Download Protein Kinase C inhibits Kv1.1 potassium channel function

Protein kinase C inhibits Kv1.1 potassium
channel function
LINDA M. BOLAND AND KATHARINE A. JACKSON
Department of Physiology and Program in Neuroscience,
University of Minnesota, Minneapolis, Minnesota 55455
Boland, Linda M., and Katharine A. Jackson. Protein
kinase C inhibits Kv1.1 potassium channel function. Am. J.
Physiol. 277 (Cell Physiol. 46): C100–C110, 1999.—The regulation by protein kinase C (PKC) of recombinant voltagegated potassium (K) channels in frog oocytes was studied.
Phorbol 12-myristate 13-acetate (PMA; 500 nM), an activator
of PKC, caused persistent and large (up to 90%) inhibition of
mouse, rat, and fly Shaker K currents. K current inhibition by
PMA was blocked by inhibitors of PKC, and inhibition was
not observed in control experiments with PMA analogs that
do not activate PKC. However, site-directed substitution of
potential PKC phosphorylation sites in the Kv1.1 protein did
not prevent current inhibition by PMA. Kv1.1 current inhibition was also not accompanied by changes in macroscopic
activation kinetics or in the conductance-voltage relationship. In Western blots, Kv1.1 membrane protein was not
significantly reduced by PKC activation. The injection of
oocytes with botulinum toxin C3 exoenzyme blocked the PMA
inhibition of Kv1.1 currents. These data are consistent with
the hypothesis that PKC-mediated inhibition of Kv1.1 channel function occurs by a novel mechanism that requires a C3
exoenzyme substrate but does not alter channel activation
gating or promote internalization of the channel protein.
phorbol ester; Shaker; voltage clamp; Xenopus oocyte
VOLTAGE-GATED K CHANNELS are a diverse family of
proteins that function in the regulation of action potentials, cardiac pacemaking, signal integration, and neurotransmitter release in excitable cells. In nonexcitable
cells, K channels help modulate hormone secretion, cell
volume regulation, cell proliferation, and lymphocyte
differentiation. The highly localized expression of the
different molecular species of K channels (10) suggests
that K channels may be specialized for different cellular functions. However, little comparative information
is available regarding the second messenger regulation
of the many different but related molecular forms of
voltage-gated K channels. An understanding of how the
different K channels are regulated by neurochemicals
is critical to understanding the diverse and important
functions of these channels.
The phosphorylation of K channels by protein kinase
C (PKC) may regulate channel function. In several cell
types, native K channel currents are affected by agents
that directly activate PKC or by receptor systems that
activate PKC through a second messenger cascade. For
example, PKC activation inhibits a K current in cultured endothelial cells (38) and K currents in brain
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stem respiratory neurons (5) but enhances a K current
in ventricular cells (36). In the Xenopus oocyte expression system, several types of recombinant, voltagegated K channels have been shown to be modulated by
PKC (18, 19, 21, 23, 29), and in some cases there is
evidence that the K channel protein is the substrate for
PKC phosphorylation (2, 3, 4, 8). However, the mechanisms and sites of kinase regulation of K channel
activity are not known for many types of cells and K
channels.
Xenopus oocytes are a useful system for studying the
regulation of ion channel activity. The oocytes express a
variety of native components of second messenger
cascades including the G proteins Go and Gq (25),
phospholipase C (14), and PKC (26). Thus the molecular mechanisms of receptor-channel coupling can be
studied by using the coexpression of foreign cell surface
receptor and channel proteins linked by the native
oocyte second messenger pathways.
In the present study, we tested for possible physiological differences in the modulation by PKC of recombinant voltage-gated K channels. We found that PKC
activation had different consequences for K currents
encoded by genes from the Shaker (Kv1), Shab (Kv2),
Shaw (Kv3), and Shal (Kv4) subfamilies. PKC-mediated downregulation of Kv1.1 currents occurs by a
novel mechanism that is inhibited by the C3 exoenzyme
of botulinum toxin without an effect on channel gating
or a significant reduction in K channel protein present
at the membrane surface.
MATERIALS AND METHODS
Expression of K channels. Plasmids containing the cDNAs
encoding voltage-gated K channels were linearized, and
capped RNAs were synthesized in vitro with Ambion (Austin,
TX) RNA polymerase kits. Similarly, plasmids containing
turkey P2Y or human P2U receptor cDNAs (gifts from T. K.
Harden, Univ. of North Carolina, Chapel Hill, NC) were
linearized and transcribed. RNA was purified by use of the
RNAid kit (Bio 101, Vista, CA) and stored at ⫺80°C in diethyl
pyrocarbonate (DEPC)-treated water. Transcription reaction
products were checked for size by agarose gel electrophoresis,
and RNA concentrations were determined by spectrophotometry (Beckman DU-7000) by using the equation 1 A260 unit ⫽
40 µg/ml RNA, where A260 is absorbance at 260 nm. K channel
cDNAs were generous gifts from M. Covarrubias (mKv4.1), R.
Joho (rKv2.1), O. Pongs (rKv1.1 and rKv3.4), L. Salkoff
(rKv4.2), and B. Tempel (mKv1.1). We also used ShakerH4
from Drosophila melanogaster and a deletion mutant Sh⌬
(residues 6–46 deleted) that removes fast, N-type inactivation (13).
Oocytes were harvested from Xenopus laevis (Xenopus I,
Ann Arbor, MI), previously injected with human chorionic
gonadotropin. Female frogs were anesthetized by immersion
in 0.2% 3-aminobenzoic acid ethyl ester (Sigma Chemical).
