From the Department of Pathology, Anatomy and Cell Biology, Jefferson Medical College, Philadelphia, Pennsylvania 19107
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ABSTRACT |
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Protein kinase C inhibits inactivation gating of Kv3.4 K+ channels, and at least two NH2-terminal
serines (S15 and S21) appeared involved in this interaction (Covarrubias et al. 1994. Neuron. 13:1403-1412). Here
we have investigated the molecular mechanism of this regulatory process. Site-directed mutagenesis (serine
alanine) revealed two additional sites at S8 and S9. The mutation S9A inhibited the action of PKC by ~85%, whereas
S8A, S15A, and S21A exhibited smaller reductions (41, 35, and 50%, respectively). In spite of the relatively large
effects of individual S
A mutations, simultaneous mutation of the four sites was necessary to completely abolish
inhibition of inactivation by PKC. Accordingly, a peptide corresponding to the inactivation domain of Kv3.4 was
phosphorylated by specific PKC isoforms, but the mutant peptide (S[8,9,15,21]A) was not. Substitutions of negatively charged aspartate (D) for serine at positions 8, 9, 15, and 21 closely mimicked the effect of phosphorylation
on channel inactivation. S
D mutations slowed the rate of inactivation and accelerated the rate of recovery from
inactivation. Thus, the negative charge of the phosphoserines is an important incentive to inhibit inactivation.
Consistent with this interpretation, the effects of S8D and S8E (E = Glu) were very similar, yet S8N (N = Asn) had
little effect on the onset of inactivation but accelerated the recovery from inactivation. Interestingly, the effects of
single S
D mutations were unequal and the effects of combined mutations were greater than expected assuming a simple additive effect of the free energies that the single mutations contribute to impair inactivation. These
observations demonstrate that the inactivation particle of Kv3.4 does not behave as a point charge and suggest
that the NH2-terminal phosphoserines interact in a cooperative manner to disrupt inactivation. Inspection of the
tertiary structure of the inactivation domain of Kv3.4 revealed the topography of the phosphorylation sites and
possible interactions that can explain the action of PKC on inactivation gating.
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INTRODUCTION |
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A-type K+ channels activate, open, and inactivate in response to membrane depolarization. In excitable tissues, inactivation of these channels helps to control the
frequency of repetitive spike firing, to shape the action
potential (Hille, 1992) and to regulate the amplitude
of back-propagating action potentials (Hoffman et al.,
1997
). There are two well-characterized mechanisms of
inactivation of voltage-gated K+ channels: N- and C-type.
The N-type mechanism is determined by an amphipathic region at the NH2 terminus of the channel protein, which acts as an internal blocking particle that occludes the inner mouth of the pore (Hoshi et al., 1990
;
Demo and Yellen, 1991
; Ruppersberg et al., 1991
; Murrell-Lagnado and Aldrich, 1993a
, 1993b
; Baukrowitz
and Yellen, 1995
; Gomez-Lagunas and Armstrong, 1995
).
The C-type mechanism is determined by residues that
contribute to the K+-selective pore and is thought to involve cooperative intersubunit interactions that alter
the structure of the outer mouth of the channel (Hoshi
et al., 1991
; Lopez-Barneo et al., 1993
; Baukrowitz and
Yellen, 1995
; Ogielska et al., 1995
; Panyi et al., 1995
;
Starkus et al., 1997
).
Inactivation of voltage-gated K+ channels may be
modulated by various physiological factors (e.g., protein kinases, auxiliary subunits, redox potential, and
external K+). Phosphorylation of these channels by
protein kinases is of importance because this post-translational modification appears to be associated with
physiological processes that underlie synaptic plasticity
(Levitan, 1994; Kandel et al., 1995
; Jonas and Kaczmarek, 1996
). Several reports have shown that protein
kinases regulate N-type inactivation of A-type K+ channels by phosphorylating specific amino acids (Covarrubias et al., 1994
; Drain et al., 1994
; Roeper et al., 1997
).
However, the molecular mechanisms of action are not
yet known. PKA-dependent phosphorylation appears to
be necessary for rapid N-type inactivation of Drosophila
ShakerD K+ channels (Drain et al., 1994
). Two serines
(S507 and S508) located at the cytoplasmic COOH-terminal domain of ShakerD are involved in this process.
Thus, it was proposed that PKA indirectly modulates N-type inactivation by phosphorylation of regulatory
sites at the COOH terminus of ShakerD. CaMK II
(Ca++/calmodulin-dependent protein kinase) slows inactivation of Kv1.4 by phosphorylation of S123 (Roeper
et al., 1997
). This residue is located downstream of the
inactivation domain at the NH2 terminus. Roeper et al.
(1997)
postulated that phosphorylation of S123 may slow inactivation either by reducing the flexibility of
the "chain" that links the NH2-terminal inactivation
particle with the core of the channel, or by interfering
with the binding of the inactivation particle to its receptor at the inner mouth of the channel. Another study
reported that C-type inactivation of Kv1.3 is modulated at three putative phosphorylation sites (Kupper et al.,
1995
) but the mechanism of action was not examined.
