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The Journal of Neuroscience, December 1, 2000, 20(23):8610-8617
The Human Skeletal Muscle Na Channel Mutation R669H Associated
with Hypokalemic Periodic Paralysis Enhances Slow Inactivation
Arie F.
Struyk1,
Kylie
A.
Scoggan3,
Dennis E.
Bulman3, 4, and
Stephen C.
Cannon1, 2
Departments of 1 Neurology, Massachusetts General
Hospital, and 2 Neurobiology, Harvard Medical School,
Boston, Massachusetts 02114, and 3 Department of Medicine,
Division of Neurology, and 4 Ottawa Hospital Research
Institute, Ottawa, Ontario K1H 7W9 Canada
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ABSTRACT |
Missense mutations of the human skeletal muscle voltage-gated Na
channel (hSkM1) underlie a variety of diseases, including hyperkalemic
periodic paralysis (HyperPP), paramyotonia congenita, and
potassium-aggravated myotonia. Another disorder of sarcolemmal excitability, hypokalemic periodic paralysis (HypoPP), which is usually
caused by missense mutations of the S4 voltage sensors of the L-type Ca
channel, was associated recently in one family with a mutation in the
outermost arginine of the IIS4 voltage sensor (R669H) of hSkM1 (Bulman
et al., 1999 ). Intriguingly, an arginine-to-histidine mutation at the
homologous position in the L-type Ca2+ channel
(R528H) is a common cause of HypoPP. We have studied the gating
properties of the hSkM1-R669H mutant Na channel experimentally in human
embryonic kidney cells and found that it has no significant effects on
activation or fast inactivation but does cause an enhancement of slow
inactivation. R669H channels exhibit an ~10 mV hyperpolarized shift
in the voltage dependence of slow inactivation and a twofold to
fivefold prolongation of recovery after prolonged depolarization. In
contrast, slow inactivation is often disrupted in HyperPP-associated Na
channel mutants. These results demonstrate that, in R669H-associated HypoPP, enhanced slow inactivation does not preclude, and may contribute to, prolonged attacks of weakness and add support to previous evidence implicating the IIS4 voltage sensor in
slow-inactivation gating.
Key words:
depolarization; inactivation; Na channel; hypokalemic
periodic paralysis; mutation; skeletal muscle
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INTRODUCTION |
The human skeletal muscle voltage-gated
sodium channel (hSkM1) mediates sarcolemmal excitability because of its
rapid activation and subsequent fast inactivation in response to
membrane depolarization. In addition to its fast gating, which takes
place over milliseconds, the Na channel enters a more prolonged
"slow"-inactivated state, over hundreds of milliseconds to seconds,
in response to sustained membrane depolarizations. The structural basis
of Na channel slow inactivation is not known but appears to be
different from conformational rearrangements associated with fast
inactivation (Rudy, 1978; Cummins and Sigworth, 1996; Featherstone et
al., 1996; Vedantham and Cannon, 1998). Slow inactivation may modulate
Na channel availability, and thus sarcolemmal excitability, in response
to prolonged shifts in membrane potential (Chandler and Meves, 1970;
Almers et al., 1983; Ruff et al., 1988).
Gain-of-function mutations of the skeletal muscle Na channel gene
SCN4A have been identified in a variety of disorders of muscle membrane excitability: potassium-aggravated myotonia,
paramyotonia congenita (PMC), and hyperkalemic periodic paralysis
(HyperPP) (for review, see Cannon, 2000). Attacks of weakness in
HyperPP and PMC are characterized by a depolarization-induced loss of membrane excitability. Depolarization arises from an aberrant inward Na
current conducted by mutant channels with disrupted fast inactivation
(Cannon et al., 1993; Hayward et al., 1996). Slow inactivation is
impaired by three of the known HyperPP/PMC-linked mutations (Cummins
and Sigworth, 1996; Hayward et al., 1997, 1999). This defect is thought
to promote weakness by dampening the ability of the Na channels to turn
off the persistent Na current and thereby to permit rapid (within
seconds) repolarization and recovery of excitability (Ruff, 1994;
Hayward et al., 1997, 1999).
Similar to HyperPP, hypokalemic periodic paralysis (HypoPP) is a rare
autosomal-dominant disorder characterized by intermittent attacks of
weakness, often lasting for hours to days. In HypoPP, however, weakness
is accompanied by a decrease in serum potassium concentration, and
myotonic stiffness does not occur (Rüdel et al., 1984; Cannon,
1998). During an attack, the sarcolemma becomes depolarized and
inexcitable (Rüdel et al., 1984). The basis of this
depolarization remains speculative, although changes in ATP-sensitive K
channels have been promoted recently as direct precipitants (Ruff,
1999; Tricarico et al., 1999). Genetic analysis of HypoPP patients has
identified three missense mutations in the  1 subunit of
the skeletal muscle voltage-gated L-type calcium channel: R528H, R1239H, and R1239G (Fontaine et al., 1994; Jurkat-Rott et al., 1994;
Ptacek et al., 1994). These mutations disrupt the outermost arginine
residues of the S4 membrane-spanning segments of either domain II or IV
of the L-type Ca channel. A variety of gating defects have been
reported for these mutants (Sipos et al., 1995; Lapie et al., 1996;
Morrill et al., 1998; Morrill and Cannon, 1999), with little further
insight into the pathophysiology of the disease.
Recently, two members of a French family were described with HypoPP, in
which no L-type Ca channel mutation was identified. Single-strand
conformational polymorphism analysis revealed an abnormal conformer of
the Na channel gene SCN4A. A missense mutation that converts
an arginine at position 669 to histidine (R669H) was identified that
cosegregated with the disease phenotype (Bulman et al., 1999). This
arginine is conserved in all voltage-gated Na channels, and the R669H
mutation was not found as an incidental polymorphism in an analysis of
100 normal individuals. The Na channel R669H mutation and the Ca
channel R528H mutation both replace the outermost arginine of the IIS4
voltage sensors of the respective channels by histidine.
We have studied the gating properties of the hSkM1-R669H mutant
channels expressed heterologously in human embryonic kidney (HEK)
cells. The kinetics and voltage dependence of activation and fast
inactivation were not significantly altered by this mutation. Surprisingly, slow inactivation was profoundly affected. In contrast to
some HyperPP mutations, in which slow inactivation is disrupted, R669H
enhances slow inactivation, predominantly via prolonged recovery from
slow inactivation and a hyperpolarized shift in the voltage dependence
of slow inactivation. These results help to define the role of slow
inactivation in disorders of muscle membrane excitability and support
previous work implicating the IIS4 voltage sensor in slow-inactivation gating.
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MATERIALS AND METHODS |
Expression of sodium channels. cDNA encoding the
adult isoform of the human muscle sodium channel subunit
hSkM1 (George et al., 1992) was used as a template for in
vitro site-directed mutagenesis using the Transformer kit
(Clontech, Palo Alto, CA). Primers CGTCGACGGATCGGGACATGTCCCGATCCCCTATGG
and GACTGTCTGTCCTACACTCCTTCCG were used for selection and
introduction of the missense mutation, respectively. In addition to
introduction of the arginine-to-histidine missense codon, a
translationally silent EclHK1 site was introduced into the cDNA-coding
sequence by the mutant oligomer for the purpose of rapid screening. The
human  1 subunit cDNA was subcloned into the EcoRI site of
the mammalian expression vector pcDNA1 (McClatchey et al., 1993). An
independently derived hSkM1-R669H Na channel mutant clone (D. Bulman)
was used in identical experimental protocols to confirm the results
seen with our construct.
