 200 nM PMA plus 1.0 µM FSK were significantly greater
than those for 200 nM PMA or 1.0 µM FSK alone,
* P 0.05.
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To determine if CFTR activation by PGF involved the activation of PKC,
the effects of the PKA- and PKC-selective inhibitors, H-89 and
staurosporine, on agonist-dependent
I
efflux were determined.
In Fig. 4, rates of agonist-dependent I
efflux were plotted after
subtraction of control rates in the absence of agonist. For each
agonist, data are presented without inhibitor, with 10 µM H-89, and
with 1 µM staurosporine. Neither inhibitor had a significant effect
on the control rate of I
efflux (data not shown). PMA-dependent
I
efflux was significantly
inhibited by staurosporine by 57 ± 6% (n = 6) but was not inhibited by H-89.
A similar pattern was observed with 0.01 µM PGF, in which
staurosporine inhibited I
efflux by 31 ± 6% (n = 8),
whereas H-89 had no effect. In contrast, FSK-dependent
I
efflux was significantly
inhibited by H-89, 30 ± 4% (n = 5), but not by staurosporine. A similar pattern was observed with 1.0 µM PGF: H-89 inhibited I
efflux by 76 ± 5% (n = 3),
whereas staurosporine had no effect. These results suggest that, at low
PGF concentrations, PGF-dependent I
efflux involves
activation of PKC, whereas, at high concentrations, a PKA-dependent
pathway may be involved in channel activation.

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Fig. 4.
Inhibition of agonist-dependent
I efflux by H-89 and
staurosporine (stauro). Rates of
I efflux were assayed with
indicated agonists in the absence (open bars) and presence of 10 µM
H-89 or 1.0 µM staurosporine. In all cases, paired control rates in
the absence of agonist (0.128 ± 0.005 min 1) were subtracted
from rates in the presence of agonist; neither H-89 nor staurosporine
had any effect on the control rate. Data are means ± SE
(n 3). Where indicated (**), rates
in the presence of inhibitor were significantly less
(P 0.01) than the uninhibited
rate.
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Agonist-stimulated cAMP levels.
The role of PKA in PGF-dependent CFTR activation was also assessed by
comparing cAMP levels and rates of
I
efflux. In Fig.
5, rates of PGF- and FSK-dependent
I
efflux were plotted
against cAMP levels obtained under the same experimental conditions.
For both PGF and FSK, rates of
I
efflux were increased
when cAMP levels increased, but the slope of the regression line for
PGF (54 ± 15 min
1 · nmol
1 · mg)
was significantly greater (P < 0.01)
than that for FSK (8.0 ± 0.07 min
1 · nmol
1 · mg).
Thus, at comparable levels of cAMP, PGF produced a greater increase in
channel activity. Two explanations are possible for this result. PGF
may stimulate a localized production of cAMP that is tightly coupled to
PKA-dependent stimulation of CFTR, whereas FSK increases cAMP levels
throughout the cell. This would require a higher level of total cAMP
with FSK than with PGF to achieve the same amount of PKA-dependent CFTR
activity. Alternatively, PGF may activate a cAMP-independent pathway
that enhances PKA-dependent CFTR activation.

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Fig. 5.
Dependence of agonist-stimulated
I efflux on intracellular
cAMP. Rates of I efflux
were assayed as described in Fig. 1, and peak rates were determined. In
parallel experiments, cAMP levels were assayed by radioimmunoassay.
Rates of I efflux with 0, 0.01, 0.1, and 1.0 µM FSK ( ); 0.01, 0.1, and 1.0 µM PGF ( );
200 nM PMA ( ); and 200 nM PMA plus 0.1 µM FSK ( ) were plotted
as a function of intracellular cAMP. Data for both
I efflux and cAMP are means ± SE (n 6). Lines for FSK and
PGF were obtained by linear regression; slopes were significantly
different at P 0.01.
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To determine if PKC enhanced PKA-dependent CFTR activation without
increasing cAMP levels, I
efflux and cAMP were measured during stimulation with 200 nM PMA and
with 200 nM PMA plus 0.1 µM FSK. In both cases, PMA produced a
significant increase in channel activity with no significant increase
in cAMP (Fig. 5). In a similar fashion, 0.01 µM PGF produced a
significant increase in channel activity with no increase in cAMP
levels. These results demonstrate that, in addition to enhancing PKA-dependent channel activation (Fig. 3), PKC can stimulate CFTR without increasing cAMP levels. They also suggest that PGF-dependent CFTR activation might involve the activation of PKC.
