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The Journal of Neuroscience, October 1, 2001, 21(19):7481-7490
Functional Effects of Two Voltage-Gated Sodium Channel Mutations
That Cause Generalized Epilepsy with Febrile Seizures Plus Type 2
Jay
Spampanato1,
Andrew
Escayg2,
Miriam H.
Meisler2, and
Alan L.
Goldin1
1 Department of Microbiology and Molecular Genetics,
University of California, Irvine, California 92697-4025, and
2 Department of Human Genetics, University of Michigan, Ann
Arbor, Michigan 48109-0618
 |
ABSTRACT |
Two mutations that cause generalized epilepsy with febrile seizures
plus (GEFS+) have been identified previously in the
SCN1A gene encoding the 
subunit of the
Nav1.1 voltage-gated sodium channel (Escayg et al., 2000
).
Both mutations change conserved residues in putative voltage-sensing S4
segments, T875M in domain II and R1648H in domain IV. Each mutation was
cloned into the orthologous rat channel rNav1.1, and the
properties of the mutant channels were determined in the absence and
presence of the 
1 subunit in Xenopus oocytes. Neither
mutation significantly altered the voltage dependence of either
activation or inactivation in the presence of the 1 subunit. The
most prominent effect of the T875M mutation was to enhance slow
inactivation in the presence of 1, with small effects on the
kinetics of recovery from inactivation and use-dependent activity of
the channel in both the presence and absence of the 1 subunit. The
most prominent effects of the R1648H mutation were to accelerate
recovery from inactivation and decrease the use dependence of channel
activity with and without the 1 subunit. The DIV mutation would
cause a phenotype of sodium channel hyperexcitability, whereas the DII
mutation would cause a phenotype of sodium channel hypoexcitability,
suggesting that either an increase or decrease in sodium channel
activity can result in seizures.
Key words:
epilepsy; sodium channels; electrophysiology; mutations; GEFS+; SCN1A
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INTRODUCTION |
Generalized epilepsy with febrile
seizures plus [GEFS+; Mendelian Inheritance in Man (MIM) #604233] was
first described by Scheffer and Berkovic (1997) as an autosomal
dominantly inherited syndrome with a complex and heterogeneous clinical
phenotype (Scheffer and Berkovic, 1997; Singh et al., 1999). This
syndrome is distinct from febrile seizures, which occur in ~5% of
all children under the age of 6 years and typically are not associated
with increased risk of epilepsy in adolescence and adulthood (Nelson
and Ellenberg, 1976; Commission on Classification and Terminology of
the International League Against Epilepsy, 1989). Scheffer and Berkovic
created the diagnosis of GEFS+ to identify families in which febrile
seizures persist beyond 6 years of age and are associated with
generalized epilepsies such as absences, myoclonic seizures, atonic
seizures, and myoclonic-astatic epilepsy (Scheffer and Berkovic, 1997;
Singh et al., 1999).
Linkage analysis identified two genetic loci for GEFS+. GEFS+ type 1 (GEFS+ 1) maps to the chromosomal region 19q13.1. A loss-of-function mutation in SCN1B, the gene encoding the voltage-gated
sodium channel 1 subunit, was identified in a family with GEFS+ 1 (Wallace et al., 1998). The mutant 1 subunit binds to but does not
modulate either neuronal Nav1.2 or skeletal
muscle Nav1.4 sodium channels (Moran and Conti,
2001). GEFS+ type 2 (GEFS+ 2) maps to the chromosomal region 2q21-q33
(Baulac et al., 1999; Moulard et al., 1999; Lopes-Cendes et al.,
2000), the location of the neuronal sodium channel gene SCN1A. Escayg et al. (2000) identified the mutations T875M
and R1648H in SCN1A, which encodes the
Nav1.1 sodium channel subunit, in two
families with GEFS+ 2. This marked the first time that a voltage-gated
sodium channel subunit had been implicated directly in the etiology
of inherited epilepsy (for review, see Steinlein and Noebels,
2000).
The sodium channel subunit is a 260 kDa transmembrane protein with
four homologous domains (DI-DIV), each with six transmembrane segments
(S1-S6). Both GEFS+ 2 mutations change conserved residues in putative
voltage-sensing S4 segments, T875M in DII and R1648H in DIV. Because
sodium channels are responsible for the initiation and propagation of
action potentials in neurons, a sodium channel mutation that makes the
neuron hyperexcitable could result in seizures (Dichter, 1991, 1994).
The effects of the R1648H mutation have been examined in two different
sodium channels, the neuronal rNav1.2 channel
(Kühn and Greeff, 1999) and the skeletal muscle hNav1.4 channel (Alekov et al., 2000), and the
effects of the mutation differed in the two studies.
To determine the effects of the two GEFS+ 2 mutations in the sodium
channel in which they are expressed normally, we cloned each
mutation into rNav1.1, the rat channel that is
orthologous to the human channel encoded by the SCN1A gene.
The human and rat orthologs are 98% identical in amino acid sequence.
The electrophysiological properties of the channels were determined in
the absence and presence of the 1 subunit by expression in
Xenopus oocytes. The DII mutation enhanced slow inactivation
and had small effects on the kinetics of recovery from inactivation and
use-dependent activity of the channel. The DIV mutation dramatically
accelerated recovery from inactivation and decreased the use dependence
of channel activity.
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MATERIALS AND METHODS |
Site-directed mutagenesis. pNaRat1 encodes a
functional rNav1.1 cDNA clone positioned between
the 5' and 3' noncoding regions of the Xenopus -globin
gene, downstream of a T7 RNA polymerase promoter and upstream of a
poly(A+) tail region (Smith and Goldin,
1998). A 1.9 kb EcoRI-AccI fragment containing
the domain II S4 segment and a 1.7 kb
BstEII-NotI fragment containing the domain IV S4
segment were subcloned into pBSTA and pGEM18B,
respectively. Mutagenesis was conducted with the QuikChange
Site-Directed Mutagenesis kit (Stratagene, La Jolla, CA) by using
the following oligonucleotide primers: T875M (sense), GCAAAGTCCTGGCCCATGCTGAACATGCTCATTAAG; T875M (antisense),
CTTAATGAGCATGTTCAGCATGGGCCAGGACTTTGC; R1648H (sense),
GGATTGGACGAATCCTACACCTGATCAAAGGCGCC; and R1648H (antisense), GGCGCCTTTGATCAGGTGTAGGATTCGTCCAATCC.