Cultures of HEK cells and their transient transfection were performed
as described previously (Hayward et al., 1996). Briefly, supercoiled
plasmid DNA encoding 1.25 µg of either wild-type or R669H mutant Na
channel subunits along with 2.5 µg (fourfold molar excess) of the
1 subunit expression plasmid and a CD8 marker were used to transfect
HEK cell monolayers in 35 mm dishes by the calcium phosphate
precipitation method. Two to three days after transfection, the HEK
cells were briefly trypsinized and passaged to a 22 mm round glass
coverslip for electrophysiological recording. Individual
transfection-positive cells were identified by labeling with anti-CD8
antibodies cross-linked to microbeads (Dynal, Great Neck, NY) (Jurman
et al., 1994).
Whole-cell recording. Na currents were measured by the use
of conventional whole-cell recording techniques as described previously (Hayward et al., 1996). Recordings were made with an Axopatch 200A
amplifier (Axon Instruments, Foster City, CA). The output was filtered
at 10 kHz and digitally sampled at 40 kHz using an LM900 interface
(Dagan, Minneapolis, MN). Data were stored to a 486-based computer
using AxoBasic (Axon Instruments) data acquisition software.
Patch electrodes were fabricated from borosilicate capillary tubes with
a multistage puller (Sutter, Novato, CA). The shank of the pipette was
coated with Sylgard, and the tip was heat-polished to a final tip
resistance in the bath solution of 0.5-2.0 M . At least 80% of the
series resistance was compensated by the analog circuitry of the
amplifier, and the leakage conductance was corrected by digital scaling
and subtraction of the passive current elicited by a 25 mV
depolarization from the holding potential. Cells with peak currents of
<1 nA after step depolarization from  120 to 10 mV were excluded to
reduce potential contamination by small endogenous Na currents
occasionally observed in untransfected HEK cells. In addition, cells
with peak currents >20 nA were excluded to reduce series resistance
errors. Only cells with series resistances of <5 M were included in
the data set. Individual cells were allowed to equilibrate for 7-10
min after achieving internal access before acquiring data. We
occasionally observed cells with Na currents that differed from typical
Na channel gating behavior, characterized by higher amplitude
persistent Na current after a 10 msec depolarization and a prolonged
 h. To reduce effects of these modal-gating
shifts, data from these cells were not included in the final analysis.
For all experiments except those using a reversed Na gradient, the
internal pipette solution contained (in mM): 105 CsF, 35 NaCl, 10 EGTA, and 10 HEPES, pH 7.4 by CsOH. Fluoride was used in the
pipette solution to prolong seal stability for the purposes of
slow-inactivation protocols. The bath contained (in mM):
140 NaCl, 4 KCl, 2 CaCl2, 1 MgCl2, 5 glucose, and 10 HEPES, pH 7.4 by NaOH.
For the reversed Na gradient experiments, the internal pipette solution
contained (in mM): 130 NaCl, 10 HEPES, and 10 EGTA; the
bath solution contained (in mM): 150 choline-Cl, 10 HEPES,
2 CaCl2, and 0.2 CdCl2. All
recordings were made at room temperature.
Data analysis. The data were analyzed and displayed by the
use of a combination of computer programs: AxoBasic, Excel (Microsoft Corporation), and Origin (MicroCal). Conductance was calculated as:
where the reversal potential
Erev was measured experimentally for
each cell. Steady-state fast and slow inactivation were fitted to a
Boltzmann function with a nonzero pedestal,
I0, calculated as:
where V1/2 is the half-maximum
voltage and k is the slope factor. The kinetics of fast
inactivation was quantified from single-exponential fits to the
macroscopic current decay and to the relaxation between a closed and
inactivated state revealed by two-pulse protocols at voltages
between 40 and 60 mV. The time constant of the decay was
estimated by fitting macroscopic Na currents I to a single exponential plus a constant term, I , as:
where Imax is the maximal
amplitude and t is the pulse duration.
The time course of recovery from fast inactivation was measured by the
use of a two-pulse protocol. A 40 msec conditioning pulse to 10 mV
was applied to fast inactivate the channels fully, followed by a return
to the recovery potential (between 120 and 80 mV) for a variable
interval. The fraction of available (recovered) channels was assayed
with a test pulse to 10 mV, and the time course of recovery was fit
to the equation:
I is the peak amplitude of the current, which was
normalized to the peak amplitude of a reference current,
Iref, elicited by a pulse to 10 mV
before application of the conditioning pulse. A is the
maximal extent of recovery, t is the recovery pulse
duration, and is the time constant of recovery.
After prolonged depolarization, Na channel availability recovers with a
complex time course characterized by multiple exponential components
(Cummins and Sigworth, 1996). An "intermediate" component, IM, recovers within 100-300 msec,
slow-inactivated channels (IS) recover
within 1-3 sec, and ultraslow-inactivated channels
(IU) recover over minutes. Recovery from
intermediate (IM)- and slow (IS)-inactivated states was measured by
the use of a two-pulse protocol with conditioning pulse lengths of up
to 60 sec. Between trials, channels were allowed to recover fully by
holding the membrane at 120 mV for a period of three times the
conditioning pulse duration (up to 90 sec). For conditioning pulse
lengths of >60 sec, a sequential recovery protocol was used in which a single conditioning pulse was followed by a series of brief test pulses
during the recovery interval. For the sequential recovery protocol,
peak current from each test pulse was normalized to a reference current
(Iref), measured as the mean peak
value from four separate step depolarizations to 10 mV from the
recovery potential, before the conditioning pulse.
Itest/Iref
values were then fit with a two-exponential decay function:
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A1 and
A2 are the amplitudes of the two
components, 1 and 2
are the time constants, t is the recovery time, and
I0 is the fractional current at time
0. Data from cells whose current failed to recover within 10% of the
reference current were discarded.
Symbols with error bars indicate means ± SEM. Statistical
significance was determined by the unpaired t test with
p values noted in the text.
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RESULTS |
HyperPP-associated Na channel mutations have gain-of-function
defects caused by disrupted fast inactivation or, in some cases, by a
hyperpolarized shift in activation (Cannon, 2000). We sought to
determine whether defects of fast gating, and in particular fast
inactivation, might account for the HypoPP phenotype in our family.
Wild-type and R669H mutant Na channel cDNAs under the control of a cytomegalovirus promoter were transiently transfected into
HEK cells, along with the human isoform of the 1 subunit. Whole-cell voltage-clamp recording was used to study the gating characteristics of the channels. On average, cells transfected with
wild-type hSkM1 cDNA had slightly larger peak Na currents (7.0 ± 0.8 nA; n = 36) than did those transfected with R669H
(5.6 ± 0.4 nA; n = 55), but the difference was
not statistically significant (p = 0.10).