In vivo CFTR phosphorylation.
To further examine PGF-dependent CFTR activation, agonist-dependent in
vivo phosphorylation was assayed. A representative autoradiogram of in
vivo-phosphorylated CFTR that was immunoprecipitated and then resolved
by gel electrophoresis is shown in Fig. 6.
Relative to control (lane 1), CFTR
phosphorylation was increased 3.7-fold with 1 µM FSK
(lane 2), 2-fold with 0.01 µM PGF
(lane 3), 3-fold with 1 µM PGF
(lane 4), and 1.6-fold with 200 nM
PMA (lane 6). In the presence of 1 µM PGF plus 1 µM FSK (lane
5), CFTR phosphorylation was 5.7-fold greater than
that of the control. To determine the roles of PKA and PKC in
PGF-dependent CFTR phosphorylation, agonist-dependent phosphorylation
was assayed in the presence of 10 µM H-89 or 1 µM staurosporine. In
general, inhibitor-dependent decreases in CFTR phosphorylation were
smaller than decreases in I
efflux. As shown in Fig. 7, PMA-dependent
phosphorylation was significantly inhibited by staurosporine (38 ± 5%, n = 5) but not by H-89. A similar
pattern was observed with 0.01 µM PGF, in which staurosporine
inhibited CFTR phosphorylation by 27 ± 6%
(n = 4), whereas H-89 had no effect.
In contrast, FSK-dependent phosphorylation was significantly inhibited
by H-89 by 31 ± 11% (n = 6) but
not by staurosporine. For 1 µM PGF, H-89 inhibited phosphorylation by
16 ± 5% (n = 4), whereas 1 µM
staurosporine was without effect. At 10 µM, staurosporine decreased 1 µM PGF-stimulated phosphorylation by 43 ± 16%
(n = 3) (data not shown).

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Fig. 6.
Agonist-dependent in vivo CFTR phosphorylation in NIH-CFTR cells.
Agonist-dependent in vivo phosphorylation and immunoprecipitation were
performed as described in METHODS.
CFTR was resolved by SDS-PAGE and detected by autoradiography. In the
representative autoradiogram, cells were stimulated with no agonist
(lane 1), 1.0 µM FSK
(lane 2), 0.01 µM PGF
(lane 3), 1.0 µM PGF
(lane 4), 1.0 µM PGF plus 1.0 µM
FSK (lane 5), and 200 nM PMA
(lane 6). Data are representative of
3 experiments.
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Fig. 7.
Inhibition of agonist-dependent in vivo CFTR phosphorylation by H-89
and staurosporine. Agonist-dependent CFTR phosphorylation was assayed
with the indicated agonists in the absence and presence of 10 µM H-89
(open bars) or 1.0 µM staurosporine (solid bars). Data are expressed
as a percentage of the uninhibited value for each agonist. Data are
means ± SE (n 3). Statistically
significant inhibitions at P 0.05 (*) and P 0.01 (**) are
indicated.
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V8 protease and trypsin digests of in vivo-phosphorylated CFTR.
The sites of in vivo CFTR phosphorylation were examined to determine if
channel activation with PMA or PGF was due to phosphorylation by
kinases other than PKA. The purpose of these studies was not to
identify specific phosphorylation sites but to determine if the pattern
of phosphorylation with these agonists differed from FSK. In an initial
study, in vivo-phosphorylated CFTR was immunoprecipitated and resolved
by electrophoresis before digestion with
Staphylococcus aureus V8 protease.
After digestion, the resulting peptides were separated by
electrophoresis. Figure 8 shows a
representative autoradiogram of the resolved phosphopeptides. Treatment
of cells with agonist produced three prominent phosphopeptides of
molecular masses of 17.8, 13.3, and 9 kDa. An additional
phosphopeptide with a molecular mass of ~16 kDa was seen when cells
were stimulated with PMA or PGF (lanes
2-5). Although the 16-kDa band was most clearly
seen with PMA (lane 4), the band was
seen in all lanes in which cells were treated with PGF
(lanes 2,
3, and
5) and not in the lane in which
cells were treated with FSK (lane
1). None of these phosphopeptides is thought to be
degradation products of another phosphopeptide, because the peptide
patterns were not altered by changes in the concentration of protease
or the length of digestion (W. W. Reenstra and S. Raman, unpublished
observations). These results suggest that PMA and PGF stimulate CFTR
phosphorylation on at least one site that is not phosphorylated with
PKA. Because the additional peptide observed with PMA and PGF appears
to have the same molecular mass, the data suggest that PGF activates
PKC-dependent phosphorylation.