Other aspects of voltage-dependent gating and expression of K+ channels are modulated by protein kinases
(Perozo and Bezanilla, 1991
; Jonas and Kaczmareck,
1996). There also, little is known about the underlying
molecular mechanisms.
We have previously shown that PKC eliminates N-type
inactivation of an A-type K+ channel encoded by Kv3.4
(Covarrubias et al., 1994). This study demonstrated that
mutation of two serines (S15A, S21A and S[15,21]A) partially inhibited the effect of PKC on N-type inactivation. However, these mutations were not capable of explaining the totality of the PKC effect. In the present
study, we demonstrate that there are two additional
sites (S8 and S9) and that the four serines (S8, S9, S15,
and S21) are sufficient to account for the PKC-mediated inhibition of inactivation, but each site does not
contribute equally to this action. Another goal of this
study was to gain insights into the mechanism of PKC
action. Because the net positive charge of the inactivation particle of Shaker K+ channels is known to be critical to achieve rapid inactivation (Murrell-Lagnado and
Aldrich, 1993a
), the NH2-terminal phosphoserines could simply neutralize the overall positive charge and
consequently slow inactivation. In agreement with this
possibility, our previous study (Covarrubias et al., 1994
)
showed that substitution of aspartate (D) for serine at
position 15 produced a partial inhibition of inactivation. Here, we have analyzed all possible S
D substitutions at positions 8, 9, 15, and 21 to address two questions. (a) Are all phosphorylation sites functionally equivalent? By mutating one residue at a time or in various
combinations, we have tested whether the net charge of
the inactivation particle is the sole determinant of the
rate of inactivation. (b) Do the phosphorylation sites act
independently? Because of the close proximity of the phosphorylation sites, it is conceivable that interactions
between these sites are involved in the mechanism of
PKC action. By examining the free energy that single
S
D mutations contribute to disrupt inactivation, we
have determined whether the effects of combined S
D
mutations are the result of simple additivity. Finally, aided by the nuclear magnetic resonance-based structure of the inactivation particle of Kv3.4 (Antz et al.,
1997
), we developed a plausible working hypothesis that
can explain the structural changes that the inactivation
gate may undergo upon phosphorylation by PKC. This
hypothesis represents one of the first opportunities to
relate the actual three-dimensional structure of a voltage-gated K+ channel to a particular function and its
regulation. Preliminary results were previously reported
in abstract form (Covarrubias et al., 1997
).
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MATERIALS AND METHODS |
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Mutagenesis
A cDNA encoding Kv3.4 was maintained as described previously
(Covarrubias et al., 1994). S
A substitutions (except S[8,9,15]A and S[8,9,21]A) were created as described before using the Altered Sites II in vitro Mutagenesis System (Promega Corp., Madison, WI) or an overlap polymerase chain reaction strategy (Covarrubias et al., 1994
). All other amino acid substitutions were
made using the QuickChange Site-Directed Mutagenesis Kit as
described by the manufacturer (Stratagene Inc., La Jolla, CA).
Briefly, complementary pairs of oligonucleotides (22mer-29mer)
containing the appropriate nucleotide substitutions were prepared at the Nucleic Acid Facility, Kimmel Cancer Center (Thomas
Jefferson University, Philadelphia, PA). These oligonucleotides
were subsequently used as primers for the complete synthesis of
both strands of the Kv3.4 plasmid. We used 20 ng of Kv3.4 plasmid as template, 5 U Pfu DNA polymerase (Stratagene Inc.),
primers, and free nucleotides in a total volume of 100 µl. After
strand synthesis (12-18 cycles), 10 U of DpnI were added to the
reaction mixture to digest the original Kv3.4 methylated plasmid
template (37°C, 1-2 h). The restriction endonuclease was heat
inactivated (65°C, 15 min), and the mixture used to transform DH5
cells by electroporation. Base substitutions were confirmed by automated DNA sequencing at the Nucleic Acid Facility, Kimmel Cancer Center. It should be noted that QuickChange
does not involve a polymerase chain reaction. Pfu DNA polymerase (containing 3'
5' exonuclease activity or proofreading
activity) simply catalyzes the extension step of the mutagenesis reaction replicating the template with a mutagenic primer. Nevertheless, to confirm that base misincorporations were unlikely under our reaction conditions, we read 105 DNA sequences (~500
bp, each) created by QuickChange. Approximately 45 of these sequences correspond to the region that surrounds the S4-S5 loop
of three distinct K+ channels (Kv3.4, Kv4.1, and dShaw); the rest
correspond to the NH2-terminal region of Kv3.4 (58) or the region surrounding the S6 region of Kv4.1 (2). This analysis did
not reveal nucleotide errors introduced by Pfu DNA polymerase
activity. In addition, the mutants characterized here did not exhibit any unexpected properties, and subcloning of some mutated sequences (S8D and S[8,15,21]D) back into the wild-type
cDNA did not result in different phenotypes. Various studies
have investigated the fidelity of this enzyme and other thermostable DNA polymerases (Kunkel, 1988
; Lundberg, et al., 1991;
Flaman et al., 1994
; Cline et al., 1996
). They found that Pfu DNA
polymerase yields the highest fidelity with an error rate {[mutation frequency]/([base pairs] [effective duplication])} on the order of 1-2 × 10
6. This is at least 10× better than Taq polymerase. By applying this formula, we predicted a mutation frequency of ~2%. Assuming that all sequences are equally
vulnerable to errors and that each analyzed sequence is an independent trial, we expected at least two sequences containing one
undesired mutation. Thus, it appears that under our assay conditions, which do not involve PCR, the mutation frequency is overestimated. cRNA for microinjection was produced as described
elsewhere (Jerng and Covarrubias, 1997
).