For each mutant, DH5 bacteria were transformed via electroporation
with DpnI-treated DNA. The electroporation was performed in
parallel with chemical transformation of XL-1 Blue bacteria, using the
procedures described in the kit. The products of both transformations
were plated on LB-ampicillin agar plates and incubated at 37°C for
>16 hr. Clones were selected for each mutant, and the desired
mutations were verified by the ThermoSequenase Radiolabeled Terminator Cycle Sequencing Kit (United States Biochemicals,
Cleveland, OH) and the following primers: T875M (sequencing primer),
GACACTCAGCCTGGTAGAAC, and R1648H (sequencing primer), TCCTCTCCATTGTAGGAATG.
At least one clone of the four clones sequenced for each mutant was
found to contain the desired mutation with no extraneous mutations. A
1.6 kb EcoRI-MfeI fragment from the domain II S4 subclone containing the mutation T875M and a 1.7 kb
BstEII-NotI fragment from the domain IV S4
subclone containing the mutation R1648H were cloned into the original
full-length pNaRat1 plasmid. The new plasmids were termed DII, which
contains the T875M mutation, and DIV, which contains the R1648H mutation.
Expression and electrophysiology. RNA was transcribed
in vitro from NotI-linearized DNA templates with
the T7 mMessage mMachine kit (Ambion, Austin, TX). The quality of the
transcribed mRNA was confirmed by glyoxal gel analysis. Stage V oocytes
were removed from adult female Xenopus laevis
frogs and prepared as previously described (Goldin, 1991). Oocytes were
incubated in ND-96 media, which consisted of (in
mM) 96 NaCl, 2 KCl, 1.8 CaCl2, 1 MgCl2, and 5 HEPES, pH 7.5, supplemented with 0.1 mg/ml gentamycin, 0.55 mg/ml
pyruvate, and 0.5 mM theophylline. RNA encoding
rNav1.1, DII, and DIV channels was injected at
~20 ng/oocyte to obtain current amplitudes between 0.8 and 6 µA.
When the channels were coexpressed with the 1 subunit, a 1:10 molar
ratio of -to-1 RNA was injected. Oocytes were incubated at 20°C
for >40 hr in ND-96 before voltage clamping.
Sodium currents were recorded by using either the two-electrode voltage
clamp (TEVC) at room temperature or the cut-open oocyte CA-1 High
Performance Oocyte Voltage Clamp (Dagan Instruments, Minneapolis, MN)
at 20°C, with a DigiData 1321A interface (Axon Instruments, Foster
City, CA) and pClamp 8.0 software (Axon Instruments) as previously
described (Patton and Goldin, 1991; Kontis et al., 1997). On the TEVC
the currents were recorded in ND-96 without supplements in the absence
and presence of 400 nM tetrodotoxin (TTX). Capacitive
transients were eliminated by subtraction of TTX-recorded currents.
Slow-gated properties were analyzed via a baseline subtraction method
in which the average current amplitude recorded during the last 1 msec
of a given test pulse was subtracted from the peak current amplitude of
that same test pulse before normalization.
The voltage dependence of activation was analyzed via a step protocol
in which oocytes were depolarized from a holding potential of 
100 mV
to a range of potentials from 95 to +50 mV in 5 mV increments. Peak
currents were normalized to the maximum peak current and plotted
against voltage. To calculate a reversal potential, we fit the
resulting I-V curve of each data set individually with the
equation:
![I=[1+<UP>exp</UP>(<UP>−</UP>0.03937·z·(V−V<SUB>1/2</SUB>))]<SUP><UP>−</UP>1</SUP>·g·(V−V<SUB><UP>r</UP></SUB>),](/content/vol21/issue19/fulltext/7481/img001.gif)
|
(1)
|
where I is the current amplitude, z is the
apparent gating charge, V is the potential of the given
pulse, V1/2 is the half-maximal voltage,
g is a factor related to the number of open channels during
the given pulse, and Vr is the
reversal potential. Then conductance was calculated directly by using
the equation:
|
(2)
|
where G is conductance and I, V, and
Vr are as described above. The
conductance values were fit with the two-state Boltzmann equation:
![G=1/(1+<UP>exp</UP>[<UP>−</UP>0.03937·z·(V−V<SUB>1/2</SUB>)]),](/content/vol21/issue19/fulltext/7481/img003.gif)
|
(3)
|
where z is the apparent gating charge, V
is the potential of the given pulse, and
V1/2 is the potential for half-maximal activation.
The voltage dependence of steady-state inactivation was determined via
a two-step protocol in which a conditioning pulse was applied from a holding potential of 100 mV to a range of potentials from 100 to +15 mV in 5 mV increments for 100 msec, followed immediately by a test pulse to 5 mV. The peak current amplitudes during the subsequent test pulses were normalized to the peak current
amplitude during the first test pulse, plotted against the potential of
the conditioning pulse, and fit with the two-state Boltzmann equation:
![I=1/(1+<UP>exp</UP>[(V−V<SUB>1/2</SUB>)/a]),](/content/vol21/issue19/fulltext/7481/img004.gif)
|
(4)
|
where I is equal to the test pulse current amplitude,
V is the potential of the conditioning pulse,
V1/2 is the voltage for half-maximal
inactivation, and a is the slope factor.
Recovery from inactivation was analyzed with three separate two-pulse
protocols. Each protocol began with a conditioning depolarization from
a holding potential of 100 to 5 mV for 50 msec, which inactivated >95% of the channels. This was followed by a decreasing recovery time
interval at 100 mV and a test depolarization to 5 mV. The three
protocols differed only in the maximum length of recovery time and the
time interval by which that recovery period decreased: 25 msec maximum
and 1 msec decrements in the short protocol, 200 msec maximum and 5 msec decrements in the intermediate protocol, and 3000 msec maximum and
100 msec decrements in the long protocol. Fractional recovery was
calculated by dividing the maximum current amplitude during the test
pulse by the maximum current amplitude of the corresponding
conditioning pulse. The recovery data were fit with either the double
exponential equation:
![I=1−[A<SUB>1</SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB>1</SUB>)+A<SUB>2</SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB>2</SUB>)],](/content/vol21/issue19/fulltext/7481/img005.gif)
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(5)
|
or the triple exponential equation:
![I=1−[A<SUB>1</SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB>1</SUB>)+A<SUB>2</SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB>2</SUB>)+A<SUB>3</SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB>3</SUB>)],](/content/vol21/issue19/fulltext/7481/img006.gif)
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(6)
|
where A1,
A2, and
A3 are the relative percentages of
current that recovered with the time constants
1, 2, and
3, and t is the recovery time.