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Fig. 8.
Staphylococcus aureus V8 protease
digest of in vivo phosphorylated CFTR. In vivo phosphorylated CFTR,
isolated by immunoprecipitation and gel electrophoresis, was digested
with V8 protease, and the resulting peptides were resolved by SDS-PAGE.
Phosphorylated peptides were observed by autoradiography. In the
typical autoradiogram, CFTR was stimulated with 1 µM FSK
(lane 1), 0.01 µM PGF
(lane 2), 1 µM PGF
(lane 3), 200 nM PMA
(lane 4), and 1 µM PGF + 1 µM FSK (lane 5). Molecular
mass markers are indicated, and position of a phosphopeptide observed
after stimulation with PMA or PGF, but not with FSK, is indicated by
arrow. Data are representative of 3 experiments.
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To confirm the results obtained with V8 protease, in vivo
phosphorylated CFTR was also subjected to tryptic digestion and 2-D
peptide mapping. In Fig. 9, representative
2-D phosphopeptide maps are shown for in vivo phosphorylated CFTR after
stimulation with 1 µM FSK
(A), 200 nM PMA
(B), 0.01 µM PGF
(C), and 1 µM PGF (D). Although more than 10 phosphopeptides were resolved, four (phosphopeptides 1-4) were
highly phosphorylated and showed agonist-specific differences. In the
absence of added agonist (data not shown), there was little
phosphorylation of these peptides. CFTR activation with FSK caused
peptides 1 and 2 to become highly phosphorylated. On the basis of
previously published phosphopeptide maps, phosphopeptides 1 and 2 were
assigned to tryptic peptides containing S737 and S795, respectively
(5). Previous studies have shown FSK-dependent phosphorylation of these
serines as well as S660, S700, and S813 (10). In our studies, S813
(peptide 5) was not heavily phosphorylated and we did not observe
phosphopeptides containing S660 and S700. On the basis of previously
published maps, the phosphopeptide containing S660 should be below the
phosphopeptide containing S795 and the phosphopeptide containing S700
should be to the left of that containing S795. In contrast to
stimulation with FSK, stimulation with PMA (Fig.
9B) led to high levels of
phosphorylation on peptides 1 and 2 and on peptides 3 and 4. Phosphopeptide 3 could contain S768, but we have been unable to
identify phosphopeptide 4. With FSK-dependent stimulation,
phosphopeptides 3 and 4 were phosphorylated to far lower levels than
phosphopeptides 1 and 2. Irrespective of the identification of
phosphopeptides 3 and 4, the increased phosphorylation of peptides 3 and 4, relative to that of peptides 1 and 2, was taken as a hallmark of
PKC-dependent CFTR phosphorylation. As shown in Fig. 9,
C and
D, at both low and high PGF
concentrations, peptides 1-4 were phosphorylated. These patterns
are clearly different from that with FSK and resemble that with PMA.
This suggests that PGF-dependent CFTR stimulation involves
phosphorylation by PKC. However, these data do not rule out
PGF-dependent CFTR phosphorylation by PKA but, when combined with the
inhibitor studies at high PGF concentrations (Figs. 4 and 7), are most
consistent with PGF-dependent CFTR activation involving CFTR
phosphorylation by both kinases.

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|
Fig. 9.
Two-dimensional (2-D) phosphopeptide maps of tryptic digestion of in
vivo phosphorylated CFTR. In vivo phosphorylated CFTR, isolated by
immunoprecipitation and gel electrophoresis, was digested with trypsin,
and resulting peptides were separated by electrophoresis and ascending
chromatography. Phosphopeptides were identified with a Storm 860 PhosphorImager (Molecular Dynamics). Peptide maps of CFTR stimulated
with 1 µM FSK (A), 200 nM PMA
(B), 0.01 µM PGF
(C), and 1 µM PGF
(D) are shown. Phosphopeptides
(1-5) and origin are indicated on each map, with protein kinase A
phosphorylation sites indicated. Data are representative of 3 experiments.