Microinjection of Xenopus Oocytes and Electrophysiological Recording
Wild-type and mutant Kv3.4 cRNA was microinjected into defolliculated Xenopus oocytes (~50 ng/cell) using a Nanoject microinjector (Drummond, Broomall, PA). Whole-oocyte currents were
recorded 2-10 d after injection using the two-microelectrode
voltage-clamp technique (TEV-200; Dagan Corp., Minneapolis,
MN). Microelectrodes were filled with 3 M KCl (tip resistance was
<1 M). Bath solution contained: 96 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, pH 7.4, adjusted with
NaOH. Phorbol 12-myristate-13-acetate (PMA)1 was purchased
from Sigma Chemical Co. (St. Louis, MO). Current traces were
digitized at 250-500 µs/point after low-pass filtering at 1-2 kHz.
The average voltage offset recorded at the end of an experiment
was generally small (0.5 ± 2.4 mV, n = 107) and was subtracted
from the command voltage when analyzing prepulse inactivation
curves. The leak current was subtracted off-line assuming ohmic
leak. Capacitive currents were subtracted on-line using a P/4 protocol or off-line using a scaled noise-free template generated
from a current trace with no active time-dependent currents (elicited by a depolarization to
80 mV). Experiments were conducted at 23°C using a temperature-controlled microscope stage
(PDMI-2; Medical Systems Corp., Greenvale, NY).
Data Acquisition and Analysis
Voltage-clamp protocols and the acquisition of data were controlled by a 486 desktop computer interfaced to a 12-bit A/D
converter (Digidata 1200 using pClamp 6.0; Axon Instruments,
Foster City, CA). Data analysis was conducted using Clampfit
(pClamp 6, Axon Instruments), Sigmaplot (Jandel Scientific, San
Rafael, CA), or Origin 4.1 (Microcal, Northampton, MA). The
slow inactivation that remained after PMA application exhibited
complex and variable kinetics (Covarrubias et al., 1994; see Fig. 1
A). This is probably the result of variable degrees of phosphorylation of multiple serines in four subunits of the channel tetramer.
Therefore, to simplify the comparative analysis of wild-type and
S
A mutant channels, the normalized current integral was
computed (current integral/peak current) before and after
PMA. In all cases, this analysis included currents elicited by 900-ms
step depolarizations to +50 mV. To analyze inactivation of the S
D mutants, the decaying relaxation of the currents was described
assuming a sum of two exponential terms as described previously
(Jerng and Covarrubias, 1997
). Because the S
D mutations
studied here mainly affected the time constant of the fast component and the relative weight of this component correlated with
changes in the steady state level of the current (see RESULTS), we
used the following relation to estimate the rate of inactivation
(ki): ki = (1
Is)/
i, where Is is the sum of the relative weights of
the slow and steady state components of the current and
i is the
time constant of fast inactivation (Murrell-Lagnado and Aldrich,
1993a
; Panyi et al., 1995
). According to the theory of absolute reaction rates (Gutfreund, 1995
), the following relation was used to
estimate the free energy change that a particular mutation contributes to inhibition of inactivation:
Gobs =
RT ln(ki h/kB T),
where R, T, h, and kB have their usual meaning (Gutfreund,
1995
). Then, relative to wild type, the free energy that a single
mutation contributes to inhibition of inactivation is:
Gobs =
Gobs(wild type)
Gobs(mutant).
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If the effect of multiple mutations results from the additive effect of single mutations, the predicted free energy change that a
multiple mutation contributes to inhibition of inactivation is:
G(multiple mutation) =
G(single mutation 1) +
G (single mutation 2) + ...; therefore, assuming additivity, the predicted inactivation rate (kpred) associated with a multiple mutation is kpred = (kBT/h) exp(
Gpred/RT), where
Gpred is the predicted free energy change [
Gpred =
Gobs(wild type) +
G(multiple mutation)]. A deviation from this prediction suggests that the mutations do not act independently. A similar procedure was used to examine the free energy that mutations contribute to destabilization of the inactivated state of the channel.