0363-6143/99 $5.00 Copyright r 1999 the American Physiological Society
K CHANNEL INHIBITION BY PKC
Oocytes were released by gentle agitation for 2 h in 1 mg/ml
collagenase D (Boehringer Mannheim) dissolved in a Ca-free
OR-2 solution containing (in mM) 96 NaCl, 2 KCl, 1 MgCl2,
and 5 HEPES, pH 7.6, with NaOH. After the first hour, the
solution was replaced with fresh enzyme. Subsequently,
oocytes were extensively washed with Ca-free OR-2, and
stage V and VI oocytes were collected with a dissecting
microscope. Defolliculated oocytes were injected the same day
or the following day with 50–100 nl of RNA (0.01–0.3 ng/nl)
dissolved in DEPC-treated water. Oocyte survival was best in
a frog Ringer solution of (in mM) 96 NaCl, 1 KCl, 0.75 CaCl2,
0.8 MgCl2, and 10 HEPES, pH 7.4, with NaOH, 50 U/ml
penicillin G, and 50 µg/ml streptomycin. Oocytes were maintained at 16–18°C in this solution (changed daily) for 1–6
days before electrophysiological recording.
Mutagenesis. Mutations were introduced into the mKv1.1
K channel cDNA in a Bluescript KS(⫹) vector (Stratagene) by
oligonucleotide-directed mutagenesis. Miniprep DNA was
prepared from isolated colonies with a DNA isolation kit
(Promega). Mutant DNA was identified by dideoxy sequencing with Sequenase-modified DNA polymerase (United States
Biochemical) and electrophoresis on 6% polyacrylamide urea
sequencing gels. For each site studied, two mutant colonies
were sequenced and prepared and tested separately.
Electrophysiology. Macroscopic K currents from oocytes
were recorded with a two-electrode voltage clamp equipped
with an OC-725B amplifier (Warner Instrument, Hamden,
CT). Voltage-measuring and current-passing electrodes were
filled with 3 M KCl and had resistances between 0.3 and 1.1
M⍀. Currents were sampled at 5 kHz and filtered at 2 kHz
with an eight-pole Bessel filter (Frequency Devices, Haverhill, MA). All recordings were done at room temperature
(⬃20°C). Oocytes were clamped at ⫺70 or ⫺80 mV (as noted
in figure legends) and pulsed every 1–5 s to various test
potentials to activate K channels.
Oocytes were perfused continuously with an external solution containing (in mM) 96 NaCl, 2 KCl, 1.8–2 CaCl2, 1
MgCl2, and 5 HEPES, pH 7.4, with NaOH. External solution
changes were made by manually switching a valve that
controlled gravity-driven solution flow across the oocyte
recording chamber. Data were recorded on 80586-based computers equipped with National Instruments AT-MIO boards
and interfaced with the experimental setup. Data acquisition
software was written and generously donated by G. Yellen.
Data were transferred to commercially available software
programs for analysis and the production of figures. Data are
expressed as means ⫾ SE.
Biotinylation and recovery of surface proteins. Kv1.1expressing oocytes (54–60/treatment) were washed twice
with 2 ml of frog Ringer solution and then incubated in a
labeling solution of 1 mg/ml EZ-Link NHS-SS-biotin [sulfosuccinimidyl 2-(biotinamido) ethyl-1,3-dithiopropionate; Pierce,
Rockford, IL] dissolved in frog Ringer solution, pH 7.4. We
used modifications to the methods of Stern-Bach et al. (28).
Cells were labeled for 2 h at 21°C with gentle mixing on a
laboratory shaker and then washed twice for 10 min each in
frog Ringer solution with gentle mixing. Biotin-labeled oocytes were then washed once with 1 ml ice-cold homogenization buffer (20 mM Tris · HCl, 200 mM NaCl, pH 7.4) containing a protease inhibitor cocktail: 1 mM phenylmethylsulfonyl
fluoride (PMSF), 2 µg/ml aprotinin, 1 µM pepstatin (all from
Boehringer Mannheim), and 2 µg/ml antipain and 1 µg/ml
leupeptin (both from Sigma). Oocytes were then homogenized
by several passages through a pipette tip in 200 µl ice-cold
homogenization buffer with protease inhibitors. Membranebound proteins were solubilized by 1.5% Triton X-100 (Sigma).
Samples were incubated for 1 h on ice and centrifuged for 30
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min (16,000 g at 8°C). The supernatant was carefully removed to a fresh tube. A 5-µl aliquot was removed and
represented the total protein (T) fraction. The remaining
supernatant was incubated for 2 h with a 20-µl bed volume of
UltraLink immobilized monomeric avidin beads (Pierce; 50%
slurry equilibrated in homogenization buffer with 1.5% Triton
X-100) with gentle mixing on a laboratory rotator at 8°C. The
resulting biotin-avidin complex was washed five times by
suspension in 1 ml homogenization buffer (with the protease
inhibitor cocktail and 1.5% Triton X-100). The samples were
centrifuged in a PicoFuge (Stratagene) (1 min, 325 g), and the
supernatant was carefully removed. Membrane proteins were
released from the biotin-avidin complex by adding 25–50 µl
loading buffer with reducing agent [0.13 M Tris · HCl (pH 6.8),
10% glycerol, 2% SDS, 150 mM dithiothreitol, 0.01% bromphenol blue] and gently mixed for 15 min at 21°C. Beads were
removed by centrifugation for 2 min at 10,000 g. The resulting supernatant was collected as the membrane-bound protein (M) fraction. In some experiments, protein concentration
was determined by the bicinchoninic acid protein assay
(Pierce), according to the manufacturer’s instructions.
SDS-PAGE and Western blots. The oocyte lysates were
denatured in Laemmli sample buffer (161–0737; Bio-Rad) for
5 min at 100°C and electrophoresed on an SDS-7.5 or 10%
polyacrylamide gel. To obtain a sufficient signal on the
Western blot, we generally loaded the gel with 1–2 µl of the T
fraction (represents total Kv1.1 protein in 0.25–0.5 oocytes)
and the full volume of the M fraction (represents membrane
Kv1.1 protein in 50 oocytes). After electrophoresis, separated
proteins were transferred to an Immobilon-P polyvinylidene
difluoride (PVDF) membrane (Millipore, Bedford, MA) by
electroblotting. Rainbow colored protein molecular weight
markers (Amersham Life Sciences) were used.