|
|
 |
DISCUSSION |
CFTR is activated by PKA-dependent phosphorylation, and there is
growing evidence for PKC-dependent activation (7, 8, 15, 16, 24). In
this study, we have attempted to determine if, under in vivo
conditions, PKC can modulate PKA-dependent CFTR activation. In addition
to examining the effects of FSK and PMA on CFTR activity, we examined
the effects of an endogenous agonist, PGF. The studies were carried out
in transfected NIH/3T3 cells, a cell line in which PGF stimulates both
adenylate cyclase (11) and phospholipase C (11, 18, 26). We have
confirmed that PGF stimulates adenylate cyclase, a result that was not
observed by a recent study (26). It is not known if PGF stimulates both signal transduction pathways by binding to two distinct receptors or if
both signal transduction pathways are coupled to a single receptor.
Irrespective of this, PGF is a physiological agonist that may use both
PKA- and PKC-dependent signal transduction pathways to activate CFTR.
Previous studies in excised patches have demonstrated that
PKC-dependent phosphorylation activates CFTR and has an additive effect
on PKA-dependent channel activity (3, 24). Under in vivo conditions,
PMA stimulates CFTR in transfected C127 cells, cardiac myocytes, and
pancreatic acinar cells (7, 8, 16). Our studies demonstrate that, under
in vivo conditions, PKC phosphorylates and activates CFTR. In addition,
we demonstrate that in vivo phosphorylation by PKC has an additive
effect on PKA-dependent CFTR activity. Our results also suggest that
PGF-dependent CFTR activation involves the stimulation of both PKA- and
PKC-dependent signal transduction pathways.
Our work shows that PGF stimulates CFTR and that the increase in CFTR
activity for a given level of cellular cAMP is greater with PGF than
with FSK. Several explanations are possible for this finding.
1) PGF, as opposed to FSK, might
increase the membrane potential, thereby increasing the rate of
I
efflux without causing a
corresponding increase in channel number and open probability.
2) CFTR activation by PGF, but not
by FSK, might cause a localized increase in cAMP near CFTR. This could allow PGF to stimulate CFTR with less total cellular cAMP.
3) PGF could activate CFTR by a
mechanism that involves the activation of both PKA and PKC. Because we
were unable to observe an effect of extracellular
K+ on PGF-dependent
I
efflux and the sites of
in vivo CFTR phosphorylation with FSK and PGF differ, it is most likely
that CFTR activation by PGF involves stimulation of another kinase
other that PKA. On the basis of the similarities among PGF- and
PMA-dependent channel activation, in vivo phosphorylation, and
inhibition by kinase-selective inhibitors, it is most likely that
PGF-dependent CFTR activation involves PKC.
This study demonstrates CFTR activation by both PKC- and PKA-dependent
signal transduction pathways and demonstrates that when both kinases
are activated there is an additive effect on CFTR activity. The
additive effects of PKA and PKC may be relevant to in vivo channel
activation, where cells are likely to be stimulated by several agonists
that, in concert, activate multiple signal transduction pathways. Our
data also suggest that activation of PKC may amplify PKA-dependent
stimulation at low cAMP levels. Such a mechanism may be important under
physiological conditions in which submaximal levels of cAMP-dependent
agonists produce less than maximal changes in intracellular cAMP.
Recent studies have demonstrated CFTR activation by the PKC-dependent
pathway in several mammalian tissues, including rabbit pancreatic
acinar cells and guinea pig myocytes (7, 16). The effects of the combined activation of both signaling pathways on CFTR activity have
not been described in these systems. Studies of CFTR activation by
agonists, or agonist combinations, that activate several signal transduction pathways will lead to a better understanding of the in
vivo function of this channel.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Jonathan Cohn for providing us with access to data before
publication and Sasikala Raman for expert technical assistance.
 |
FOOTNOTES |
This work was supported by grants from the Cystic Fibrosis Foundation
(K. A. Yurko-Mauro and W. W. Reenstra), Cystic Fibrosis Research (W. W. Reenstra), and the Nemours Foundation (W. W. Reenstra).
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. §1734 solely to indicate this fact.
Address for reprint requests: W. W. Reenstra, Dept. of Clinical
Science, Alfred I. duPont Hospital for Children, PO Box 269, Wilmington, DE 19899-0269.
Received 11 February 1998; accepted in final form 18 May 1998.
 |
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