In this case, we assumed that the observed rate of recovery from
inactivation (kr) = 1/(fast time constant of recovery from inactivation). Although the time course of recovery from inactivation
of wild-type and some mutant channels was best described assuming the sum of two exponential terms (the fast component contributed to >65% of the time course; see RESULTS), a simple exponential rise was sufficient for several mutants that exhibited
faster recoveries from inactivation. For K+ channels that exhibit
N-type inactivation, the fast phase of the recovery from inactivation is associated with the exit of the inactivation particle from
the pore (Demo and Yellen, 1991
).
Synthetic Peptides and Phosphorylation In Vitro
The wild-type peptide corresponding to the first 28 amino acids
of Kv3.4 was provided by Dr. R.W. Aldrich (Stanford University, Stanford, CA) and handled as described before (Covarrubias et al., 1994). The mutant peptide S[8,9,15,21]A was purchased
from Multiple Peptide Systems (San Diego, CA). All peptides
were purified (>95%) by reverse phase chromatography. The
peptides were applied to a C8 column attached to the FPLC
(Pharmacia LKB Biotechnology Inc., Piscataway, NJ) and the adsorbed material was eluted with a linear acetonitrile gradient
(0-30%) in 0.1% trifluoroacetic acid. Recombinant rat PKC isoforms (
,
, and
) were provided by Dr. C.D. Stubbs (Jefferson
Medical College). Phosphorylation in vitro was carried out using
a filter assay as described previously (Slater et al., 1993
). To evaluate whether a peptide can act as a substrate of PKC, this assay was
designed to measure the initial rates of phosphate incorporation.
The functionality of the isozymes was always confirmed using the
myelin-basic-protein as the control substrate. Phosphorylation
data are expressed as molar fraction of phosphorylated peptide
in the presence of diacylglycerol.
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RESULTS |
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Inhibition of Inactivation by PKC Is Associated with Phosphorylation of Four NH2-Terminal Serines
The first 28 amino acids at the NH2 terminus of the
Kv3.4 protein constitute its inactivation gate (Ruppersberg et al., 1991; Rettig et al., 1992
; Covarrubias et al.,
1994
). This sequence includes four putative PKC phosphorylation sites at positions 8, 9, 15, and 21 (underlined; MISSVCVSSYRGRKSGNKPPSKTCLKEE). Previously, we showed that S15 and S21 are important contributors to the action of PKC on N-type inactivation,
but are not sufficient to explain the totality of the inhibition of inactivation by PKC (Covarrubias et al., 1994
).
To identify the remaining residues and further investigate their contribution, we examined a series of new
S
A substitutions affecting of S8, S9, S15, and S21
(see also subsequent section). Oocytes expressing the
wild-type and mutant currents were exposed to a phorbol ester (PMA, 20 nM) to activate PKC. In sharp contrast to the wild-type currents that exhibit a dramatic inhibition of inactivation induced by PKC activation, currents
expressed by the quadruple mutant (S[8,9,15,21]A) were not changed in the presence of PMA (Fig. 1, A and C).
Thus, S8, S9, S15, and S21 are the most likely phosphorylation sites associated with inhibition of inactivation by PKC.
To verify this result, we conducted in vitro phosphorylation assays using peptides that correspond to the first 28 amino acids of Kv3.4 (the NH2-terminal inactivation domain of the channel) as substrates. We found that three
PKC isoforms (,
, and
) phosphorylated the Kv3.4
peptide but, as expected, did not significantly phosphorylate the quadruple mutant peptide S[8,9,15,21]A (Fig. 2). The small signal observed with S[8,9,15,21]A corresponds to unspecific phosphate incorporation observed with unrelated peptides that have no phosphate
acceptors (e.g., the ShakerB inactivation domain; data
not shown). These results confirmed that the NH2-terminal inactivation domain of Kv3.4 is phosphorylated
by PKC and that this modification is associated with
elimination of N-type inactivation by PKC in the intact
channel.
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Evidence of Nonequivalent and Nonindependent Phosphorylation Sites at the NH2 Terminus of Kv3.4
In the previous section, we showed that PKC-mediated
inhibition of inactivation involves multiple phosphorylation sites. Thus, to understand the mechanism that disrupts inactivation, it is necessary to know: (a) whether
the phosphorylation sites are functionally equivalent,
and (b) whether the phosphorylation sites act independently. To investigate these questions, we examined the
effect of PKC activation on single, double, and triple
S A mutants. The current integral (see MATERIALS AND
METHODS) was computed before and after application
of 20 nM PMA. In the absence of PMA, the current integrals exhibited little or no difference between the
wild-type and mutant channels (Fig. 1 D, top). Note that
the time courses of the currents were similar (e.g., Fig.