Fast-gating behavior
Representative current traces elicited by a series of step
depolarizations from a holding potential of 120 mV to voltages ranging between 75 and +80 mV for wild-type (WT) and R669H
channels are shown in Figure 1. Both the
activation and inactivation properties of the R669H current appeared to
be qualitatively identical to that of wild type.
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Figure 1.
Representative currents from cells expressing WT
or R669H channels. Currents were elicited by a series of voltage steps
from a holding potential of 120 mV to voltages ranging between 75
and +80 mV.
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Three different protocols were used to quantify the kinetics of fast
inactivation. The development of inactivation was measured over the
voltage range 30 to +40 mV by fitting the current decay during a test
depolarization. For membrane potentials between 60 and 40 mV, the
relaxation between closed and fast-inactivated states was measured with
a two-pulse protocol as the time-dependent inactivation of the peak
current during variable duration conditioning pulses. Finally, the time
constant of recovery from fast inactivation was determined by the use
of a two-pulse protocol with a variable recovery gap between the
conditioning and test pulses. Recovery was measured at potentials
ranging between 120 and 80 mV. Figure 2A displays the voltage
dependence of the resulting time constants on a semilogarithmic scale.
Time constants for both entry and recovery were comparable for R669H
channels and wild-type ones. The small increase in observed for
R669H was not statistically different from that of WT. The transition
rate from open to the fast-inactivated state is estimated best from the
asymptotic value of at strongly depolarized potentials. We examined
this microscopic inactivation rate in greater detail by using a
reversed Na gradient (high internal/low external) to shift
Erev away from +50 mV and thereby
increase the amplitude of the Na current. Superposition of individual
trials showed no apparent differences between R669H and WT (Fig.
2B). On average, the single-exponential fit of the current decay elicited by a step depolarization from 120 to +50 mV
yielded identical fast-inactivation time constants for wild-type (0.26 ± 0.008 msec; n = 3) and R669H (0.26 ± 0.007 msec; n = 4) channels.
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Figure 2.
Fast-inactivation kinetics of R669H channels was
indistinguishable from that of wild type. A, Time
constants of fast inactivation at voltages between 30 and +40 mV for
wild-type (open circles; n = 7) and
R669H (closed circles; n = 10)
channels were determined by fitting a single exponential to the
macroscopic current decay during step depolarizations from a holding
potential of 120 mV. At more hyperpolarized potentials (40 and 60
mV), a two-pulse protocol was used to measure the time course of fast
inactivation from closed states for wild-type (open
squares; n = 6) and R669H (closed
squares; n = 7) channels. The kinetics of
recovery from fast inactivation for wild-type (open
triangles; n = 6) and R669H (closed
triangles; n = 6) channels was determined
by the use of a standard two-pulse protocol with a 40 msec conditioning
pulse at 10 mV and recovery potentials of 80, 100, or 120 mV.
B, The limiting rate of macroscopic inactivation at
strongly depolarized potentials was identical for WT (open
circles) and R669H (closed circles) channels.
Superimposed traces show outward Na+
currents recorded with a reversed Na+ gradient (see
Materials and Methods) by depolarization to +50 mV from a holding
potential of 120 mV. The mean time constant of current decay was
identical for wild-type (0.26 ± 0.008 msec; n = 3) and R669H (0.26 ± 0.007 msec; n = 4)
channels.
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The peak conductance-voltage relationship of R669H channels (Fig.
3) had an increased slope and a very small
rightward shift, estimated to be a 3 mV depolarized shift in the
midpoint of the fitted curve in comparison with that of wild type
(Table 1). The voltage dependence of
steady-state fast inactivation was measured as the relative peak
current elicited after a 300 msec conditioning pulse. Data from each
cell were fit with a Boltzmann function, and amplitude-normalized data
from separate cells were pooled as shown by mean values and SEMs in
Figure 3. No significant difference was noted for the midpoint or the
voltage sensitivity between R669H and wild-type channels (Table 1).
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Figure 3.
Voltage dependence of fast gating was not affected
by the R669H mutation. The voltage dependence of fast inactivation of
wild-type (open circles; n = 8) and
R669H (closed circles; n = 11)
channels was measured as the peak INa
elicited by a test depolarization to 10 mV, after 300 msec
conditioning pulses to potentials ranging from 140 to 35 mV.
Fitted curves are derived from mean parameter values of
 2-minimized fits to a Boltzmann function plus a constant
for each cell. The conductance-voltage curves for
wild-type (open squares; n = 8) and R669H (closed squares;
n = 16) channels were derived from the peak
INa elicited by a series of 10 msec test
pulses to the potentials ranging from 75 to +80 mV. Reversal
potential was fitted as an independent parameter for each cell.
Curves for wild type and R669H are computed from means
of parameter values estimated from 2-minimized fits of a
Boltzmann function for each cell. Mean parameter values and SEMs are
listed in Table 1.
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Slow inactivation
Studies of HyperPP mutants suggest that, whereas defects in fast
gating may precipitate episodes of paralysis, disruption of Na channel
slow inactivation may contribute to the maintained depolarization
responsible for prolonged attacks (Hayward et al., 1999). We therefore
sought to characterize slow inactivation of the R669H mutant.
The rate of entry to the slow-inactivated state was measured by the use
of a two-pulse protocol as described in the Figure 4A inset. Cells
were held at 120 mV, and a conditioning pulse of varying duration was
applied. A 20 msec return to Vhold was interposed between the conditioning and test pulses, which allowed complete recovery from fast inactivation. The current elicited by a
subsequent pulse to 10 mV was normalized by the response to a
preconditioning reference pulse, plotted against the conditioning pulse
duration, and fit with a single-exponential function. The rate of entry
to slow inactivation was similar for wild-type and R669H channels at
voltages between 80 and 10 mV (Fig. 4). The extent of slow
inactivation, however, was greater for R669H as shown by the lower
relative INa after long conditioning
pulses.
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Figure 4.
Entry to slow inactivation was enhanced for R669H
channels. Entry to slow inactivation was measured as the peak
INa elicited by a test depolarization to
10 mV, after a conditioning pulse of varying duration. Current
amplitudes were normalized to a reference peak,
INa, elicited by a depolarization to
10 mV from a holding potential of 120 mV. Conditioning and test
pulses were separated by a 20 msec repolarization to 120 mV to
allow for full recovery from fast inactivation (A,
inset, pulse protocol). Data for wild type (open
circles; n = 4) and R669H (closed
circles; n = 9) were fit to a
single-exponential relaxation to a steady-state value,
S. Curves were generated
from means of parameter values. A,
Vcond = 10 mV; WT, = 390 ± 140 msec, S = 0.18 ± 0.03; R669H, = 510 ± 74 msec,
S = 0.09 ± 0.01. B, Vcond = 40 mV; WT,
= 2400 ± 60 msec, S = 0.29 ± 0.03; R669H, = 2400 ± 120 msec,
S = 0.18 ± 0.01. C, Vcond = 80 mV; WT,
= 7100 ± 2200 msec, S = 0.88 ± 0.03; R669H, = 4200 ± 800 msec,
S =0.65 ± 0.05.