The PVDF membrane was rinsed in distilled water and
blocked with 4% Blotto [4% nonfat dried milk in Tris-buffered
saline, pH 7.6, with 0.1% Tween 20 (TBS-T)] for 1 h at room
temperature or overnight at 4°C. The blots were incubated
with primary antibodies in sealable pouches. We used 1 µg/ml
anti-rat Kv1.1 monoclonal antibody (Upstate Biotech or gift
from J. Trimmer), diluted in 4% Blotto, for 1 h at room
temperature on a shaking platform. The anti-Kv1.1 antibody
was raised against residues 458–476, part of the putative
intracellular COOH terminus of rat brain Kv1.1. After primary antibody incubation, the blot was washed five times in
TBS-T for 5 min each. The blot was then incubated with the
secondary antibody for 1 h on a shaking platform in a Pyrex
dish. We used a 1:80,000 dilution of goat anti-rat IgG conjugated with horseradish peroxidase (Sigma) and diluted in 4%
Blotto. Finally, the blot was washed three times in TBS-T for
10 min each. Immunolabeled proteins were detected by the
ECL Plus enhanced chemiluminescence detection system
(Amersham Life Science). X-ray films were scanned and
analyzed with a model GS-700 scanning densitometer and
Molecular Analyst software (Bio-Rad).
A crude synaptosomal membrane fraction was prepared
from freshly dissected adult mouse brains as described by
Trimmer (33), and an aliquot (6 µg protein/lane) was included
as a positive control for Kv1.1 expression. Extracts from
uninjected oocytes do not react with the Kv1.1 antibody (data
not shown).
We probed the M and T fractions with an anti-actin
primary antibody (Sigma) that specifically recognized an
⬃46-kDa band present in the mouse brain extract and in
purified actin, run as a positive control for the antibody. By
Western blotting, native oocyte actin was detected only in the
T fractions, not in the M fractions prepared from either
uninjected oocytes or Kv1.1-expressing oocytes (data not
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K CHANNEL INHIBITION BY PKC
shown). We probed for actin because it is a ubiquitous
cytoskeletal protein that is known to associate with some
plasma membrane proteins, but actin itself is not an integral
membrane protein. Thus the lack of anti-actin reactivity in
the M fraction is consistent with a lack of contamination of
the M fraction with proteins not embedded in the plasma
membrane.
Drugs. Phorbol 12-myristate 13-acetate (PMA), phorbol
12-monomyristate (PMM), 4␣-phorbol, and staurosporine were
from Sigma Chemical. Phorbol esters were prepared as stock
solutions in DMSO and then diluted into standard bath
solution before recording; the final concentration of DMSO
was ⱕ0.1%. Bisindolylmaleimide I and ATP (Na⫹ salt) were
from Boehringer Mannheim. For pharmacological experiments, staurosporine (prepared in DMSO) and the C3 exoenzyme of botulinum toxin (prepared in DEPC-water) were
injected into oocytes 2 days after injecting channel RNA. On
the basis of an estimate of 500 nl for oocyte volume, staurosporine was injected to achieve a final (estimated) cytoplasmic
concentration of 0.9 µM. C3 exoenzyme was injected at
50–100 pg/oocyte. Control experiments for staurosporine
included 0.1% DMSO-injected oocytes. Control experiments
for C3 exoenzyme included injection of an equal concentration
of C3 exoenzyme that had been heated to ⬎110°C for 2 h and
allowed to cool to room temperature before injection. Oocytes
injected with C3 or ‘‘heat-inactivated’’ C3 were then incubated
at different temperatures before electrophysiology, as described in RESULTS.
RESULTS
PMA application inhibits Shaker K currents. Bath
application of the PKC activator PMA (500 nM) caused
large inhibition of Shaker subfamily K channel currents expressed in frog oocytes (Fig. 1A). PMA applied
to oocytes expressing mouse Kv1.1 (n ⫽ 8), rat Kv1.1
(n ⫽ 7), Drosophila Sh⌬ and ShakerH4 (n ⫽ 4), and
mouse Kv1.3 (n ⫽ 3) caused nearly complete inhibition
in some cases, with an average of 73% current inhibition. The time courses and magnitudes of the inhibition
were the same regardless of the Shaker channel species. Voltage-gated K channels from the Shab (Fig. 1B)
and Shal (Fig. 1C) subfamilies were much less affected
by PMA application than were K channels from the
Shaker subfamily (Fig. 1D). At the membrane potential
indicated in the legend for Fig. 1, levels of current
inhibition were compared, either at the peak current
(for inactivating currents) or at the end of a 60- to
80-ms pulse if currents lacked N-type inactivation.
Because of the selectively large inhibitory effect of PMA
on Shaker subfamily K currents, we further studied the
inhibition of mKv1.1 K channels.
Inhibition of K current by PMA always began after a
short delay (Fig. 2A), probably limited by the permeation of PMA through the plasma membrane and the
accumulation to an unknown concentration within the
cell. Drug application was always marked when the
drug solution was switched on. For comparison of
the rates of PMA effect, in this perfusion system, K
channel block by 30 mM external tetraethylammonium
(TEA) is maximal within 10 s [time constant for block
(␶on) ⫽ 3.1 ⫾ 1.1 s; n ⫽ 3]. The time course of current
inhibition by PMA followed a characteristic pattern
with a half time to maximal effect of 8.2 ⫾ 0.6 min (n ⫽
8 representative cells) and a maximal effect after ⬃15
min of external washing of the oocyte. Short applications of PMA were sufficient to produce this longlasting inhibition of K current [Fig. 2A; see also Fig. 6A
(curve labeled ⫹C3, 16°C)], which did not recover in up
to 2 h, the longest time period recorded. Over the course
of hour-long recordings, K current inhibition was never
seen in control oocytes not treated with PMA. The PMA
time course (Fig. 2A) suggests that K channel activity
was steadily and slowly decreasing, and the pattern
was not indicative of sudden oocyte death or perfusion
artifacts. K current inhibition was also observed with
shorter applications of higher concentrations of PMA or
longer applications of lower concentrations. For comparison and analyses, we standardized the PMA applications at 500 nM for 1 min and measured the steadystate current inhibition 15 min after the PMA
application, with test pulses to 0 or ⫹20 mV. We were
careful to perform all experiments on defolliculated
oocytes because, in early experiments, we noticed that
1-min applications of 500 nM PMA were without effect
on mKv1.1 channels expressed in oocytes with intact
follicle cells.