1, A-C). By contrast, in the presence of PMA, the current integrals of the mutants differed significantly from
that of the wild type (Fig. 1 D, bottom). S9A and S[8,9]A exhibited a greater reduction in their responses to
PMA (13 and 10% of the control response, respectively) than S8A, S15A, S21A, and S[15,21]A (59, 64, 50, and 39% of the control response, respectively). Thus, the presence of S9 appears to be more critical
than that of S8, S15, and S21. Moreover, the results in
Fig. 1 D also showed that the sites do not act independently. Addition of the remaining current integrals of
the mutants in the presence of PMA did not account
for the results observed with wild-type channels. For instance, the sum of the remaining current integrals
from S[8,9]A and S[15,21]A in the presence of PMA
was only ~50% of the normalized current integral
from wild type. If the sites were independent, this result
would be unexpected because phosphorylation of S8, S9, S15, and S21 accounts for the effect of PMA on current inactivation (Fig. 1, A and C). In addition, the apparent contribution of S15 and S21 to the total effect of
PMA on inactivation is considerable (~50%) when
these residues were mutated to alanine (individually or
in combination; Fig. 1 D). Yet only ~10% of the response to PMA remained when both S15 and S21 are
available (see S[8,9]A). Even lower responses were observed when S15 or S21 were the only available sites (see
S[8,9,21]A and S[8,9,15])A, respectively). Although the
pooled data (from different batches of oocytes) did not reveal a statistically significant difference between
S[8,9,15,21]A and S[8,9,21]A (P
0.5), this mutant
channel exhibited a small but consistent disruption of
inactivation in the presence of PMA (Fig. 1 B). Therefore, the sole contribution of S15 and S21 to the effect
of PKC on inactivation is small (when the other sites are not available). Later experiments will confirm this
observation more directly. Overall, these results suggest
that the ability of a phosphorylated site (or set of sites)
to disrupt inactivation depends on whether other sites
are phosphorylated. Thus, interactions between NH2-terminal phosphoserines play a critical role in determining the disruption of inactivation by PKC.
Serine to Aspartate Mutations at Positions 8, 9, 15, and 21 Mimic the Effect of PKC on K+ Channel Inactivation
To further investigate the interactions described above
and how they may disrupt inactivation of Kv3.4, we
studied the biophysical properties of serine aspartate (S
D) mutants. Like phosphorylation, the S
D
mutations incorporate negatively charged side chains. Although there is no sequence similarity between the
NH2-terminal regions of ShakerB and Kv3.4, both share a
net positive charge within the first 25 amino acids (+2
and +5, respectively). Furthermore, positive charges
are clearly clustered in the sequence. Previous studies
with ShakerB K+ channels have demonstrated that the
net positive charge of the inactivation domain determines long-range electrostatic interactions that facilitate the diffusion of the inactivation domain to the inner mouth of the pore (Murrell-Lagnado and Aldrich,
1993a
). If the inactivation domain in Kv3.4 behaves as a
point charge, phosphorylation could impair inactivation by simply reducing or reversing the net positive
charge of the inactivation domain (from +5 to
3, if
all sites are phosphorylated). To test this hypothesis, we
created S
D mutations affecting positions 8, 9, 15, and 21 individually and in all possible combinations.
We found that the currents mediated by these mutant
channels (in the absence of PMA) mimic the action of
PKC on K+ channel inactivation (Figs. 3 and 4). For example, wild-type currents recorded in the presence of
PMA resemble the currents expressed by S[8,9,15,21]D.
Similarly, S[8,9]A currents in the presence of PMA
(S15 and S21 are available for phosphorylation) closely
resemble currents expressed by S[15,21]D. The similarity was not as striking when comparing S[15,21]A currents in the presence of PMA (S8 and S9 are available
for phosphorylation) and currents expressed by S[8,9]D.
This was mainly due to the larger sustained current of
S[8,9]D. Such a difference may have occurred because: (a) aspartates at 8 and 9 may destabilize the inactivated
state to a greater extent than phosphoserines at the
same positions; (b) S[15,21]A per se reduces the ability
of phosphoserines at 8 and 9 to disrupt inactivation;
and (c) with S[8,9]D, all channels are modified, whereas
phosphorylation of S[15,21]A may be limited by the degree of PKC activation by PMA. In spite of this finding, S[15,21]A currents in the presence of PMA exhibited
intermediate kinetics between that of wild-type and
S[8,9]A channels recorded in the presence of PMA,
and S[8,9]D currents exhibited intermediate kinetics
between that of S[8,9,15,21]D and S[15,21]D currents (Fig. 3). These observations demonstrated that S
D
mutations reproduce a phenotype that is reasonably
equivalent to that of constitutively phosphorylated
channels and that the presence of negative charges at
positions 8, 9, 15, and 21 may be important to inhibit
N-type inactivation.