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The voltage dependence of slow inactivation was determined by using a
40 sec conditioning pulse (Fig. 5,
inset), which was sufficiently long to allow slow
inactivation to approach steady state, as shown in Figure 4. A 20 msec
gap at 120 mV was used to allow channels to recover from fast
inactivation, and the fraction of channels not slow inactivated was
measured as the relative peak current elicited by a subsequent test
depolarization to 10 mV. Slow inactivation was enhanced by R669H. The
steady-state voltage dependence for R669H was shifted by 10 mV in the
hyperpolarized direction compared with wild-type channels (Table
2), and the maximal extent of slow
inactivation at strongly depolarized potentials was greater (Fig.
5, smaller relative current,).
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Figure 5.
Steady-state slow inactivation was enhanced for
R669H channels. The voltage dependence of slow inactivation was
measured as the peak INa elicited by a test
depolarization to 10 mV, after a 40 sec conditioning pulse to
voltages ranging from 130 to 10 mV for wild type (open
circles; n = 7) and R669H (closed
circles; n = 10). Current amplitudes were
normalized to a reference peak, INa,
elicited by a depolarization to 10 mV from a holding potential of
120 mV. Conditioning and test pulses were separated by a 20 msec
repolarization to 120 mV to allow recovery from fast
inactivation (inset, pulse protocol). Smooth
curves for wild-type data (dashed line) and
R669H (solid line) are computed from means of parameter
values estimated from 2-minimized fits to a Boltzmann
function plus a constant term for each cell. Parameter values are
listed in Table 2.
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With prolonged depolarization, Na channel inactivation is a complex
process with multiple kinetically defined states with different entry
and recovery rates (Cummins and Sigworth, 1996). Variations in the
length of the conditioning pulse alter the population of these states,
as evidenced by changes in the relative amplitudes of the
multiexponential relaxation components (Hayward et al., 1997). We
designed protocols to examine the recovery of channel availability
after a wide range of conditioning pulse durations. Two different pulse
protocols were used: two pulse and sequential. The two-pulse protocol
was used for conditioning pulses of durations between 40 msec and 60 sec (Fig. 6, inset ). Full
recovery between trials was verified by monitoring the current elicited
by a reference pulse to 10 mV that preceded the conditioning pulse.
The recovery interval between trials was generally three times the
duration of the conditioning pulse, and peak current during the
reference pulse did not vary by >5%. Recovery curves from 40 msec
conditioning pulses for both wild-type and R669H channels were well fit
with a single exponential (Fig. 6A), consistent with
the data from Figure 4 suggesting that a negligible fraction of the
channels enter a slow-inactivated state within that time. The time
constant for recovery from fast inactivation at 120 mV was similar
for WT (3.2 ± 0.49 msec; n = 6) and R669H
(4.0 ± 0.38 msec; n = 13) channels. For
conditioning pulses of 1 sec or longer (Fig. 6B,C), however, a double-exponential relaxation function was required to
obtain a reasonable fit. After a 1 sec conditioning pulse (Fig. 6B), ~60% of WT and R669H channels recovered
within 10 msec, and we interpret this as the fraction of channels that
were fast inactivated. The second component recovered more slowly for
R669H (530 ± 65 msec; n = 9) than for WT
(110 ± 12 msec; n = 11) channels. This component
of recovery (typically 100-300 msec at 120 mV) is thought to reflect
a population of channels that is in an intermediate-inactivated state, designated IM, and the data in
Figure 6, B and C, show that recovery from
IM was prolonged by R669H. The time course of recovery after a 60 sec conditioning pulse (Fig. 6C) was
also well fit by a two-exponential relaxation, plus a small offset to
account for the small fraction of channels (~10-15%) that recovered before the first time point at 10 msec. The bulk of WT and R669H channels recovered with the kinetics of
IM, and like the data after a 1 sec
conditioning pulse (Fig. 6B), recovery of this
component was impeded for R669H (930 ± 140 msec) compared with WT
(200 ± 18 msec). The second exponential component in Figure
6C recovered on the order of seconds and reflects the
fraction of slow-inactivated channels
(IS). Recovery of the
IS component is also prolonged by R669H
(16 ± 5.5 sec; n = 5) compared with WT (4.4 ± 0.86 sec; n = 6). The estimate for the time constant
of the IS component is limited by its
small amplitude, ~10% of the total current. In the next set of
experiments even longer duration conditioning pulses of several minutes
were used to obtain a better measure of the rate of recovery for the
IS component.
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Figure 6.
Recovery from an intermediate inactivation state
(IM) was prolonged for R669H channels.
In A-C, a two-pulse protocol
(inset) was used to measure recovery from
inactivation for wild-type (open circles) and R669H
(closed circles) channels. The relative amplitude of
different kinetic components of recovery varied with the duration of
the conditioning pulse to 10 mV of 40 msec (A),
1 sec (B), or 60 sec (C).
The time course of recovery was measured as the relative peak
INa at 10 mV, as a function of the
recovery interval at 120 mV. Channels were allowed to recover fully
at 120 mV between trials, as ascertained by the reference peak
INa measured by a pulse to 10 mV before
application of the conditioning pulse. The amplitude of this
Iref was used to compute the relative degree
of recovery for the subsequent test pulse to 10 mV. A,
After a 40 msec conditioning pulse, recovery was well fit by a single
exponential that was comparable for WT (3.2 ± 0.49 msec;
n = 6) and R669 (4.0 ± 0.38 msec;
n = 13) channels. B, Recovery after
a 1 sec conditioning pulse had two exponential components. The fast
component was similar for WT and R669H channels, whereas the slower one
(IM) was clearly prolonged for R669H
(530 ± 65 msec; n = 9) compared with WT
(110 ± 12 msec; n = 11). C,
After a 60 sec conditioning pulse, the time course of recovery had
three components: fast (<10 msec; ~15% of total; not resolved by
this protocol), intermediate (100-500 msec; ~70% of response), and
slow (5-15 sec; ~15% of response). A double-exponential fit of the
data was used to estimate the time constants of the intermediate and
slow components. As in B, recovery of the intermediate
component was markedly slower for R669H than for WT, and recovery of
the slow component was moderately prolonged [16 ± 5.5 sec
(n = 5) for R669H compared with 4.4 ± 0.86 sec (n = 6) for WT].
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Because conditioning pulse durations >1 min and their associated slow
recovery times are impractical for two-pulse protocols, a sequential
protocol with a single conditioning pulse followed by a series of short
test depolarizations was used to monitor the time course of recovery
after such prolonged conditioning pulses (Fig.