Pharmacological specificity. Phorbol esters such as
PMA are membrane-permeant drugs that bind with
high affinity to the diacylglycerol site of PKC. To test
whether the inhibition of mKv1.1 K channel currents
by PMA was due to the activation of native oocyte PKC,
we used inactive phorbols and inhibitors of PKC activation. Two structural analogs of PMA that do not activate PKC in in vitro phosphorylation assays (for review
see Ref. 22), 4␣-phorbol and PMM, were ineffective
inhibitors of K channel current (Fig. 2, B, C, and F).
The analog 4␣-phorbol always caused a small and
reversible inhibition of current (Fig. 2B), unlike the
persistent inhibition produced by PMA (Fig. 2A). PMM
caused a small but insignificant (unpaired t-test; P ⬍
0.05) inhibition of mKv1.1 K current 15 min after a
1-min application of 500 nM drug (Fig. 2, C and F).
Prior and concurrent bath application of the specific
PKC inhibitor bisindolylmaleimide (31) prevented the
large and persistent inhibition of K current by PMA
(Fig. 2F). In some oocytes treated with the inhibitor, we
also tested PMA applications longer than 1 min (Fig.
2D) and still did not observe a significant inhibition, as
seen for PMA application to oocytes not treated with
PKC inhibitor (compare with Fig. 2A). Staurosporine, a
nonspecific kinase inhibitor, is also a potent inhibitor of
PKC (30) and was injected into channel-expressing
oocytes 1–2 h before the experiment to achieve an
estimated concentration of 0.9 µM in the oocyte (see
MATERIALS AND METHODS ). Injection of oocytes with
staurosporine also blocked the K current inhibition by
PMA (Fig. 2E). Taken together (Fig. 2F), these pharmacological data support the idea that the inhibition of K
current by PMA application is due to the activation of
PKC.
Receptor-dependent coupling of PKC to K currents.
The inhibition of Shaker subfamily K channel currents
by PMA persisted with up to 2 h of washing. This might
be explained by either the persistent activation of PKC
K CHANNEL INHIBITION BY PKC
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Fig. 1. PMA inhibition (⬎PMA) of different K channels.
Acute application of phorbol 12-myristate 13-acetate
(PMA) largely inhibits K channel currents from Shaker
subfamily (A). K channel currents from mouse, rat, and
fly clones are shown. Sh⌬ is deletion mutant of ShakerH4 (ShH4) with N-type inactivation removed. For
comparison, the effect of PMA on K channels from Shab
(B) and Shal (C) subfamilies are shown. D: comparison
of percent inhibition of peak K channel current for the
three subfamilies. Data are means ⫾ SE for n cells. PMA
inhibition was measured 15 min after a 1-min bath
perfusion with 500 nM PMA. Oocytes were clamped at
⫺80 mV and stepped to 0 or ⫹20 mV.
by PMA that accumulates and cannot be washed from
the oocyte membrane or the presence of a stable
phosphorylation event. To distinguish between these
ideas, we studied K channel modulation by receptorstimulated PKC activation. We coinjected oocytes with
RNA encoding the mKv1.1 K channel and a P2Y purinergic receptor, previously shown to activate phospholipase C in stably transfected astrocytoma cells (11). In
coinjected oocytes (Fig. 3A), the P2Y agonist ATP (100
µM) mimicked the effect of PMA inhibition of mKv1.1 K
current (n ⫽ 6). ATP had no persistent effect on mKv1.1
K channels in oocytes injected only with channel RNA
(Fig. 3B; n ⫽ 4 cells).
An unexpected finding was that the inhibition of K
current by activation of P2Y receptors required ⬃10 min
to reach maximal effect, was nearly complete, and
persisted after ATP was washed off, similar to the time
course and long-lasting effects of the PMA-induced
inhibition. This was true even for short applications of
ATP that were followed by up to 30 min of washing.
Unlike PMA, which could accumulate in the oocyte,
water-soluble compounds such as ATP should wash
completely from the perfusion chamber. G proteincoupled phospholipase C activation would be expected
to be a transient event, dependent on agonist-receptor
binding. The K channel block by TEA, in comparison, is
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K CHANNEL INHIBITION BY PKC
Fig. 2. Pharmacological specificity of inhibition of Kv1.1
current. Current amplitude time courses for 5 different
oocytes expressing Kv1.1 currents are shown. A: protein
kinase C (PKC) activator PMA (500 nM) was applied by
bath perfusion for 1 min, followed by washing. Phorbol
analogs 4␣-phorbol (B) and phorbol 12-monomyristate
(PMM; C) were also applied at 500 nM by bath perfusion
for 1 min. D: the kinase inhibitor bisindolylmaleimide
was applied before and concurrent with a long 7.5-min
bath application of 500 nM PMA. E: effect of bath
application of PMA (500 nM, 1 min) for an oocyte
previously injected with staurosporine (see MATERIALS
AND METHODS for details). All oocytes were clamped at
⫺70 to ⫺80 mV, and K currents were measured with
100-ms pulses every 5 s to 0 mV. Occasional interruptions to this protocol were made to run other currentvoltage protocols. F: composite pharmacological data.
Bars, mean percentages of K current (⫾SE) inhibited 15
min after start of 1-min bath application of phorbol
ester. Results from mKv1.1, rKv1.1, and Sh⌬ were not
different and were combined for analysis. We tested
following numbers of cells: PMA, 22; PMM, 6; 4␣phorbol, 6; bisindolylmaleimide, 3; and staurosporine, 4.
fully reversible within 30 s in the same recording
system [time constant for recovery from block (␶off) ⫽
8.0 ⫾ 1.2 s; n ⫽ 3]. So the ongoing K current inhibition
in the absence of ATP might be explained by the
initiation of a PKC-dependent event(s) that cannot be
reversed or halted under these experimental conditions. The stable event could be phosphorylation in the
absence of an appropriate phosphatase required to
dephosphorylate the protein substrate. These data
suggest that the delayed time to reach steady-state
inhibition and the persistence of the inhibition are not
simply explained by a prolonged accumulation of hydrophobic PMA in the oocyte membrane.