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Serine to Aspartate Mutations at Positions 8, 9, 15, and 21 Slow Inactivation and Destabilize the Inactivated State in a Nonadditive Manner
Single S D mutants exhibited different degrees of impaired inactivation and, as shown above, a quadruple
mutation (S[8,9,15,21]D) was necessary to closely mimic
the complete effect of PKC (Fig. 4 A). To examine the
kinetics of inactivation of the S
D mutants, the decaying relaxation of the currents was described assuming the sum of two exponential terms. Analysis of the
wild-type currents gave the following best-fit parameters at +50 mV:
f = 7.5 ± 1.4 ms,
s = 35 ± 9.6 ms,
Wf = 0.82 ± 0.05, Ws = 0.14 ± 0.04, and Wss = 0.03 ± 0.02 (mean ± SD, n = 32;
f and
s are the fast and slow
time constants, respectively; Wf and Ws are the corresponding relative weights of the fast and slow components of current decay, respectively; and Wss is the relative weight of the steady state level of the current). Notice that the fast component dominates the kinetics of
current decay (>80%) and that inactivation is nearly
complete (>95%). S
D mutations slowed both
f and
s. However, while Wf was greatly reduced (from 0.84 to
0.02 at the extremes between wild type and S[8,9,21]D),
Ws stayed relatively constant (the largest changes were
0.15 and 0.1 above and below the wild-type value, respectively). Parallel to a decrease in Wf, there was an increase in Wss (0.03 and 0.97 at the extremes between
wild type and S[8,9,21]D). This result suggested that the inactivated state of the channel has been destabilized by the mutations (see below). We focused the
analysis on the fast component of current decay because it was more sensitive to the mutations, and
changes that affect the relative weight of this component correlated with changes in the steady state level of the current. First, we estimated the rate constants of inactivation from the decay of the macroscopic currents
(MATERIALS AND METHODS; Table I). These values were
sorted by size and plotted as a Tukey box plot (Tukey,
1977
). Then, to test whether the simple additive effect
of the single mutants can account for the effects observed with multiple mutants, we calculated the free energies that the mutations contribute to impair inactivation, and plotted the predicted and observed rate constants (MATERIALS AND METHODS; Fig. 4 B; Table I).
Clearly, the extent of inhibition of inactivation in the
single mutants is not equivalent (S8D > S9D > S15D > S21D) and the predicted rates differed significantly
from the observed ones. Thus, as suggested earlier,
NH2-terminal phosphoserines that regulate N-type inactivation in Kv3.4 K+ channels are not functionally
equivalent and interact favorably to eliminate N-type
inactivation in Kv3.4. For instance, S21D alone caused
no effect on inactivation, but enhanced the effects of
S8D or S9D when combined in the double mutants
S[8,21]D and S[9,21]D. Similarly, inhibition of inactivation by S[8,15]D and S[9,15]D was enhanced by the
presence of S21D in triple mutants S[8,15,21]D and
S[9,15,21]D.
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Single S D mutants also exhibited different degrees of accelerated recovery from inactivation at
100
mV (S8D > S9D
S15D > S21D; Fig. 5 A), and the
multiple mutations caused greater acceleration of recovery from inactivation than expected from the corresponding single mutations (see below). Kinetic analysis
revealed that recovery from inactivation of wild-type
Kv3.4 channels was best described assuming the sum of
two exponential terms. The fast and slow time constants were 538 ± 198 and 3,304 ± 960 ms, respectively,
and their corresponding relative weights were 0.66 ± 0.04 and 0.30 ± 0.04 (mean ± SD, n = 21). S
D mutations decreased both time constants and, in all cases,
the weight of the fast component dominated the time
course of recovery from inactivation (>65%). In fact,
recoveries from inactivation for S[8,9]D, S[8,15]D,
S[8,21]D, S[8,15,21]D, and S[9,15,21]D (those channels that exhibited the fastest recoveries) were well described assuming a simple exponential rise. The estimated rates of recovery from inactivation (MATERIALS AND METHODS) are summarized in Fig. 5 B and Table I.
The recoveries from inactivation of other triple mutants and the quadruple mutant were not examined
quantitatively because they exhibited little inactivation
during a 900-ms pulse to +50 mV. Resembling the onset of macroscopic inactivation (Fig. 4), the effect of
multiple mutations on recovery from inactivation
could not be explained assuming additivity of the free
energies that single mutations contribute to destabilization of the inactivated state (MATERIALS AND METHODS; Table I). The predicted rate constants of recovery
were systematically slower than the observed values
(Fig. 5 B). Overall, however, deviation from additivity
appeared moderate with respect to the results obtained
from the analysis of the onset of macroscopic inactivation (Fig. 4 B).
The results presented above demonstrate that S D
mutations not only reduced the rate of inactivation but
also increased the rate of recovery from inactivation.
Assuming three interconnected states (closed, open,
and inactivated), we expected a depolarizing shift of
the prepulse inactivation curve and an increased level of the noninactivating current. Accordingly, most of
the S
D mutants that exhibited significantly slower
rates of inactivation (Fig. 4) and a destabilized inactivated state (Fig. 5) also showed a rightward shift of the
midpoint of prepulse inactivation and an increased
current level at the foot of the curve (Fig. 6; Table I).
Also, as expected from the analyses of the rates of inactivation and recovery from inactivation, the midpoint
potentials of prepulse inactivation for various double
S
D mutants did not exhibit linear additivity (Fig. 6,
legend). Some mutants that exhibited a small or moderate acceleration of the recovery from inactivation did
not show an increased level of the noninactivating current after a 10-s conditioning prepulse to depolarized
membrane potentials (Fig. 6; Table I). These mutant
channels inactivated almost completely like the wild
type because they slowly entered a second more stable inactivated state (possibly from closed and open states).