7). The brief and relatively infrequent test
depolarizations do not alter the recovery from
IM and slower components, as judged by
pilot studies showing indistinguishable recovery curves using conventional two-pulse or this sequential protocol (data not shown). The peak INa elicited from these test
depolarizations was normalized to the reference "maximal"
INa measured as the average response from four step depolarizations preceding the conditioning pulse. After
a 240 sec conditioning pulse, ~60% of the channels recovered with a
time course typical of IM, 35% recovered
with the kinetics of IS, and ~5% of the
channels were fast inactivated only (Fig. 7). As shown by the first
data point in the recovery curve, the fraction of channels fast
inactivated only was smaller for R669H (0.05 ± 0.01;
n = 11) compared with WT (0.09 ± 0.01;
n = 7). This difference is consistent with the reduced
availability of R669H channels in the "steady-state" protocol of
Figure 5. As in the two-pulse protocol of Figure 6, data from the
sequential protocol show a dramatic slowing for the recovery of the
IM component of R669H (1600 ± 220 msec; n = 11) compared with that of WT (230 ± 21 msec; n = 7). The second exponential component of
recovery, corresponding to IS, was also
slowed for R669H (37 ± 4.0 sec compared with 15 ± 0.7 sec
for WT), but the relative change was not as great as that observed for
the IM component. During these very prolonged protocols, we noted a tendency for the peak amplitude of
INa during recovery eventually to
become slightly greater than the mean value measured for the reference
pulses. This increase presumably represents further recovery of
chronically inactivated channels, despite our waiting for at least 10 min at 120 mV before starting the sequential recovery protocol.
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Figure 7.
Recovery after long-duration conditioning pulses
was prolonged for R669H mutants. A 240 sec conditioning pulse to 10
mV was applied from a resting potential of 120 mV. Recovery from
inactivation for wild-type (open circles) and R669H
(closed circles) channels was measured by determining
peak INa during 5 msec test depolarizations
to 10 mV from the recovery potential of 120 mV at the times
indicated. Peak amplitudes were normalized to a reference response
obtained as the mean of four separate pulses to 10 mV elicited before
the conditioning pulse (see inset, pulse protocol).
Because relaxation from the various inactivated states varied over five
orders of magnitude, a logarithmic scale is used to show the time
course of recovery. Recovery of the intermediate component was
sevenfold slower for R669H than for WT, whereas the slow component was
prolonged only approximately twofold. Smooth curves for
wild type (dashed line) and R669H (solid
line) were generated using means of parameter values estimated
from [chi]2-minimized fits of a double-exponential decay
function plus a constant, S, for each
cell. The two exponentials represent recovery of the intermediate- and
slow-inactivated components, and the constant term is the proportion of
channels that fully recovered within 20 msec at 120 mV. The relative
amplitudes and time constants were as follows: WT,
S = 0.091 ± 0.007, A1 = 0.59 ± 0.04, 1 = 230 ± 21 msec, A2 = 0.33 ± 0.02, 2 = 15000 ± 720 msec, n = 7; R669H, S = 0.051 ± 0.007, A1 = 0.61 ± 0.05, 1 = 1600 ± 220 msec, A2 = 0.41 ± 0.04, 2 = 37000 ± 4000 msec, n = 11.
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Composite data from all conditioning pulse durations revealed that
wild-type channels relaxed from IM to the
resting/available state with a time constant of 190 ± 23 msec
(n = 28) and from the slowly developing
(IS) state with a relaxation time constant of 11 ± 3.0 sec (n = 17). In comparison, R669H
channels exhibited relaxation time constants of 900 ± 97 msec
(n = 33) or approximately fivefold longer than that of
wild type for the IM component
(p < 0.001) and 23 ± 4.8 sec
(n = 24) or approximately twofold longer than that of
wild type (p < 0.005) for the
IS component. There was no difference in
the time constant for recovery of the fast component (3.5 ± 0.17 msec for WT and 4.2 ± 0.15 msec for R669H). These estimates were
obtained by simultaneously fitting the recovery data after the 40 msec,
1 sec, 60 sec, and 240 sec conditioning pulses with a set of four
triple-exponential curves that were constrained to have the same values
for the time constants. The relative amplitude of each component varied
with the conditioning pulse duration [for details, see Hayward et al.
(1996)].
The excitability defect in HypoPP muscle occurs in association with
hypokalemia (serum [K+] of <2.5
mM). We tested whether the gating abnormalities observed for R669H were sensitive to extracellular
[K+]. Reducing the bath
[K+] from 4 to 1 mM did not
alter the gating of R669H or of WT channels. In R669H channels,
recovery of the IM and
IS components were still slowed fivefold
and twofold, respectively (data not shown).
Use-dependent inactivation during a pulse train
Enhanced slow inactivation of Na channels has been shown to
augment the use-dependent reduction of peak
INa during prolonged trains of brief
depolarizations (Wang and Wang, 1997), such as might occur in skeletal
muscle during vigorous activity. We tested whether use-dependent
inactivation of Na channels during a pulse train was enhanced in R669H
channels. Depolarizing pulses to +10 mV for 3 msec were applied at 5 or
20 Hz, from a holding potential of 90 mV. Figure
8A shows that a 5 Hz train
applied for 2 min produced a modest degree of use dependence for WT
channels (relative peak current, 0.80 ± 0.03; n = 5), whereas a significantly greater fraction of the R669H channels
became unavailable in response to the same train (relative peak
current, 0.65 ± 0.03; n = 5). This
difference was more pronounced with higher-frequency pulse trains. The
use-dependent reduction of INa during
a 20 Hz pulse train was two times greater for R669H channels (relative
residual current, 0.25 ± 0.03; n = 6) than for WT
ones (residual relative current, 0.53 ± 0.04; n = 6).
View larger version (19K):
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|
Figure 8.
Use-dependent inactivation of
INa during pulse trains was greater for
R669H than for wild-type channels. Trains of 3 msec pulses to +10 mV
from a holding potential of 90 mV were performed at 5 Hz
(A) and 20 Hz (B). Peak
INa for wild-type (open
circles; n = 6) and R669H (closed
circles; n = 6) channels is plotted against
the cumulative duration of the pulse train for every 10th pulse
(A) or 40th pulse (B).
Amplitudes were normalized to the value measured for the first pulse in
the train.
|
|
| |
DISCUSSION |
We studied the gating properties of the skeletal muscle Na channel
mutation R669H, a rare cause of hypokalemic periodic paralysis located
at the outermost charged residue of the IIS4 voltage sensor. In
contrast to Na+ channel mutations
associated with paralytic attacks in hyperkalemic periodic paralysis or
paramyotonia congenita that disrupt fast inactivation and, in some
cases, also disrupt slow inactivation, we found that R669H has
negligible effects on fast gating but profoundly enhances slow
inactivation. Slow inactivation was augmented in R669H by an ~10 mV
hyperpolarizing shift in voltage dependence, by more complete
steady-state slow inactivation at depolarized potentials, and by a much
slower rate of recovery. By using conditioning pulses of different
lengths, we were able to demonstrate that the prolonged time course of
R669H recovery was caused by a fivefold to sevenfold slower recovery of
the intermediate component of inactivation and a more modest (twofold)
prolongation of recovery from the slow-inactivated state. The rate of
entry to the intermediate- and slow-inactivated states was not
significantly altered.
Role of IIS4 in Na+ channel gating
The effects of the R669H mutation on gating reported here are
consistent with results of other mutagenesis studies performed on
distal IIS4 residues in both skeletal muscle and other isoforms of the
Na+ channel. A charge-neutralizing
arginine-to-cysteine substitution at the same position in hSkM1 (R669C)
caused only a modest depolarizing shift of the
G-V relationship of 5-7 mV (Mitrovic et al.,
1998; Groome et al., 1999), similar to the small 3 mV rightward shift reported here. In the human cardiac Na channel isoform hH1, mutation of
the outermost arginine in IIS4 to histidine caused a similar modest
depolarizing shift in the conductance-voltage relationship and no
change in the voltage dependence of inactivation (Chen et al., 1996).