Current gating after inhibition by PKC. We plotted
conductance-voltage (G-V) relationships and scaled current traces during the inhibition of K current to study
the impact of PKC activation on current gating. We
observed overlap of the control and inhibited-K-current
G-V profiles. The large inhibition of the K current by
PMA (Fig. 4A) or ATP application to cells coexpressing
K channels and P2Y receptors (Fig. 4B) was not accompanied by a significant change in the voltage range of
activation or the slope of the activation curve. Furthermore, mKv1.1 K channel currents were inhibited by
PKC activation without a significant effect on current
kinetics (Fig. 4C). Scaling the inhibited current to
match the steady-state control current showed that
inhibition by PMA did not alter macroscopic activation
kinetics (Fig. 4D). This demonstrates that the large
inhibition measured cannot be simply explained by an
effect on K channel activation gating. This is in contrast
to other reports that PKC phosphorylation may change
the voltage dependence of ion channel activation gating, possibly through direct phosphorylation of the
channel protein (1, 24). With activating pulses up to
10 s long, PMA inhibition also did not alter C-type
K CHANNEL INHIBITION BY PKC
Fig. 3. Functional coupling of K current inhibition to purinergic
receptors. Xenopus oocytes were injected 2 days before recording with
a combination of RNAs for mKv1.1 and purinergic P2Y receptors (A)
or RNA encoding mKv1.1 alone (B). K current amplitude time course
is shown for 1-min bath applications of 100 µM ATP. Test pulses were
to 0 mV from a holding potential of ⫺80 mV. Inset: magnitude
(mean ⫾ SE) of K current inhibition by 100 µM ATP was measured 13
min after washing for cells coexpressing K currents and P2Y purinergic receptors (hatched box; n ⫽ 6) or cells expressing only K currents
(mKv1.1 only; n ⫽ 4).
inactivation rates (data not shown). In ShakerH4 channels with intact N-type inactivation (Fig. 1A) and in
fast-inactivating channels of the Shal subfamily (Fig.
1C; rKv4.2), PMA inhibition also did not change the
rate of inactivation (see also Ref. 21).
PKC activation does not inhibit all K channel subfamilies. PKC activation does not inhibit to the same degree
all recombinant voltage-gated K channels expressed in
oocytes. As reported previously for a human Shaw
subfamily K channel (8), we confirmed that PMA
application largely removed the N-type inactivation of
rKv3.4 (in 5 of 5 cells; data not shown). In coinjected
oocytes, we found that functional coupling to purinergic
receptor activation largely removed the N-type inactivation of rKv3.4 (in 3 of 3 cells; data not shown). Because
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this PKC-mediated pathway has been well studied (2,
8), it provided an interesting comparison for the time
course of the PKC-mediated event we studied. We
observed that the inhibition of inactivation of rKv3.4 by
either PMA or purinergic receptor activation required
5–10 min to reach maximal effect and was a relatively
stable event, similar to the PKC-dependent inhibition
of Shaker subfamily K channels. Thus the time courses
and levels of stability of the PKC phosphorylation
events for the different groups of oocytes studied may
be similar even though PKC activation may have
different consequences for the function of different K
channel proteins. In cells expressing multiple types of
K channels, PKC activation could be an important and
complex process for regulating cellular excitability and
signaling.
Site-directed mutagenesis of PKC phosphorylation
sites. We identified only three potential PKC phosphorylation sites on mKv1.1 channels, based on the consensus sequence identified by Woodgett et al. (37): Ser/ThrX-Lys/Arg, where X is generally an uncharged residue.
Residues 318 and 322 in mKv1.1 channels are present
in the S4-S5 linker, and residue 442 is in the S6 region,
all thought to be intracellular in current working
models of the channel. The PKC phosphorylation sequence at Thr-318 is conserved in most voltage-gated K
channels (except Kv1.5 and Kv1.6), whereas the phosphorylation sequence at Ser-322 is conserved throughout the Shaker subfamily and in Drosophila Shab and
Shaw. The phosphorylation sequence at Ser-442 in
mKv1.1 is shared only by Kv1.1 channels of the Shaker
subfamily.
Using mutagenesis experiments, we tested the hypothesis that PKC inhibition of mKv1.1 K channel
current is due to phosphorylation of one of these three
consensus PKC sites, present in four-fold symmetry in
the homotetrameric K channel. We replaced, one at a
time, the native serine or threonine residue at position
318, 322, or 442 in mKv1.1 with alanine or valine.
These conservative substitutions remove the PKC phosphorylation sequence. The mutant channels were sequenced, expressed, and tested for inhibition by PMA.