The protocol used to determine the rate of recovery
from inactivation did not allow a significant entry into
that inactivated state (Fig. 5, legend).
Electrostatic and Steric Interactions Contribute to the Effects of
NH2-Terminal S D Mutations on Inactivation of Kv3.4
To investigate the relative importance of an electrostatic effect versus steric hindrance, we examined two additional substitutions: S8N (N = Asn), S8E (E = Glu). These mutations allowed us to inquire whether the negative charge (S8E and S8D) and/or the size of the side chain (S8N) altered the inactivation properties of the channel. We chose to study the eighth position because the S8D mutant alone exhibited the largest disruption of inactivation (Figs. 4 and 5). S8E slowed the onset of inactivation to the same extent as S8D, whereas S8N only slightly slowed this process (Fig. 7, A and B). Thus, the slower onset of inactivation of the S8D mutant channels is not likely to be caused by the steric effect of the mutation. As previously suggested, the negative charge appears to be the main factor that explains the slow inactivation of the phosphorylated channels. The presence of steric hindrance was, however, more apparent on the recovery from inactivation (Fig. 7, C and D). S8E increased the rate of recovery from inactivation by approximately fivefold, and S8D and S8N also accelerated the recovery from inactivation but to a more moderate extent (approximately three- and twofold, respectively). This small difference between S8D and S8N indicates that the charge of the side chain plays a minor role on the recovery from inactivation. Thus, by contrast to the onset of inactivation, the stability of the inactivated state is more sensitive to the topography of the inactivation particle, which can be altered by phosphorylation.
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DISCUSSION |
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This study investigated the molecular physiology of the
NH2-terminal phosphorylation sites that regulate inactivation gating of an A-type K+ channel. The main results
show that: (a) PKC acts on four phosphate acceptors
(S8, S9, S15, and S21) within the inactivation domain because mutation of these residues to alanine is necessary and sufficient to remove the action of PKC on
channel inactivation. Moreover, a peptide that corresponds to the inactivation domain of Kv3.4 (the first 28 amino acid) was phosphorylated by three PKC isoforms, but the mutant peptide S[8,9,15,21]A was a
much poorer substrate. (b) Single S A mutants exhibited partially reduced effects of PKC on inactivation.
The remaining effects of PKC were, however, not equal
among the single mutants and linear summation of
these effects could not explain a nearly complete elimination of inactivation caused by PKC. (c) S
D substitutions at the putative phosphorylation sites mimicked
PKC action. The inactivation properties conferred by
single S
D mutations were also not equivalent, and
multiple S
D mutations appeared to interact in a cooperative manner. Thus, a simple reduction or neutralization of the net positive charge of the inactivation domain of Kv3.4 is not sufficient to explain the data. (d)
Analysis of the S
N and S
E mutants revealed that
a combination of electrostatic and steric factors contribute to disruption of inactivation by S
D substitutions or phosphorylation at the NH2-terminal domain.
Structural Basis of PKC Action on Kv3.4 Inactivation
To explain how phosphorylation of a set of four serines
at positions 8, 9, 15, and 21 by PKC causes elimination
of rapid N-type inactivation in Kv3.4, we have considered two possible mechanisms. First, the inactivation
domain could behave as a point charge (Murrell-Lagnado and Aldrich, 1993a). In such a case, the factor that determines the onset of inactivation would be the
net charge of the inactivation domain. Accordingly, the
addition of negatively charged phosphate groups could
simply reduce or neutralize the net positive charge of
the inactivation domain. Then, individual phosphoserines should behave equivalently and act independently. Although aspartates at positions 8, 9, 15, and 21 can mimic PKC action (as predicted by this hypothesis), our data showed that these positions are not functionally equivalent and that multiple mutations interact
(Figs. 1, and 4-6). Also, the net charge change does not
appear to be critical because monoanionic aspartates can closely mimic the action of dianionic phosphates
introduced by phosphorylation. Thus, the inactivation
domain of Kv3.4 does not behave as a simple point
charge.
Second, the topography and electrostatic profile of
the inactivation domain may determine binding to its
receptor in the channel. In this case, the localization of
an active group is more critical. In agreement with this
possibility, the effects of mutations at S8 and S9 were,
in general, more dramatic than those of mutations at
S15 and S21 (Figs. 1-6) and, while the rate of inactivation was mainly sensitive to electrostatic interactions,
the rate of recovery from inactivation was mainly sensitive to steric hindrance (Fig. 7). Additional results also
suggested the presence of other structural changes that
could result from the combination of electrostatic and
steric effects. For instance, S D mutations decreased
the rate of inactivation and increased the rate of recovery from inactivation, but the energetic effects of the
mutations were not linearly additive (Figs. 4-6). Analysis of multiple mutations in other proteins has shown
that deviation from simple free energy additivity may
reflect conformational changes induced by the substitutions or functional coupling between the mutated
residues (Andersen and Koeppe, 1992
). For tyrosyl-tRNA synthetase, crystallographic data have confirmed this interpretation (Carter et al., 1984
; Fothergill and
Fersht, 1991
). Similarly, the contribution of individual
deep pore residues to the conductance and tetraethyl
ammonium block in certain K+ channels is not additive
(Kirsch et al., 1992
). This appears to reflect coupling
between deep pore residues and the importance of the
backbone structure in the pore region. Also, as shown
for certain metabolic enzymes of known structure (glycogen phosphorylase and isocitrate dehydrogenase),
phosphoserines may alter function by a combination of
local and allosteric effects (Johnson and Barford, 1993
;
Johnson and O'Reilly, 1996
).