Use dependence was analyzed at frequencies of 10, 20, and 39 Hz by
using 17.5 msec depolarizations to 10 mV from a holding potential of
100 mV. The protocols were performed for 2 sec at 10 Hz, 2.5 sec at
20 Hz, and 2.56 sec at 39 Hz, which was long enough for the current to
have reached an equilibrium value in each case. Peak current amplitudes
were normalized to the peak current amplitude during the first
depolarization and plotted against pulse number.
The kinetics of fast inactivation were analyzed by using the cut-open
oocyte voltage clamp, with the bath solution maintained at 20°C with
an HCC-100A Temperature Controller (Dagan). The external solution
consisted of (in mM) 120 sodium methanesulfonate, 10 HEPES,
and 1.8 calcium methanesulfonate, pH 7.5. The internal solution
consisted of (in mM) 88 K2SO4, 10 EGTA, 10 HEPES,
and 10 Na2SO4, pH 7.5. P/4
subtraction was used to eliminate capacitive transients and leak
currents. Currents were elicited via a step protocol in which oocytes
were depolarized from a holding potential of 100 mV to a range of
potentials from 95 to +50 mV in 5 mV increments. Inactivation time
constants were determined by the Chebyshev method to fit each trace
with either the single exponential equation:
![I=A<SUB><UP>Slow</UP></SUB>·<UP>exp</UP>[<UP>−</UP>(t−K)/&tgr;<SUB><UP>Slow</UP></SUB>]+C,](/content/vol21/issue19/fulltext/7481/img007.gif)
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(7)
|
or the double exponential equation:
![I=A<SUB><UP>Fast</UP></SUB>·<UP>exp</UP>[<UP>−</UP>(t−K)/&tgr;<SUB><UP>Fast</UP></SUB>]+A<SUB><UP>Slow</UP></SUB>·<UP>exp</UP>[<UP>−</UP>(t−K)/&tgr;<SUB><UP>Slow</UP></SUB>]+C,](/content/vol21/issue19/fulltext/7481/img008.gif)
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(8)
|
where I is the current,
AFast and
ASlow are the relative proportions of
current inactivating with the time constants
Fast and
Slow, K is the time shift,
and C is the steady-state asymptote. The time shift was
selected manually as the point at which the macroscopic current began
to inactivate exponentially.
The voltage dependence of steady-state slow inactivation was determined
via a modified two-step protocol in which a conditioning pulse was
applied from a holding potential of 120 mV to a specified range of
potentials between 120 and 10 mV for a period of 60 sec. The
conditioning pulse was followed immediately by a hyperpolarization to
120 mV for 20 msec to allow for recovery from fast inactivation and a
test pulse to 5 mV. The peak current amplitudes during the subsequent
test pulses were normalized to the peak current amplitude during the
first test pulse, plotted against the potential of the conditioning
pulse, and fit with the two-state Boltzmann equation described earlier:
![I=1/(1+<UP>exp</UP>[(V−V<SUB>1/2</SUB>)/a]).](/content/vol21/issue19/fulltext/7481/img009.gif)
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(9)
|
The rate of entry into the slow-inactivated state was analyzed
by using a two-step protocol with a variable length conditioning pulse,
followed by a test pulse. The conditioning potential of 45 mV was
applied from a holding potential of 120 mV for a variable length of
time beginning with 0 sec and ending with 60 sec in 5 sec increments.
The conditioning pulse was followed immediately by a hyperpolarization
to 120 mV for 20 msec to allow for recovery from fast inactivation
and a test pulse to 5 mV. The peak current amplitudes during the
subsequent test pulses were normalized to the peak current amplitude
during the first test pulse, plotted against the period of the
conditioning pulse, and fit with the double exponential equation:
|
(10)
|
where I is the current,
AFast and
ASlow are the relative proportions of
current inactivating with the time constants
Fast and
Slow, and t is the period of
the conditioning pulse. The rate of entry into the slow-inactivated
state also was analyzed with a 10 mV conditioning pulse over a period
of 28.5 sec. At this potential, however, the conditioning pulse length
was varied in a nonlinear manner to collect more data during the first
10 sec, because >80% of the current inactivated in <10 sec.
Recovery from slow inactivation was analyzed with two separate
two-pulse protocols. Each protocol began with a conditioning depolarization from a holding potential of 120 to 5 mV for 60 sec,
which inactivated >95% of the channels. This was followed by a
decreasing recovery time interval at 120 mV, an additional 20 msec at
120 mV, and a test depolarization to 5 mV. Like the fast
inactivation recovery protocols, these protocols differed only in the
maximum length of recovery time and the time interval by which that
recovery period decreased. The short protocol used a 14 sec maximum and
2 sec decrements and the long protocol used a 60 sec maximum and 10 sec
decrements. Fractional recovery was calculated by dividing the maximum
current amplitude during the test pulse by the average maximum current
amplitude during five single-step depolarizations to 5 mV recorded
before each recovery protocol and was plotted against the length of the
recovery interval. The recovery data were fit with a double exponential
equation:
![I=1−[A<SUB><UP>Fast</UP></SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB><UP>Fast</UP></SUB>)+A<SUB><UP>Slow</UP></SUB>·<UP>exp</UP>(<UP>−</UP>t/&tgr;<SUB><UP>Slow</UP></SUB>)],](/content/vol21/issue19/fulltext/7481/img011.gif)
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(11)
|
where I is the current,
AFast and
ASlow are the relative percentages of
current that recovered with the time constants
Fast and
Slow, and t is the recovery time.
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RESULTS |
The GEFS+ 2 mutations do not affect the voltage dependence of
activation or inactivation significantly
Escayg et al. (2000) previously identified two mutations in the
SCN1A gene encoding the Nav1.1 sodium
channel that cause GEFS+ 2. The two mutations are a substitution of
methionine for threonine 875 (T875M) in domain II and a substitution of
histidine for arginine 1648 (R1648H) in domain IV. To determine how
these mutations affect sodium channel function, we constructed each
mutation in the orthologous rat channel rNav1.1.
Then the properties of the mutant channels were compared with those of
the wild-type channel expressed in Xenopus oocytes by using
the cut-open oocyte voltage clamp and the two-electrode voltage clamp.