Channel mRNAs for T318A and S442A mutants gave
rise to K currents with no functional differences in
macroscopic gating or current stability recorded. In
addition, both mutants demonstrated K current inhibition after PMA application, with effects not significantly different (unpaired t-test) from those found in
control experiments. K currents from the T318A mutant were inhibited by 83.3 ⫾ 6.8% (n ⫽ 6), and
currents from the S442A mutant were inhibited by
69.5 ⫾ 6.3% (n ⫽ 5). In addition, channels expressing
both the T318A and S442A mutations were also inhibited by PKC activation (76.8 ⫾ 10%; n ⫽ 5). For
comparison, wild-type mKv1.1 currents were inhibited
by 73.4 ⫾ 5.2% (n ⫽ 8). The only difference in these
mutant channels was a current amplitude smaller than
that for wild-type channels, after injections of the same
quantity of RNA (generally 2- to 3-fold-smaller currents). The mutant S322A did not express functional K
channel currents, even though all three of the mutant
C106
K CHANNEL INHIBITION BY PKC
Fig. 4. Comparison of conductance-voltage (G-V)
and current kinetics. A: G-V relationships for
control K current and for mKv1.1 K currents
after submaximal (35 or 60%) inhibition by PMA
(500 nM, 1-min application). B: G-V relationships
for control K current and for K current after
maximal (84% in this cell) inhibition by ATP (100
µM; 1-min application) in an oocyte coinjected
with RNAs for mKv1.1 and P2Y receptors. Conductances were normalized to the value at ⫹20 mV
and were calculated from the equation G ⫽
[I/(Vm-EK )], where I is the peak current at command voltage (Vm ) and EK is the equilibrium
potential (mean EK ⫽ ⫺71 mV from tail currents). EK was calculated from oocytes with small
K currents, for which holding potential quickly
returns to Vm. G-V data were fitted to single
Boltzmann functions with voltage required for
half-maximal activation (V½; midpoint) ranging
from ⫺30 to ⫺33 mV and k (slope factor) ranging
from 7.0 to 10 mV for fitted curves. C: in another
cell, the control K currents before application of
PMA (500 nM, 1 min), when ⬃50% inhibited
(8-min wash), and after maximal inhibition (20min wash) are shown on the same scale. Currents
shown are from ⫺70 mV to a test potential of ⫹20
mV. D: K current at 8 min is scaled to match
maximal steady-state amplitude of control current to compare current kinetics.
RNAs directed membrane protein expression that could
be identified on a Western blot with a Kv1.1-specific
primary antibody (not shown). We conclude that at
least two of the three consensus PKC phosphorylation
sites are not required for the PKC-mediated inhibition
of mKv1.1 K current. We cannot rule out the possibility
that additional, unidentified phosphorylation sites exist in the native, folded protein.
Assay for channel internalization. As shown in Fig. 4,
the inhibition of mKv1.1 K channel current was not
accompanied by changes in the channels’ G-V relationship or kinetics of activation, suggesting that the
channels that contribute the remaining current are not
functionally different from the channels that account
for the inhibited current. Instead, the inhibition of K
current may reflect a reduction in the number of
functional K channels by, for example, downregulating
the K channels from the membrane surface. Indeed,
several pumps, transporters and ion channels (7, 9, 34),
and G protein-coupled receptors (12, 35) are thought to
be up- or downregulated by insertion or internalization
from the plasma membrane, and these processes may
also require a phosphorylation event.
The possible movement of biotin-labeled Kv1.1 K
channels from the membrane surface to a nonmembrane (cytosolic) compartment was determined by avidin recovery of the biotinylated surface proteins and
subsequent immunoblotting for Kv1.1 protein. Groups
of oocytes were placed in single wells of a 24-well plate
and exposed for 2 min to one of the following, dissolved
in extracellular recording solution: 0.01% DMSO, 500
nM 4␣-phorbol, or 500 nM PMA. The pharmacological
treatments paralleled the physiological experiments,
although we used a 2-min drug application because the
oocytes were in a static bath rather than a perfusion
stream. Over the next 30 min, oocytes were washed a
minimum of 7 times by exchanging the drug solution
for normal recording solution. Thirty minutes after
beginning the washing, 4–7 oocytes from each drug
treatment and untreated controls were randomly selected and tested for current magnitude. The remaining intact oocytes were incubated with NHS-SS-biotin,
a membrane-impermeant reagent, to label Kv1.1 channels and other proteins also present at the cell surface.
After being labeled, 50 intact oocytes were washed, and
total protein extracts or membrane extracts were prepared (see MATERIALS AND METHODS ). Samples were
electrophoresed, blotted to nitrocellulose, and probed
with an antibody specific for Kv1.1 protein. The antibody recognizes a sequence present in the COOHterminal region that is thought to be intracellular.
No detectable differences were found in comparing
the densities of Kv1.1 protein in membrane fractions
from oocytes treated with DMSO alone, 4␣-phorbol, or
PMA (Fig. 5A), even though oocytes from the very same
treatment wells showed a ⬎90% inhibition of Kv1.1 K
current by PMA (Fig. 5B). To confirm that the membrane extracts can be used to detect different levels of
K CHANNEL INHIBITION BY PKC
Fig. 5. Immunoblot analysis of total (T) and biotin-recovered membrane (M) fractions of Xenopus oocytes, probed with an anti-Kv1.1
antibody. A: each lane represents extract from oocytes from same frog
injected with same volume of same mRNA preparation and allowed to
express channels for 2 days. Oocytes were treated for 2 min with
either vehicle (0.01% DMSO), 500 nM 4␣-phorbol (4␣), or 500 nM
PMA and then washed for 30 min before being labeled with NHS-SSbiotin. T and M extracts were prepared as described in MATERIALS AND
METHODS. T lanes represent 0.5 oocytes, and M lanes represent 50
oocytes. Positive control is 6 µg of crude mouse brain (br) extract and
shows a single band migrating at ⬃83 kDa. Extracts from uninjected
oocytes show no signal (data not shown on this gel). The Kv1.1
protein in the T fraction runs ⬃58–63 kDa (at lower intensity this is
resolved into two bands), and Kv1.1 protein in M fraction shows 2
bands at ⬃59 and ⬃64 kDa; these are approximations because
Rainbow molecular mass markers (MW) are not linearly related to
migration distance (see manufacturer’s literature). B: densitometric
data showing fraction of total signal that is represented by membranebound channel protein (M/T) are compared with amplitudes of K
channel currents in drug-treated oocytes relative to control (untreated) oocytes from same experiment. For analysis, densities of the
2 bands for each M lane were combined and divided by density of
corresponding T lane (1 or 2 bands, depending on intensity). Data
points represent means ⫾ SE for 4 experiments. Electrophysiological
measurements are means ⫾ SE for 4 experiments with 7 oocytes
each, except for the current amplitude for DMSO, which is average of
2 experiments with 7 oocytes each. Immunoblot images were visualized by autofluorography using enhanced chemiluminescence and
analyzed with a Bio-Rad GS-700 scanning densitometer and Molecular Analyst software.
channel protein that correlate with different magnitudes of K channel current, we compared the Kv1.1
protein signals with the K current amplitudes at 0.5–3
days postinjection. Importantly, as the current amplitude increased with time of expression, so did the
density of the protein signal on the Western blot (data
not shown). However, in our experiments, the Kv1.1
current decreases by 90% on activation of PKC, without
a significant change in the membrane protein level. In
addition, large inhibition of Kv1.1 current by PMA was
not accompanied by a change in the capacitative tran-
C107
sient induced in the oocyte membrane by the voltage
step (data not shown).