Studies of slow inactivation were not reported for the above mutations.
Kontis and Goldin (1997) studied slow inactivation of the rat
brain IIa Na channel in which the second outermost positive charge of
IIS4 was neutralized by substitution with tryptophan. In agreement with
our findings for R669H, the voltage dependence of slow inactivation was
shifted to the left, although in their mutation the shift was >10 mV
(Kontis and Goldin, 1997). This same study demonstrated that the
voltage dependence of slow inactivation was preserved in a
charge-conserving arginine-to-lysine missense mutation at the same
position, supporting the idea that the effects on slow inactivation
seen in our work result from changes in the charge density within IIS4.
Histidine has a pK of 6-9, depending on its immediate environment;
thus substitution of histidine for arginine is expected to result in at
least a partial neutralization of the positive charge normally present at this position. However, we did not observe any change in fast- or
slow-gating kinetics or voltage dependence when the external pH was
varied between 6.5 and 8.0 for R669H Na channels (data not shown).
Several lines of experimental evidence have suggested that the S4
segments do not contribute equally to voltage-dependent gating in
Na+ channels. Mutations in IVS4
preferentially disrupt fast inactivation, without altering activation
(Chahine et al., 1994; Chen et al., 1996). Using site-directed
fluorescent labeling of each S4 segment, Cha et al. (1999) showed that
changes in fluorescence signals from domains I and II voltage sensors
follow activation and are not immobilized by fast inactivation, whereas
signals from domains III and IV S4 segments tracked fast inactivation
and its associated charge immobilization. Kontis and Goldin (1997)
examined the effects of charge-neutralizing mutations in the second and
fourth positively charged residues in each of the four S4 domains of
the RBIIa Na+ channel. Mutations in the
second position of domains I and II, but not III or IV, caused
pronounced shifts in the voltage dependence of slow inactivation,
whereas only small changes were observed for activation or fast
inactivation. Together with our findings of enhanced slow inactivation
by R669H in the first charged residue of IIS4, these data suggest that
the S4 segment in domain II has a specialized role in regulating the
voltage dependence of slow inactivation, particularly the charged
residues at the first and second positions.
Pathogenesis of hypokalemic periodic paralysis
How enhanced slow inactivation caused by the R669H mutation
results in the HypoPP phenotype is not immediately evident. Individuals with R669H-associated HypoPP have prolonged (hours to days) episodes of
paralysis in conjunction with a reduction in serum potassium concentration, identical to the clinical presentation of HypoPP associated with L-type Ca2+ channel
mutations (Bulman et al., 1999). The final common pathway in the
development of weakness, for both Ca2+ and
Na+ channel-based HypoPP, is most likely a
loss of excitability because of fiber depolarization and inactivation
of Na+ channels. In vitro
recordings of HypoPP muscle fibers from patients with L-type Ca channel
mutations demonstrate that sustained, aberrant depolarization of the
sarcolemma forms the basis for these attacks of paralysis (Rüdel
et al., 1984). HypoPP fibers depolarize when the bath
[K+] is lowered from 4.0 to 2.5 mM, whereas under the same conditions normal
fibers hyperpolarize by several millivolts. The hypokalemia-induced depolarization in HypoPP fibers is not prevented by blockers of Na+ (TTX) or L-type
Ca2+ (nitrendipine) channels (Rüdel
et al., 1984; Ruff, 1999). Microelectrode recordings of fibers from the
hSkM1-R669H family have never been made, and consequently the resting
potential of the affected fibers during attacks is unknown. It remains
a possibility that changes in muscle membrane excitability in this
family differ fundamentally from the more common L-type
Ca2+ channel-associated HypoPP.
Nevertheless, the clinical phenotype of hSkM1-R669H is sufficiently
similar to the Ca2+ channel-linked
disorder that it is unlikely that significant differences in basic
pathophysiology exist.
Our results are surprising in view of recent proposals about the role
of slow inactivation in diseases of muscle membrane excitability. A
disruption of slow inactivation is thought to promote attacks of
depolarization-induced weakness in HyperPP and PMC by impeding the
ability of slow inactivation to attenuate aberrant persistent
Na+ currents arising from impaired fast
inactivation or left-shifted activation (Hayward et al., 1999). In
contrast, our results suggest that enhanced slow inactivation may
contribute to, or at least does not preclude, the phenotype of
prolonged weakness. Similarly, Bendahhou et al. (1999) reported
recently a HyperPP-associated mutation, I1495F, in which prolonged
weakness was a prominent component of the phenotype and enhanced slow
inactivation was exhibited. The enhanced slow inactivation found in
R669H and I1495H could result in a loss of muscle excitability. The
availability of Na+ channels at the
resting potential would be decreased, and the decrease would be
exacerbated by even a small depolarization of Vrest. Although this is a plausible
mechanism for a loss of excitability, it does not explain how
hypokalemia triggers attacks or why
Vrest is (presumably) depolarized
during the attacks. We used trains of short depolarizations comparable
with the duration of single action potentials and demonstrated that
R669H channels might become "trapped" in a slow-inactivated state
in a use-dependent manner over several minutes. This, too, may
contribute to the exercise-induced loss of excitability and weakness.
Previous attempts to link functional defects mechanistically in
HypoPP-associated mutant L-type Ca2+
channels to the electrophysiological changes in muscle during an attack
have met with little success. A variety of alterations in the behavior
of mutant L-type Ca2+ channels have been
described. A common theme among this diversity is that external
Ca2+ entry is predicted to be reduced in
HypoPP muscle. Ca2+ current density is
reduced in cultured human myotubes heterozygous for the R1239H (Sipos
et al., 1995) or the R528H (Morrill et al., 1998) mutation.
Heterologous expression studies of R528H in mouse L cells (Lapie et
al., 1996) or of R528H, R1239H, or R1239G channels in
Xenopus oocytes (Morrill and Cannon, 1999) have also found a
reduced current density for mutants compared with WT. Moreover, in
oocytes we observed slowed activation for all three L-type channel
mutants and accelerated deactivation for the R1239 mutant (Morrill and
Cannon, 1999). The reduced current density and altered kinetics would
act synergistically to reduce Ca2+ entry
into HypoPP muscle fibers. The link between altered
Ca2+ entry and the episodic membrane
depolarization and hypokalemia because of the trapping of
K+ in muscle remains to be discerned.
Several investigators have promoted the idea that secondary changes in
ATP-regulated K+ channel activity might be
important in HypoPP attacks. Tricarico et al. (1999) found abnormal
subconductance states and a reduced open probability of K-ATP channels
in HypoPP fibers with the R528H L-type
Ca2+ channel mutation. In another study of
HypoPP fibers with the R528H mutation, Ruff (1999) showed that insulin
potentiates the aberrant depolarization by reducing an inwardly
rectifying K conductance. A decrement in the resting
K+ conductance is consistent with both
hypokalemia because of the trapping of K+
in the myoplasm and a depolarized shift in
Vrest.