The channels were compared in the absence and presence of the 1
subunit. Figure 1 shows sample cut-open
oocyte voltage-clamp recordings of the currents from the wild-type
(rNav1.1) and the two mutant channels during
depolarizations between 50 and +50 mV in 10 mV intervals. For all
three samples the injection of ~20 ng of RNA resulted in current
amplitudes between 1 and 5 µA, suggesting that the mutations had no
significant effects on the translation or processing of the channel
proteins. As can be seen, there were no dramatic differences in channel kinetics between the mutant and wild-type channels in either the absence or presence of the 1 subunit.
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Figure 1.
Sample sodium channel currents from wild-type
rNav1.1 and GEFS+ 2 mutant channels. Currents were recorded
for the wild-type rNav1.1, DII (T875M), and DIV (R1648H)
mutants expressed as subunits alone and as + 1 subunits.
Mutant and wild-type channels were expressed in Xenopus
oocytes, and currents were recorded at 20°C by using the cut-open
oocyte voltage clamp, as described in Materials and Methods. Membrane
depolarizations from a holding potential of 100 mV to a range of
potentials from 50 to +50 mV in 10 mV increments are shown.
Calibration: 2 msec, 200 nA.
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Because both mutations are located in S4 regions that function as
voltage sensors for the channel, it was possible that one or both
mutations might alter the voltage dependence of either activation or
inactivation. A shift in the voltage dependence of activation of the
Nav1.6 neuronal sodium channel has been
demonstrated previously to result from a mutation that causes ataxia in
medjo mice (Smith and Goldin, 1999). We
therefore compared the voltage dependence of activation and
inactivation of the mutant and wild-type channels in the absence and
presence of the 1 subunit. The data are shown in Figure
2, with the parameters of the Boltzmann
fits displayed in Table 1.
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Figure 2.
Voltage dependence of activation and
steady-state inactivation for wild-type rNav1.1 and GEFS+ 2 mutant channels. The voltage dependencies of activation
(circles) and inactivation (diamonds)
were determined for the wild-type rNav1.1 (white
symbols), DII (gray symbols), and DIV
(black symbols) mutants expressed as subunits alone
(A) and as + 1 subunits
(B). Sodium currents were recorded from a holding
potential of 100 mV by depolarizations to a range of potentials from
95 to +50 mV in 5 mV increments. Conductance values were calculated
by dividing the peak current amplitude by the driving force at each
potential and normalizing to the maximum conductance, as described in
Materials and Methods. The values shown are averages; the error bars
indicate SD. The data were fit with a two-state Boltzmann equation, and
the parameters of the fits are shown in Table 1. The voltage dependence
of inactivation was determined by using a two-step protocol in which a
conditioning pulse was applied from a holding potential of 100
mV, consisting of 100 msec depolarizations to a range of potentials
from 100 to +15 mV in 5 mV increments, followed by a test pulse to
5 mV. The peak current amplitude during each test pulse was
normalized to the current amplitude of the first test pulse and plotted
as a function of the conditioning pulse potential. The values shown are
averages; the error bars indicate SD. The data were fit with a
two-state Boltzmann equation, and the parameters of the fits are shown
in Table 1.
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The voltage dependence of activation for the DII (T875M, gray
circles) and the DIV (R1648H, black circles) mutants
was not significantly different from that of the wild-type channel
(rNav1.1, white circles) when
expressed as either the subunit alone or as + 1 subunits
(Fig. 2). There were no significant differences in either the
V1/2 or slope values for any of the
channels (Table 1). In addition, the presence of the 1 subunit did
not alter the voltage dependence of activation significantly, in
agreement with previously published data that demonstrated no
significant effects of the 1 subunit on the voltage dependence of
activation of rNav1.1 or
rNav1.2 channels (Smith and Goldin, 1998).
The voltage dependence of inactivation was not significantly different
between the DII (T875M, gray diamonds) and wild-type (rNav1.1, white diamonds) channels
when the subunits were expressed alone (Fig. 2A,
Table 1). However, the DIV (R1648H, black diamonds) mutant
subunit displayed a shift of ~8 mV in the hyperpolarized direction. When the channels were coexpressed with the 1 subunit, there were no significant differences among the DII (T875M, gray diamonds), DIV (R1648H, black diamonds), and wild-type
(rNav1.1, white diamonds) channels.
Coexpression of 1 shifted the V1/2 of
inactivation for both the wild-type and the DII channels in the
negative direction but did not alter the
V1/2 of inactivation for the DIV mutant
channel significantly. The negative shift in the
V1/2 of inactivation is similar but less
pronounced than previously observed (Smith and Goldin, 1998). In
summary, neither of the GEFS+ 2 mutations significantly altered the
voltage dependence of activation or inactivation.
The DIV mutant alters the kinetics of inactivation
Because neither mutation markedly changed the voltage
dependence of channel gating, it was possible that the neuronal
phenotype in GEFS+ 2 could result from alterations in the kinetics of
inactivation. Based on the sample currents shown in Figure 1, the DIV
mutant appeared to have small effects on inactivation kinetics when
expressed as the subunit alone. To quantify these effects, we fit
current traces similar to those shown in Figure 1 with either a single or a double exponential equation, as described in Materials and Methods. In the absence of the 1 subunit, wild-type
rNav1.1 was best fit with a single exponential
equation for potentials from 35 to +10 mV, resulting in a single
slow (Fig.
3A, white
triangles). The DII (T875M, gray triangles) and DIV
(R1648H, black triangles) channels were best fit with a
single exponential for potentials from 30 to +10 mV and 35 to +10
mV, respectively. A double exponential equation resulting in a
slow (triangles) and a
fast (circles) was used to fit
wild-type rNav1.1 and both GEFS+ 2 mutants for potentials positive to +10 mV. When expressed as the subunit alone,
the DII mutant displayed kinetics of inactivation similar to that of
the wild-type channel. In addition, the percentage of the fast
component was similar for the DII mutant and
rNav1.1 wild-type channels. The DIV mutant
displayed a slower fast than wild-type
rNav1.1 for potentials positive to +10 mV and a
faster slow for potentials between 10 and
+10 mV. The DIV mutant also inactivated a larger percentage of current
than did wild-type rNav1.1 with
fast for the potentials of +15 and +20 mV.
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Figure 3.
Kinetics of fast inactivation of
wild-type rNav1.1 and GEFS+ 2 mutant channels. Sodium
currents were recorded from oocytes expressing wild-type
rNav1.1 (white symbols and
bars), DII (gray symbols and
bars), and DIV (black symbols and
bars) channels, as described in the legend to Figure 1.