Botulinum toxin sensitivity. The specific components
of the regulatory pathway of the PKC-mediated inhibition of Kv1.1 K channels are not known. Mechanisms
ascribed to other channel regulatory pathways, such as
phosphorylation of the channel protein and either
modulation of the voltage sensor or translocation from
the membrane to the cytosol, seem unlikely to explain
this large inhibition of K current by PKC activation. In
a search for alternative mechanisms, we discovered
that the C3 exoenzyme of botulinum toxin blocked the
inhibition of Kv1.1 by PKC activation. The known
targets of C3 toxin are low-molecular mass (20–25 kDa)
G proteins, specifically RhoA and RhoC, which are
selectively modified by ADP-ribosyltransferase activity,
preventing their interaction with effector proteins (6,
27, 31).
Two days after injection with Kv1.1 channel RNA, we
injected oocytes with C3 exoenzyme and then incubated
the oocytes at different temperatures for 1–2 h. Figure
6 shows the effect of PMA application on C3-injected
oocytes. C3 inhibited PKC-mediated K channel inhibition in a temperature-sensitive fashion. The temperature dependence is, perhaps, not surprising if C3
exoenzyme mediates ADP-ribosylation of a substrate
required for K current inhibition. In control experiments, the C3 exoenzyme was heated to ⬎110°C for 2 h,
allowed to cool, and then injected into oocytes. This
heat-inactivated preparation did not inhibit PKCmediated inhibition of K channel current (Fig. 6, A and
B). Alone, none of these treatments altered the K
channel current kinetics, gating, or amplitude, compared with injection of heat-inactivated C3 toxin or no
injections.
DISCUSSION
A membrane-permeant activator of PKC inhibits
cloned Shaker subfamily K channel currents expressed
in Xenopus oocytes. The inhibition of K current is
specific for a phorbol ester known to activate PKC, and
the effects of PMA application are prevented by two
inhibitors of PKC. Agonist stimulation of coexpressed
purinergic receptors also inhibits the K channel currents, mimicking the effect of PKC activation. Together,
these data strongly suggest that direct activation of
PKC through membrane-permeant activators, or indirect activation through cell-surface receptors coupled
to phospholipase C, inhibits K channel currents of the
Shaker subfamily.
Cell surface receptors that couple to PLC may also
inhibit K channel currents in native cells. PKC is a
ubiquitous enzyme, present in high concentrations in
brain and other tissues (16), and many neurochemicals
activate phosphatidylinositol 4,5-bisphosphate hydrolysis, including ATP, histamine, thrombin, and bradykinin. The regulation of neuronal and glial K channels by
transmitter- or hormone-activated PKC may be an
important physiological mechanism for the modulation
of membrane excitability and K homeostasis. Likewise,
there may be important roles for PKC regulation of
C108
K CHANNEL INHIBITION BY PKC
Fig. 6. Effect of botulinum toxin C3 exoenzyme on PKC-mediated
inhibition of K channel currents. C3 exoenzyme was injected into
mKv1.1 K channel-expressing oocytes, and oocytes were then incubated for 1–2 h at 16, 22, or 30°C before electrophysiological
recordings, which were performed at 22°C. A: representative cells
(n ⫽ 3) for each incubation temperature. Controls included cells
injected with heat-inactivated C3 exoenzyme (HI-C3; see MATERIALS
AND METHODS for details) and then incubated at 30°C before testing
(dashed line). In all cases, PMA (500 nM) was applied for 1 min in
bath perfusion and peak currents were measured every 5 s with
pulses to 0 mV from ⫺80 mV. Occasional interruptions to this
protocol were made to run other current-voltage protocols. B: comparison of C3 treatment on K inhibition by PMA, measured 15 min after
PMA application. Bars, means ⫾ SE for 3 cells each.
nonneuronal K currents by endogenous hormones and
messenger molecules (see, e.g., Ref. 29).
The molecular mechanisms of PKC regulation of K
channels are not completely understood. In some cases,
phosphorylation of channel proteins themselves may be
the structural basis for PKC-mediated events. Consensus protein phosphorylation sites are prevalent in the
deduced amino acid sequences of cloned ion channel
proteins. Commonly located in regions thought to be
intracellular or near the cytoplasmic face are sequences
targeted by PKC, cAMP-dependent protein kinase, and
other posttranslational modification sites. Busch et al.
(3) showed that a specific Ser-to-Ala mutation in a
cloned voltage-dependent K channel subunit (ISK ) from
rat kidney prevented K current inhibition by PKC
activators. Also, Cai and Douglass (4) reported biochemical evidence that PKC phosphorylates a voltage-gated
K channel (hKv1.3) in T lymphocytes. And Covarrubias
et al. (8) suggested that inactivation of a Shaw subfamily K channel (Kv3.4) is regulated by direct phosphorylation of the channel protein. In the present study,
however, site-directed substitution of the consensus
PKC phosphorylation sites on mKv1.1 channels did not
prevent the inhibition of K current by PMA. A similar
result was reported for the inhibition by PKC activation of rKv4.2 K currents (21). Although we cannot rule
out the possibility of phosphorylation at novel or dormant sites in the folded channel protein, at least two
consensus PKC sites in Shaker subfamily K channels
are not likely to be directly phosphorylated by PKC, on
the basis of mutagenesis data. This suggests that the
substrate for phosphorylation may be a protein that
associates with or indirectly regulates the Kv1.1 channel protein.