Additional note
Since the submission of this manuscript, Jurkat-Rott et al. (2000)
have identified two additional missense mutations in an adjacent
arginine (R672H and R672G) from five families with HypoPP phenotypes.
Heterologous expression studies showed a reduced
Na+ current density and enhanced
inactivation for both mutants. Thus our findings and those of
Jurkat-Rott both imply that a reduced Na+
current density contributes to the pathogenesis of HypoPP.
| |
FOOTNOTES |
Received June 26, 2000; revised Sept. 11, 2000; accepted Sept. 15, 2000.
This work was supported by grants from the National Institutes of
Health, the National Institute of Arthritis and Musculoskeletal and
Skin Diseases Grant AR42703 to S.C.C. and the National Institute of
Neurological Diseases and Stroke Grant K08-NS02137 to A.F.S., and by a
grant from the Medical Research Council of Canada to D.E.B.
Correspondence should be addressed to Dr. Stephen C. Cannon, Department
of Neurobiology, Massachusetts General Hospital, Edwards Research
Building 417, Fruit Street, Boston, MA 02114. E-mail: cannon{at}helix.mgh.harvard.edu.
| |
REFERENCES |
-
Almers W,
Stanfield PR,
Stühmer W
(1983)
Slow changes in currents through sodium channels in frog muscle membrane.
J Physiol (Lond)
339:253-271[Abstract/Free Full Text].
-
Bendahhou S,
Cummins TR,
Tawil R,
Waxman SG,
Ptacek LJ
(1999)
Activation and inactivation of the voltage-gated sodium channel: role of segment S5 revealed by a novel hyperkalaemic periodic paralysis mutation.
J Neurosci
19:4762-4771[Abstract/Free Full Text].
-
Bulman DE,
Scoggan KA,
van Oene MD,
Nicolle MW,
Hahn AF,
Tollar LL,
Ebers GC
(1999)
A novel sodium channel mutation in a family with hypokalemic periodic paralysis.
Neurology
53:1932-1936[Abstract/Free Full Text].
-
Cannon SC
(1998)
Ion channel defects in the hereditary myotonias and periodic paralyses.
In: Molecular neurology (Martin JP,
ed), pp 257-277. New York: Scientific American.
-
Cannon SC
(2000)
Spectrum of sodium channel disturbances in the nondystrophic myotonias and periodic paralyses.
Kidney Int
57:772-779[Web of Science][Medline].
-
Cannon SC,
Brown Jr RH,
Corey DP
(1993)
Theoretical reconstruction of myotonia and paralysis caused by incomplete inactivation of sodium channels.
Biophys J
65:270-288[Web of Science][Medline].
-
Cha A,
Ruben PC,
George ALJ,
Fujimoto E,
Bezanilla F
(1999)
Voltage sensors in domains III and IV, but not I and II, are immobilized by Na+ channel fast inactivation.
Neuron
22:73-87[Web of Science][Medline].
-
Chahine M,
George Jr AL,
Zhou M,
Ji S,
Sun W,
Barchi RL,
Horn R
(1994)
Sodium channel mutations in paramyotonia congenita uncouple inactivation from activation.
Neuron
12:281-294[Web of Science][Medline].
-
Chandler WK,
Meves H
(1970)
Slow changes in membrane permeability and long-lasting action potentials in axons perfused with fluoride solutions.
J Physiol (Lond)
211:707-728[Abstract/Free Full Text].
-
Chen LQ,
Santarelli V,
Horn R,
Kallen RG
(1996)
A unique role for the S4 segment of domain 4 in the inactivation of sodium channels.
J Gen Physiol
108:549-556[Abstract/Free Full Text].
-
Cummins TR,
Sigworth FJ
(1996)
Impaired slow inactivation in mutant sodium channels.
Biophys J
71:227-236[Web of Science][Medline].
-
Featherstone DE,
Richmond JE,
Ruben PC
(1996)
Interaction between fast and slow inactivation in SkM1 sodium channels.
Biophys J
71:3098-3109[Web of Science][Medline].
-
Fontaine B,
Vale-Santos J,
Jurkat-Rott K,
Reboul J,
Plassart E,
Rime CS,
Elbaz A,
Heine R,
Guimaraes J,
Weissenbach J,
Baumann N,
Fardeau M,
Lehmann-Horn F
(1994)
Mapping of the hypokalaemic periodic paralysis (HypoPP) locus to chromosome 1q31-32 in three European families.
Nat Genet
6:267-272[Web of Science][Medline].
-
George Jr AL,
Komisarof J,
Kallen RG,
Barchi RL
(1992)
Primary structure of the adult human skeletal muscle voltage-dependent sodium channel.
Ann Neurol
31:131-137[Web of Science][Medline].
-
Groome JR,
Fujimoto E,
George AL,
Ruben PC
(1999)
Differential effects of homologous S4 mutations in human skeletal muscle sodium channels on deactivation gating from open and inactivated states.
J Physiol (Lond)
516:687-698[Abstract/Free Full Text].
-
Hayward LJ,
Brown Jr RH,
Cannon SC
(1996)
Inactivation defects caused by myotonia-associated mutations in the sodium channel III-IV linker.
J Gen Physiol
107:559-576[Abstract/Free Full Text].
-
Hayward LJ,
Brown Jr RH,
Cannon SC
(1997)
Slow inactivation differs among mutant Na channels associated with myotonia and periodic paralysis.
Biophys J
72:1204-1219[Web of Science][Medline].
-
Hayward LJ,
Sandoval GM,
Cannon SC
(1999)
Defective slow inactivation of sodium channels contributes to familial periodic paralysis.
Neurology
52:1447-1453[Abstract/Free Full Text].
-
Jurkat-Rott K,
Lehmann-Horn F,
Albaz A,
Heine R,
Gregg RG,
Hogan K,
Powers PA,
Lapie P,
Vale-Santos JE,
Weissenback J,
Fontaine B
(1994)
A calcium channel mutation causing hypokalemic periodic paralysis.
Hum Mol Genet
3:1415-1419[Abstract/Free Full Text].
-
Jurkat-Rott K,
Mitrovic N,
Hang C,
Kouzmekine A,
Iaizzo P,
Herzog J,
Lerche H,
Nicole S,
Vale-Santos J,
Chauveau D,
Fontaine B,
Lehmann-Horn F
(2000)
Voltage-sensor sodium channel mutations cause hypokalemic periodic paralysis type 2 by enhanced inactivation and reduced current.
Proc Natl Acad Sci USA
97:9549-9554[Abstract/Free Full Text].
-
Jurman ME,
Boland LM,
Liu Y,
Yellen G
(1994)
Visual identification of individual transfected cells for electrophysiology using antibody-coated beads.
Biotechniques
17:874-881.
-
Kontis KJ,
Goldin AL
(1997)
Sodium channel inactivation is altered by substitution of voltage sensor positive charges.
J Gen Physiol
110:403-413[Abstract/Free Full Text].
-
Lapie P,
Goudet C,
Nargeot J,
Fontaine B,
Lory P
(1996)
Electrophysiological properties of the hypokalaemic periodic paralysis mutation (R528H) of the skeletal muscle alpha 1s subunit as expressed in mouse L cells.
FEBS Lett
382:244-248[Web of Science][Medline].