Current traces were fit with either a single or a double exponential
equation, as described in Materials and Methods, and time constants for
the fast (fast, circles) and slow
(slow, triangles) components of
fast inactivation are plotted on a logarithmic scale in the
top panels for subunits alone
(A) and + 1 subunits
(B). The bottom panels indicate
the fraction of current inactivating with fast. In all
cases the sum of the components is one. The values shown are averages;
the error bars indicate SD. Sample sizes were rNav1.1 (4), DII (5), DIV (4), rNav1.1 + 1 (4), DII
+ 1 (5), DIV + 1 (7).
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Coexpression of both mutants with the 1 subunit resulted in
fast values that were similar to those of
wild-type rNav1.1 at all potentials at which
fast was detected (Fig. 3B). For
each mutant and wild-type rNav1.1,
fast was detected first at 25 mV. The DIV
mutant again displayed slow values that were
slightly faster than those of wild-type rNav1.1
at potentials between 5 and +10 mV, with a similar percentage of
current inactivating with fast at all
potentials. The DII mutant displayed similar kinetics to that of
wild-type rNav1.1, with a similar ratio of current inactivating with fast at all potentials.
The DIV mutant displays a more rapid recovery from inactivation
than does wild-type rNav1.1
Inactivated channels experience a latency period during which they
recover from the inactivated state to the closed state. This transition
to the closed state must occur before the channels are capable of
opening in response to subsequent membrane depolarizations. A mutation
that alters this property would result in channels that are capable of
either more rapid recovery, resulting in hyperexcitability, or less
rapid recovery, resulting in hypoexcitability. Neuronal hyperexcitability is known to be the cause of some seizure activity (Dichter, 1991, 1994). Separate disease-causing mutations in both Nav1.4 and Nav1.5 have been
shown to result in hyperexcitability via the mechanism of more rapid
recovery from inactivation (Hayward et al., 1996; Chen et al., 1998).
We therefore analyzed the effects of each GEFS+ 2 mutation on recovery
from inactivation in comparison to wild-type
rNav1.1 in the presence and absence of the 1
subunit, as described in Materials and Methods (Fig.
4, Table 1).
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Figure 4.
Recovery from inactivation for wild-type
rNav1.1 and GEFS+ 2 mutant channels. Recovery from
inactivation was determined by using three separate two-pulse protocols
for wild-type rNav1.1 (white symbols), DII
(gray symbols), and DIV (black
symbols) channels. Each protocol was performed with a holding
potential of 100 mV and consisted of a conditioning depolarization to
5 mV for 50 msec (which inactivated >95% of the channels), a
decreasing recovery time interval at 100 mV, and a test
depolarization to 5 mV. The three protocols differed only in the
maximum length of recovery time and the time interval by which that
recovery period decreased: 25 msec maximum and 1 msec decrements in the
early protocol, 200 msec maximum and 5 msec decrements in the
intermediate protocol, and 3000 msec maximum and 100 msec decrements in
the late protocol. Fractional recovery was calculated by dividing the
maximum current amplitude of the test pulse by the maximum current
amplitude of the corresponding conditioning pulse. Fractional recovery
is plotted on a log scale as a function of time for subunits alone
(A) and + 1 subunits
(B). The values shown are averages; the error
bars indicate SD.
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In the absence of 1 the DII mutant (Fig. 4A,
gray circles) displayed slower recovery than wild-type
rNav1.1 (white circles) in the
early-to-middle time period, whereas the DIV mutant (black circles) displayed faster recovery at all times. The time for ~50% recovery was 45 msec for the DII mutant, 12 msec for the wild-type rNav1.1, and <3 msec for the DIV
mutant. The DIV mutant recovered 95% of the total current in <18
msec, >20 times faster than wild-type rNav1.1
and DII mutant channels, which required between 400 and 500 msec for
comparable levels of recovery. Data for the rapidly recovering DIV
mutant were best fit with a double exponential equation, in contrast to
the DII mutant and wild-type rNav1.1 channels,
both of which were fit with triple exponential equations (Table 1).
Coexpression of the 1 subunit with wild-type
rNav1.1 has been shown previously to result in a
more rapid recovery from inactivation (Smith and Goldin, 1998).
Coexpression of the 1 subunit with the GEFS+ 2 mutants also resulted
in more rapid recovery from inactivation when compared with subunit
channels (Fig. 4B). In the presence of the 1
subunit, recovery from inactivation was similar for the DII mutant
(gray circles) and the wild-type rNav1.1 (white circles). The DIV
mutant (black circles) demonstrated more rapid recovery
from inactivation than either wild-type
rNav1.1 or DII mutant channels even when
coexpressed with the 1 subunit. The DIV mutant recovered 95 ± 1% of the total current at 8 msec, whereas the DII mutant and
wild-type rNav1.1 each required ~100 msec for
comparable recovery. In the presence of the 1 subunit, recovery from
inactivation for all three channels was best fit with a double
exponential equation (Table 1).
The DIV mutant achieved a more rapid recovery by two different effects.
When expressed as an subunit alone, a larger percentage of the
total current recovered with the shortest time constant (1) and the remaining portion of the total
current recovered with a more rapid second time constant
(2) as compared with both the wild-type
rNav1.1 channel and the DII mutant (Table 1).
When coexpressed with the 1 subunit, however, the DIV mutant
recovered with a more rapid 1 as compared with
the wild-type rNav1.1 channel, although the
percentage of current recovering with that time constant was similar
for both channels. The second time constant of recovery (2) was also faster for the DIV mutant as
compared with the wild-type rNav1.1 and DII
mutant channels.
Both GEFS+ 2 mutants display altered frequency dependence
as compared with wild-type rNav1.1
An alternative way to evaluate recovery from inactivation is to
examine the use or frequency dependence of sodium current amplitudes
during a series of depolarizations. If there is insufficient time for
complete recovery between each depolarization, then the magnitude of
the current will decrease with successive depolarizations. Therefore,
we examined the frequency dependence of both GEFS+ 2 mutants in the
absence and presence of the 1 subunit at 10, 20, and 39 Hz, as
described in Materials and Methods (Fig.
5).
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Figure 5.
Frequency dependence of wild-type
rNav1.1 and GEFS+ 2 mutant channels. Use dependence was
analyzed at 10, 20, and 39 Hz for wild-type rNav1.1
(white symbols), DII (gray
symbols), and DIV (black symbols) channels.