Inspection of the nuclear magnetic resonance-based
tertiary structure of the inactivation domain of Kv3.4
(Antz et al., 1997) allowed us to visualize the topography and electrostatic profile of critical regions and possible structural changes induced by phosphorylation.
On one face of the molecule, the side chains of S8 and
S15 are located in a region that is partly surrounded by
basic side chains (R11, R13, K14, K18, and K26). The
local electrostatic potential of this region appeared especially important because single negatively charged
substitutions at S8 (S8D and S8E) most effectively
slowed the rate of inactivation, and a neutral substitution (S8N) had little or no effect on the onset of inactivation (Fig. 7). Its topography seemed critical too because, independently of the charge, S8D, S8E, and S8N
destabilized the bound state of the inactivation particle
(i.e., accelerated the recovery from inactivation). On
another face, the side chain of S9 is located in a less polar region, and that of S21 (in close spatial relationship with the side chain of K22) is relatively more distant
from the other three sites. S9D, S15D, and S21D were
significantly less effective in disrupting inactivation
than S8D. The effects of the substitutions on inactivation were, however, greatly enhanced when they appeared in double, triple, or quadruple combinations (even when S8 remained). Energetically, this enhancement was greater than that predicted by a simple additive interaction, suggesting the presence of cooperativity (Figs. 4-6). Thus, a combination of electrostatic and
steric effects (as a result of phosphorylation or negatively charged substitutions) may serve as an incentive that favors a structural change of the inactivation domain. Phosphoserines could simply shift the structural
equilibrium toward a disordered conformation. In support of this model, recent nuclear magnetic resonance
experiments have demonstrated that S8D, S15D, and
S21D (and their phosphorylated counterparts) differentially disordered the NH2- and COOH-terminal portions of the inactivation domain (Antz et al., 1998
).
Regulation of K+ Channel Inactivation by PKC: A Physiological Scenario
Neurotransmitter receptors that activate phospholipase
C initiate the second messenger cascade that activates
PKC (Nicoll, 1988; Kandel et al., 1995
). Thus, under
physiological conditions, regulation of rapid K+ channel inactivation by PKC may be controlled by a neurotransmitter. We have previously demonstrated that
such a system can be reconstituted in Xenopus oocytes
by coexpressing a metabotropic serotonin receptor that
activates phospholipase C and Kv3.4 K+ channels (Velasco et al., 1998
). This result has important implications because Kv3.4 and metabotropic receptors linked
to phospholipase C (e.g., type 2c serotonin receptor,
muscarinic acetylcholine receptor, glutamate receptor,
etc.) may coexist in the nervous system. In a possible
physiological scenario, the neurotransmitter could be
released from inhibitory neurons acting on a presynaptic terminal. In fact, the terminal areas of major projection tracts in the brain are rich in Kv3.4 (Roeper and
Pongs, 1996
). Thus, activation of a metabotropic receptor in the synapse will trigger the second messenger
cascade that activates PKC. Both PKC
and PKC
are
found in the nervous system, and PKC
seems to be especially abundant in certain terminal areas (Nishizuka,
1988
). Phosphorylation of the inactivation particle in
Kv3.4 by PKC causes slower inactivation of this channel.
As a result, the action potential will be shortened and,
consequently, synaptic transmission will be depressed.
Alternatively, dephosphorylation by phosphatases may
broaden action potentials and enhance synaptic activity.
![]() |
FOOTNOTES |
---|
Address correspondence to Manuel Covarrubias, Department of Pathology, Anatomy and Cell Biology, Jefferson Medical College, 1020 Locust Street JAH 245, Philadelphia, PA 19107. Fax: 215-923-2218; E-mail: manuel.covarrubias{at}mail.tju.edu
Received for publication 20 February 1998 and accepted in revised form 6 May 1998.
We thank Mr. T. Harris for harvesting and injecting oocytes, Drs. A. Wei and A. Matlapudi for helping with site-directed mutagenesis, and Dr. T.B. Vyas for conducting preliminary experiments. We thank Drs. C. Antz, B. Fakler, and R. Horn for insightful discussions and critical comments to an earlier version of the manuscript. We are also grateful to Dr. C.D. Stubbs for providing recombinant PKC isoforms.
This work was supported by National Institutes of Health (NIH) grant NS32337 (M. Covarrubias). E. Beck was supported by NIH Training Grant AA07463.
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Abbreviation used in this paper |
---|
PMA, phorbol 12-myristate-13-acetate.
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