We found that the C3 exoenzyme of botulinum toxin
blocks the inhibitory effect of PKC activation on Shaker
subfamily K currents. The C3 exoenzyme of botulinum
toxin ADP-ribosylates small-molecular weight G proteins of the ras superfamily. A major target of C3
exoenzyme is thought to be RhoC (27), which has a
known role in the regulation of actin microfilament
assembly. For example, C3 exoenzyme causes morphological changes in mammalian cells due to disassembly
of actin microfilaments (6). Thus one hypothesis is that
small GTP-binding proteins, downstream from PKC
activation, mediate the inhibition of Shaker subfamily
K channel currents through interactions with actin
microfilaments.
In most cases, protein kinases are thought to be
‘‘modulators’’ or modifiers of ion channel gating. Perozo
and Bezanilla (24) found that PKC activation shifts the
voltage dependence of activation gating of a squid
giant-axon delayed rectifier K channel. They suggested
that phosphorylation of the channel protein modulates
the voltage-sensing capabilities of the channel through
an electrostatic interaction of added phosphate groups
with the voltage sensor. Also for squid axon, Augustine
and Bezanilla (1) found that MgATP modulates K
conductance through an endogenous, unidentified protein kinase. Similarly, PKC activation caused a rightward shift in the voltage dependence of activation
gating of a rat kidney K channel, and mutagenesis
experiments were used to identify the serine substrate
for the phosphorylation event (3). In addition, phosphorylation of Kv2.1 was reported to cause a depolarizing
shift in activation gating (20). In all of these studies,
the inhibitory effects of PKC may be explained by a
phosphorylation-mediated positive shift in the voltage
dependence of K channel gating. The addition of phosphate groups to the K channel protein is proposed to
increase the negative surface charge on the cytoplasmic
K CHANNEL INHIBITION BY PKC
side of the membrane, thus changing the local electric
field, rendering the channels less sensitive to activating
stimuli.
However, gating changes are not the only mechanism
by which PKC activation alters channel function. In
our experiments and those of others on Kv4.2 and
Kv4.3 K channels (21), there was no evidence for a
PKC-induced change in K channel gating. Moran et al.
(18), however, noted a small change in the voltage
dependence of activation and inactivation gating of
ShakerH4 channels. Together with our results that the
consensus PKC phosphorylation sites are not required
for the PKC-induced inhibition of mKv1.1 K channels,
our data suggest that PKC inhibits K channel function.
This could occur by reducing the number of functional
channels so that after PKC inhibition of current, the
remaining channels are normal, or by reducing the
single-channel conductance of each channel.
One mechanism for a change in the number of
functional channels is a selective endocytotic mechanism, as has been reported for hormone-activated
endocytosis of K channels in starfish oocyte plasma
membranes (17). Such a process has also been implicated in the receptor and second messenger regulation
of the plasma membrane stability of adrenergic receptors (12, 35), Na⫹ channels (9), and Na⫹-K⫹-ATPase
(34). Interestingly, the time course of agonist-induced
␤-adrenergic receptor or activity-induced Na⫹ channel
internalization parallels the time course of the inhibition of K channel current in our studies. However, we
assayed for changes in the density of Kv1.1 channel
protein present at the cell surface before and after PKC
stimulation in oocytes. Although the K channel current
is 90% inhibited by PKC activation, there is no concomitant reduction in the amount of Kv1.1 protein detected
in membrane extracts, indicating that PKC stimulation
does not cause internalization of the K channel protein.
This is consistent with the lack of observable change in
the oocyte membrane capacitance after PKC activation.
We conclude therefore, that PKC activation causes K
channels present at the cell surface to become nonfunctional or significantly reduces the average singlechannel conductance.
Interestingly, the regulation of K channel function by
PKC is clearly different for the different molecular
species of K channel. The large, nonmodulatory, PKCmediated inhibition of K channel activity exemplified
by mKv1.1 is also seen for other Shaker subfamily K
channels. However, K channels of the Shab, Shaw, and
Shal subfamilies are only slightly or not at all inhibited
by PKC activation, and, in some cases, inactivation
gating is also modulated by PKC. Covarrubias et al. (8)
first noted the elimination by PKC of N-type inactivation in a human Kv3.4 channel and have evidence for
PKC-mediated phosphorylation of four serines within
the inactivation ‘‘gate’’ (2). Likewise, we observed PKCmediated increases in current amplitude and slowing of
inactivation kinetics in rKv3.4, but not in all channels
demonstrating fast inactivation (e.g., not Sh⌬ or rKv4.2).
The different effects of PKC activation on different K
C109
channels are likely explained by differences in channel
structure, possibly at PKC phosphorylation sites, or
functional regions that may interact with other membrane-associated or cytoplasmic proteins.
Understanding the physiological importance of PKC
regulation of K channels depends on knowledge about
the molecular species of K channel subunits that
account for the native K currents. In addition, the
diversity of PKC and phospholipase isoenzymes suggests that there may be cellular or regional specificity
to receptor-activated PKC coupling in native cells. The
coupling between neurotransmitter or hormone receptors and voltage-dependent K channels could have
important effects on the roles of K channels in determining the resting membrane potential, action potential
duration, speed of repolarization, and regulation of K
homeostasis and synaptic plasticity. Regulation of K
channel availability would thereby alter neurochemical
communication and other important functions served
by K channels.
We thank Heather Hill for help with early experiments, Viet Hoa
Hoang for help with oocyte preparations and Western blots, Gary
Yellen for generously donating the data acquisition software, and T.
Kendall Harden for cDNAs for purinergic receptors. We are grateful
to Manuel Covarrubias, Rolf Joho, Olaf Pongs, Larry Salkoff, and
Bruce Tempel for K channel cDNAs. We thank James Trimmer for a
gift of anti-Kv1.1 and discussions on protein extracts and Scott
O’Grady for helpful discussions. We thank Wildcat Farms for beef
liver for feeding frogs.
This study was supported by grants from The Minnesota Medical
Foundation and the National Institutes of Health.
Address for reprint requests and other correspondence: L. M.
Boland, Dept. of Physiology, Univ. of Minnesota, 6–255 Millard Hall,
Minneapolis, MN 55455 (E-mail: [email protected]).
Received 4 November 1998; accepted in final form 7 April 1999.
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