Currents were elicited at each frequency by using 17.5 msec
depolarizations to 10 mV from a holding potential of 100 mV. Each
protocol was performed until an equilibrium current had been reached: 2 sec at 10 Hz, 2.5 sec at 20 Hz, and 2.56 sec at 39 Hz. Peak current
amplitudes were normalized to the initial peak current amplitude and
plotted against pulse number for subunits alone (A,
circles) and + 1 subunits (B,
diamonds). The values shown are averages; the error bars
indicate SD. Sample sizes were rNav1.1 (5), DII (5), DIV (5), rNav1.1 + 1 (3), DII + 1
(5), DIV + 1 (5).
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When expressed as subunits alone (Fig. 5A), the DIV
mutant (black circles) maintained the highest level of
current, with the DII mutant (gray circles) next, and
the wild-type rNav1.1 channel (white
circles) least at all frequencies. At 10 Hz the DIV mutant showed
little-to-no use-dependent decrease in current and was capable of
maintaining 94 ± 1% of the maximum current for up to 2 sec (20 depolarizations). The DII mutant showed some use dependence at 10 Hz,
reaching an equilibrium level of 74 ± 3% of the maximum current,
which was still significantly higher than the wild-type
rNav1.1 channel, which reached equilibrium at
65 ± 4% of the maximum current level. At 20 Hz the DIV mutant showed some use dependence during the 2.5 sec protocol, reaching an
equilibrium current level of 86 ± 1% of the initial peak current amplitude, as compared with 55 ± 4% for the DII mutant and only 45 ± 3% for the wild-type rNav1.1. At 39 Hz the DIV mutant demonstrated significant use dependence, with an
equilibrium current of 58 ± 1% of the initial peak amplitude,
but this level was still more than twofold higher than the equilibrium
observed for both the DII mutant and wild-type
rNav1.1 channel, which reached levels of 27 ± 4 and 22 ± 1%, respectively. Although the DII mutant reached an equilibrium current level similar to that of the wild-type rNav1.1 after 2.5 sec at 39 Hz, the mutant
channel maintained a slightly higher level of current during the first second.
When the channels were coexpressed with the 1 subunit, both GEFS+ 2 mutants and the wild-type rNav1.1 channels
displayed less use dependence as compared with subunits alone (Fig.
5B). At 10 Hz the DIV mutant (black diamonds),
the DII mutant (gray diamonds), and wild-type
rNav1.1 (white diamonds) showed no
significant use dependence, with equilibrium current levels of 97 ± 2, 95 ± 1, and 97 ± 1% of the initial peak current
amplitudes, respectively. At 20 Hz both the DIV mutant and wild-type
rNav1.1 displayed similar equilibrium current
levels, with the DIV mutant maintaining 94 ± 2% and wild-type
rNav1.1 maintaining 92 ± 1% of the initial current amplitudes. The DII mutant demonstrated more pronounced use
dependence, with an equilibrium level of 84 ± 3% of the initial peak current amplitude. At the highest frequency that was tested, 39 Hz, the three channels displayed significantly different levels of use
dependence. The DIV mutant displayed the least use dependence, maintaining 73 ± 5% of the initial peak amplitude as compared with 64 ± 1% for the wild-type rNav1.1 and
46 ± 4% for the DII mutant.
These results show that the DIV mutant is capable of carrying a larger
amount of current than the wild-type rNav1.1
channel at high frequencies of depolarization when expressed as either the subunit alone or as + 1 subunits. This property is
consistent with a phenotype of hyperexcitability. The DII mutant is
capable of carrying a larger amount of current than the wild-type
rNav1.1 channel at low frequencies of
depolarization and at early times during high frequencies of
depolarization when expressed as the subunit alone, but this effect
is reversed when the 1 subunit is coexpressed.
The DII mutant displays enhanced slow inactivation as compared with
wild-type rNav1.1
In addition to the fast-gated properties examined above, sodium
channels undergo slow-gated transitions that occur on the time scale of
seconds to minutes. Several disease-causing mutations in the skeletal
muscle sodium channel Nav1.4 have been reported to disrupt the voltage dependence of slow inactivation, disrupt entry
into the slow-inactivated state, or enhance recovery from slow
inactivation (for review, see Cannon, 2000). Because the fast-gated
properties of the DII mutant were very similar to those of the
wild-type channel, it seemed likely that this mutation might have more
of an effect on slow inactivation. Therefore, we examined the voltage
dependence of slow inactivation, the rate of entry into the
slow-inactivated state, and recovery from slow inactivation of both
GEFS+ 2 mutants in comparison to wild-type rNav1.1 in the presence of the 1 subunit, as
described in Materials and Methods (Fig.
6, Table
2).
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Figure 6.
Slow-gated properties of wild-type
rNav1.1 and GEFS+ 2 mutant channels. The slow-gated
properties were determined for the wild-type rNav1.1
(white circles), DII (gray
circles), and DIV (black circles) mutants
expressed as + 1 subunits. The voltage dependence of slow
inactivation (A) was analyzed by using a two-step
protocol consisting of 60 sec depolarizations from a holding potential
of 120 mV to a range of potentials between 120 and 10 mV,
followed by a hyperpolarization to 120 mV for 20 msec to allow for
recovery from fast inactivation and a test pulse to 5 mV. The data
were fit with a two-state Boltzmann equation, as described in Materials
and Methods, and the parameters of the fits are shown in Table 2. The
recovery from slow inactivation (B) was analyzed
by using two separate two-pulse protocols consisting of a 60 sec
depolarization to 5 mV from a holding potential of 120 mV, followed
by a variable recovery time at 120 mV, a hyperpolarization to 120
mV to allow for recovery from fast inactivation, and a test
depolarization to 5 mV. The data were fit with a double exponential
equation, as described in Materials and Methods, and the parameters of
the fits are shown in Table 2. The rate of entry into the
slow-inactivated state (C, D) was
analyzed by using a two-step protocol consisting of a variable length
conditioning pulse at either 45 or 10 mV from a holding potential
of 120 mV, followed by a hyperpolarization to 120 mV to allow
for recovery from fast inactivation and a test depolarization to 5
mV. The data were fit with a double exponential decay, as described in
Materials and Methods, and the parameters of the fits are shown in
Table 2. For each graph the values shown are averages; the error bars
indicate SD.