-
McClatchey AI,
Cannon SC,
Slaugenhaupt SA,
Gusella JF
(1993)
The cloning and expression of a sodium channel beta 1-subunit cDNA from human brain.
Hum Mol Genet
2:745-749[Abstract/Free Full Text].
-
Mitrovic N,
George Jr AL,
Horn R
(1998)
Independent versus coupled inactivation in sodium channels. Role of the domain 2 S4 segment.
J Gen Physiol
111:451-462[Abstract/Free Full Text].
-
Morrill JA,
Cannon SC
(1999)
Effects of mutations causing hypokalaemic periodic paralysis on the skeletal muscle L-type Ca2+ channel expressed in Xenopus laevis oocytes.
J Physiol (Lond)
520:321-336[Abstract/Free Full Text].
-
Morrill JA,
Brown Jr RH,
Cannon SC
(1998)
Gating of the L-type Ca channel in human skeletal myotubes: an activation defect caused by the hypokalemic periodic paralysis mutation R528H.
J Neurosci
18:10320-10334[Abstract/Free Full Text].
-
Ptacek LJ,
Tawil R,
Griggs RC,
Engel AG,
Layzer RB,
Kwiecinski H,
McManis PG,
Santiago L,
Moore M,
Fouad G,
Bradley P,
Leppert MF
(1994)
Dihydropyridine receptor mutations cause hypokalemic periodic paralysis.
Cell
77:863-868[Web of Science][Medline].
-
Rüdel R,
Lehmann-Horn F,
Ricker K,
Kuther G
(1984)
Hypokalemic periodic paralysis: in vitro investigation of muscle fiber membrane parameters.
Muscle Nerve
7:110-120[Web of Science][Medline].
-
Rudy B
(1978)
Slow inactivation of the sodium conductance in squid giant axons. Pronase resistance.
J Physiol (Lond)
283:1-21[Abstract/Free Full Text].
-
Ruff RL
(1994)
Slow Na+ channel inactivation must be disrupted to evoke prolonged depolarization-induced paralysis.
Biophys J
66:542-545[Web of Science][Medline].
-
Ruff RL
(1999)
Insulin acts in hypokalemic periodic paralysis by reducing inward rectifier K+ current.
Neurology
53:1556-1563[Abstract/Free Full Text].
-
Ruff RL,
Simoncini L,
Stuhmer W
(1988)
Slow sodium channel inactivation in mammalian muscle: a possible role in regulating excitability.
Muscle Nerve
11:502-510[Web of Science][Medline].
-
Sipos I,
Jurkat-Rott K,
Harasztosi C,
Fontaine B,
Kovacs L,
Melzer W,
Lehmann-Horn F
(1995)
Skeletal muscle DHP receptor mutations alter calcium currents in human hypokalaemic periodic paralysis myotubes.
J Physiol (Lond)
483:299-306[Abstract/Free Full Text].
-
Tricarico D,
Capriulo R,
Conte Camerino D
(1999)
Insulin modulation of ATP-sensitive K+ channel of rat skeletal muscle is impaired in the hypokalaemic state.
Pflügers Arch
437:235-240[Web of Science][Medline].
-
Vedantham V,
Cannon S
(1998)
Slow inactivation does not affect movement of the fast inactivation gate in voltage-gated Na+ channels.
J Gen Physiol
111:83-93[Abstract/Free Full Text].
-
Wang SY,
Wang GK
(1997)
A mutation in segment I-S6 alters slow inactivation of sodium channels.
Biophys J
72:1633-1640[Web of Science][Medline].
Copyright © 2000 Society for Neuroscience 0270-6474/00/20238610-08$05.00/0
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W. Ulbricht
Sodium Channel Inactivation: Molecular Determinants and Modulation
Physiol Rev,
October 1, 2005;
85(4):
1271 - 1301.
[Abstract]
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S Talon, M.-A Giroux-Metges, J.-P Pennec, C Guillet, H Gascan, and M Gioux
Rapid protein kinase C-dependent reduction of rat skeletal muscle voltage-gated sodium channels by ciliary neurotrophic factor
J. Physiol.,
June 15, 2005;
565(3):
827 - 841.
[Abstract]
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M. Bouhours, D. Sternberg, C.-S. Davoine, X. Ferrer, J. C. Willer, B. Fontaine, and N. Tabti
Functional characterization and cold sensitivity of T1313A, a new mutation of the skeletal muscle sodium channel causing paramyotonia congenita in humans
J. Physiol.,
February 1, 2004;
554(3):
635 - 647.
[Abstract]
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A. Tsujino, C. Maertens, K. Ohno, X.-M. Shen, T. Fukuda, C. M. Harper, S. C. Cannon, and A. G. Engel
Myasthenic syndrome caused by mutation of the SCN4A sodium channel
PNAS,
June 10, 2003;
100(12):
7377 - 7382.
[Abstract]
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A. F. Struyk and S. C. Cannon
Slow Inactivation Does Not Block the Aqueous Accessibility to the Outer Pore of Voltage-gated Na Channels
J. Gen. Physiol.,
September 30, 2002;
120(4):
509 - 516.
[Abstract]
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K. Hilber, W. Sandtner, O. Kudlacek, B. Schreiner, I. Glaaser, W. Schutz, H. A. Fozzard, S. C. Dudley, and H. Todt
Interaction between Fast and Ultra-slow Inactivation in the Voltage-gated Sodium Channel. DOES THE INACTIVATION GATE STABILIZE THE CHANNEL STRUCTURE?
J. Biol. Chem.,
September 27, 2002;
277(40):
37105 - 37115.
[Abstract]
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A. Kuzmenkin, V. Muncan, K. Jurkat-Rott, C. Hang, H. Lerche, F. Lehmann-Horn, and N. Mitrovic
Enhanced inactivation and pH sensitivity of Na+ channel mutations causing hypokalaemic periodic paralysis type II
Brain,
April 1, 2002;
125(4):
835 - 843.
[Abstract]
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J.-F. Desaphy, A. De Luca, P. Tortorella, D. De Vito, A. L. George Jr., and D. Conte Camerino
Gating of myotonic Na channel mutants defines the response to mexiletine and a potent derivative
Neurology,
November 27, 2001;
57(10):
1849 - 1857.
[Abstract]
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N. P. Davies, L. H. Eunson, M. Samuel, and M. G. Hanna
Sodium channel gene mutations in hypokalemic periodic paralysis: An uncommon cause in the UK
Neurology,
October 9, 2001;
57(7):
1323 - 1325.
[Abstract]
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J. Spampanato, A. Escayg, M. H. Meisler, and A. L. Goldin
Functional Effects of Two Voltage-Gated Sodium Channel Mutations That Cause Generalized Epilepsy with Febrile Seizures Plus Type 2
J. Neurosci.,
October 1, 2001;
21(19):
7481 - 7490.
[Abstract]
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J.-F. Desaphy, S. Pierno, C. Leoty, A. L. George Jr, A. De Luca, and D. C. Camerino
Skeletal muscle disuse induces fibre type-dependent enhancement of Na+ channel expression
Brain,
June 1, 2001;
124(6):
1100 - 1113.
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TOP
ABSTRACT
INTRODUCTION
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