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The voltage dependence of slow inactivation of the DII mutant (Fig.
6A, gray circles) was shifted 10 mV in the
negative direction when compared with both the DIV mutant (black
circles) and wild-type rNav1.1 (white
circles). The DII mutant also displayed a significantly steeper
slope (Table 2), thus inactivating over a smaller voltage range than
wild-type rNav1.1. Interestingly, the DIV mutant
also displayed a significantly steeper slope that was similar to that of the DII mutant, but the V1/2 of slow
inactivation was similar to that of wild-type
rNav1.1
Both GEFS+ 2 mutants and wild-type rNav1.1
displayed biphasic recovery from the slow-inactivated state (Fig.
6B, Table 2). The DIV mutant (black
circles), which was shown to recover from fast inactivation more
rapidly than wild-type rNav1.1, recovered from
slow inactivation in a manner that was very similar to that of
wild-type rNav1.1 (white circles).
However, the DII mutant (gray circles) displayed a
significantly slower slow component (slow) of
recovery than that of wild-type rNav1.1 (Table
2). This effect resulted in less complete recovery by 60 sec at 120 mV, reaching only 87 ± 5% recovery for the DII mutant as
compared with 97 ± 1% for the wild-type channels.
The rate of entry into the slow-inactivated state for both GEFS+ 2 mutants and wild-type rNav1.1 also followed a
biphasic time course (Fig. 6C,D, Table 2). Entry into slow
inactivation was tested at two potentials, 45 and 10 mV. At 45 mV
both the DIV mutant (black circles) and wild-type
rNav1.1 (white circles) displayed
significantly less slow inactivation as compared with the DII mutant
(gray circles). The DII mutant resulted in a more complete slow inactivation by inactivating a significantly larger percentage of current with the fast time component
(Fast) as compared with both wild-type
rNav1.1 and the DIV mutant. The magnitude of the
fast time constant was comparable for all three channels. The DII
mutant inactivated the remainder of the current with a significantly
more rapid slow time constant (Slow) as
compared with both the DIV mutant and wild-type
rNav1.1 (Table 2). The DII mutant fully
inactivated in 60 sec, whereas the DIV mutant and wild-type
rNav1.1 channels did not, consistent with the
results examining the voltage dependence of slow inactivation (Fig.
6A).
At 10 mV all of the channels inactivated more rapidly than at 45
mV, which was expected. However, the DII mutant channel still
demonstrated more rapid entry into the slow-inactivated state. At 10
mV the DII mutant displayed a larger percentage of current inactivating
with a faster Fast as compared with wild-type rNav1.1 (Table 2). The DIV mutant also displayed
a faster Fast at 10 mV, but it was otherwise
similar to wild-type rNav1.1. All three channels
inactivated completely within 30 sec, consistent with the results
examining the voltage dependence of slow inactivation (Fig.
6A).
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DISCUSSION |
We have shown that two sodium channel mutations that cause GEFS+ 2 alter the function of the channels. Each mutation was cloned into the
orthologous rat sodium channel (rNav1.1) and
expressed in Xenopus oocytes. Mutant channels were expressed
at comparable levels to the wild-type rNav1.1
channel, indicating that the mutations had no significant effects on
the translation or processing of the channel proteins.
The two mutations had different effects on the properties of the sodium
channel. The DIV mutation accelerated recovery from inactivation (see
Fig. 4) and decreased use dependence (see Fig. 5). The mutant channels
displayed a much more rapid two-component recovery from inactivation
when expressed both as the subunit alone and when coexpressed with
the 1 subunit. This more rapid recovery would make the mutant
channels available to initiate and propagate action potentials at a
higher frequency during a sustained depolarization. The ability of the
DIV mutant channels to sustain a rapid train of action potential firing
was demonstrated by their reduced frequency dependence. The ability of
the mutant channel to carry more current than the wild-type channel at
each frequency most likely results from the more rapid recovery of the
mutant channel from inactivation. At 10 Hz the DIV mutant is capable of
maintaining ~100% of the initial current for up to 2 sec in both the
absence and presence of the 1 subunit. At this frequency the
start-to-start interval of each successive depolarization is 100 msec,
a time span that is sufficient for the DIV mutant channels to recover
fully from inactivation. This correlation between frequency dependence
and recovery time is also true for the higher frequencies of 20 and 39 Hz. In neurons firing action potentials in rapid succession, the DIV
mutant channels that open and inactivate during a given action
potential would recover from inactivation and be ready to participate
in the firing of a second action potential 2- to 10-fold faster than
the wild-type channels. These effects are consistent with the GEFS+ 2 phenotype of hyperexcitability.
The DII mutation had minor effects on fast-gated properties, but it
greatly enhanced slow inactivation (see Fig. 6). There were two effects
on fast-gated properties. First, recovery from inactivation was slower
for the DII mutant channels as compared with the wild-type channels
when the subunits were expressed alone, but not when the channels
were coexpressed with the 1 subunit. Second, the DII mutant channels
showed less use dependence than the wild-type channels when the subunits were expressed alone but more use dependence when the 1
subunit was present. The increased use dependence probably results from
enhanced entry into the slow-inactivated state, because the DII mutant
and wild-type channels recovered from fast inactivation with similar
kinetics. At 39 Hz with 17.5 msec depolarizations to 10 mV, the
channels spend most of the 2.56 sec protocol at 10 mV. A single
depolarization to 10 mV for 1.5 sec reduced the available current to
63 ± 4% for the wild-type channels and to 34 ± 5% for the
DII mutant channels (see Fig. 6D). After
approximately the same time in the use dependence protocol (58 depolarizations), the current amplitude was reduced to 67 ± 1%
for wild-type rNav1.1 and to 53 ± 4% for
the DII mutant (see Fig. 5B). Therefore, the decrease in
current during a series of rapid depolarizations is comparable to the
decrease during a single long depolarization. This correlation between
enhanced slow inactivation and increased use dependence has been
reported by Struyk et al. (2000) for the hypokalemic periodic paralysis mutation R669H in DIIS4 of hNav1.4 and by Wang
and Wang (1997) for the N434A mutation in DIS6 of
rNav1.4. Each of these mutations shifted the
voltage dependence of slow inactivation in the hyperpolarized direction, accelerated entry into the slow-inactivated state, slowed
recovery from slow inactivation, and increased use dependence.
Enhanced slow inactivation is more consistent with a phenotype of
hypoexcitability rather than hyperexcitability, which is contrary to
what would be expected for a mutation that causes epilepsy. An extended
depolarization to a potential that elicits slow inactivation would
result in more extensive slow inactivation of the DII mutant channels,
which would decrease the ability of the neuron to fire action
potentials at high frequency. However, several skeletal muscle sodium
channel mutations that cause hypokalemic periodic paralysis with
myotonia have been shown to cause enhanced slow inactivation (Takahashi
and Cannon, 1999; Jurkat-Rott et al., 2000; Struyk et al., 2000). In
addition, decreased sodium channel activity can cause epilepsy, as
demonstrated by null mutations in the SCN1A gene in patients
with severe myoclonic epilepsy of infancy (Claes et al., 2001). One
possibility is that seizure activity results from decreased firing of
inhibitory neurons, which causes increased firing in postsynaptic neurons.
The fact that the effects of the DIV mutation are more consistent with
hyperexcitability than those of the DII mutation is consistent with the
differences in clinical severity of the two GEFS+ 2 mutations (Baulac
et al., 1999; Moulard et al., 1999; Escayg et al., 2000). The DIV
mutation results in more severe clinical sequelae, as assessed by the
presence of afebrile and generalized seizures. In a family with the DIV
R1648H mutation, 11 of 12 affected individuals presented with
generalized epilepsy in addition to febrile seizures (Baulac et al.,
1999; Escayg et al., 2000). In contrast, only 5 of 11 affected
individuals with the DII T875M mutation presented with afebrile
seizures and generalized epilepsy (Moulard et al., 1999; Escayg et al.,
2000).
Many antiepileptic drugs such as phenytoin, carbamazepine, and valproic
acid are known to block sodium channels in a use-dependent manner
(Dichter, 1991; Macdonald and Kelly, 1994), thus reducing the
equilibrium current maintained during high frequency depolarizations. These drugs slow recovery from inactivation of wild-type sodium channels by binding to and stabilizing the inactivated state (McLean and Macdonald, 1986; Macdonald and Kelly, 1994; Kuo and Lu, 1997). The
result is a use-dependent block that reduces the ability of the channel
to maintain high-frequency action potential firing during sustained
membrane depolarization. These drugs have been thought to reduce
seizure activity by decreasing the excitability of sodium channels
downstream of the actual cause of the seizure. The results for the DIV
mutant suggest that these drugs also may act directly on hyperexcitable
sodium channels in patients with GEFS+ 2.
The arginine in the S4 segment of DIV (R1648 in
rNav1.1) is absolutely conserved in mammalian and
invertebrate voltage-gated sodium channels (Goldin, 1999; Escayg et
al., 2000). The effects of substituting histidine for this arginine
have been examined by a number of investigators, and the effects have
varied depending on the sodium channel isoform that has been examined.
With respect to voltage dependence, we have shown that R1648H in
rNav1.1 shifts the voltage dependence of
inactivation slightly in the hyperpolarized direction without affecting
the voltage dependence of activation when expressed as an subunit
alone. When coexpressed with the 1 subunit, there were no
significant effects on the voltage dependence of activation or
inactivation. Kühn and Greeff (1999) analyzed the same
arginine-to-histidine mutation in rNav1.2
(R1638H) with Xenopus oocytes. They observed a shift in the
opposite (depolarized) direction for the voltage dependence of
inactivation, with no shift in the current-voltage relationship when
the subunit was expressed alone. Alekov et al. (2000) studied the
comparable mutation (R1460H) in the hNav1.4
skeletal muscle sodium channel in tsA201 cells. They observed shifts in
the negative direction for the voltage dependence of activation and
inactivation. Neither Kühn and Greeff (1999) nor Alekov et al.
(2000) examined the mutant channels in the presence of the 1 subunit.
With respect to kinetics, we have shown that the R1648H mutation in
rNav1.1 dramatically accelerates recovery from
inactivation in both the absence and presence of the 1 subunit, with
a slight slowing of inactivation when the subunit is expressed
alone. The results of Alekov et al. (2000) for the R1460H mutation in hNav1.4 are similar to our data, with slightly
slower inactivation and markedly faster recovery from inactivation when
the subunits were expressed alone. Kühn and Greeff (1999)
observed no significant effects of the corresponding R1638H mutation in
rNav1.2 on the time course of recovery from
inactivation when the subunits were expressed alone. With respect
to the phenotype of GEFS+ 2, we believe that the most relevant results
are those obtained by using the orthologous channel in the presence of
the 1 subunit, in which case the most dramatic effect is a
significant acceleration in recovery from inactivation. However, it is
possible that the presence of either the 2 or 3 subunits might
alter channel function to result in other differences between the
mutant and wild-type channels.
In summary, we have shown that two SCN1A mutations that were
identified previously as causing GEFS+ 2 alter the electrophysiological properties of the sodium channel. The DIV mutation accelerated recovery
from inactivation and reduced the frequency dependence of the channel,
whereas the DII mutation enhanced slow inactivation and increased the
frequency dependence of the channel. The DIV mutation results in a
phenotype of hyperexcitability, whereas the DII mutation results in a
phenotype of hypoexcitability, suggesting that either an increase or
decrease in sodium channel activity can result in seizures. Additional
mutations in SCN1A that cause GEFS+ 2 (Escayg et al., 2001;
Wallace et al., 2001) and a mutation in SCN2A that causes
febrile seizures associated with afebrile seizures (Sugawara et al.,
2001) have been identified since, making it likely that there will be
even more mechanisms by which sodium channel alterations cause
epilepsy. Understanding these mechanisms should provide important
insights into the etiology of epilepsy in GEFS+ 2 patients and
facilitate the development of animal models for the study of GEFS+ 2 and targeted drug discovery.
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FOOTNOTES |
Received Feb. 1, 2001; revised June 25, 2001; accepted July 20, 2001.
This work was supported by National Institutes of Health (NIH) Grants
NS26729 (A.L.G.) and NS34609 (M.H.M.). J.S. was supported by NIH
Training Grant T32-NS07444, and A.E. acknowledges a fellowship from the
American Epilepsy Society with support from UCB Pharma, Smyrna, GA. We
thank Annie Lee, Wei Zhou, and A. J. Barela for helpful
discussions during the course of this work and Mimi Reyes for excellent
technical assistance.
Correspondence should be addressed to A. L. Goldin at the above
address. E-mail: AGoldin{at}uci.edu.
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REFERENCES |
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ABSTRACT
INTRODUCTION
