Effects of local anesthetics on
Na+ channels containing the
equine hyperkalemic periodic paralysis mutation
Rajan L.
Sah,
Robert G.
Tsushima, and
Peter H.
Backx
Departments of Medicine and Physiology, University of Toronto,
Toronto, Ontario, Canada M5G 1L7
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ABSTRACT |
We examined the
ability of local anesthetics to correct altered inactivation properties
of rat skeletal muscle Na+
channels containing the equine hyperkalemic periodic paralysis (eqHPP)
mutation when expressed in Xenopus
oocytes. Increased time constants of current decay in eqHPP channels
compared with wild-type channels were restored by 1 mM benzocaine but
were not altered by lidocaine or mexiletine. Inactivation curves, which were determined by measuring the dependence of the relative peak current amplitude after depolarization to
10 mV on conditioning prepulse voltages, could be shifted in eqHPP channels back toward that
observed for wild-type (WT) channels using selected concentrations of
benzocaine, lidocaine, and mexiletine. Recovery from inactivation at
80 mV (50-ms conditioning pulse) in eqHPP channels followed a
monoexponential time course and was markedly accelerated compared with
wild-type channels (
WT = 10.8 ± 0.9 ms;
eqHPP = 2.9 ± 0.4 ms). Benzocaine slowed the time course of recovery
(
eqHPP,ben = 9.6 ± 0.4 ms
at 1 mM) in a concentration-dependent manner. In contrast, the recovery
from inactivation with lidocaine and mexiletine had a fast component
(
fast,lid = 3.2 ± 0.2 ms;
fast,mex = 3.1 ± 0.2 ms),
which was identical to the recovery in eqHPP channels without drug, and
a slow component (
slow,lid = 1,688 ± 180 ms;
slow,mex = 2,323 ± 328 ms). The time constant of the slow component of the
recovery from inactivation was independent of the drug concentration,
whereas the fraction of current recovering slowly depended on drug
concentrations and conditioning pulse durations. Our results show that
local anesthetics are generally incapable of fully restoring normal WT
behavior in inactivation-deficient eqHPP channels.
sodium channel; inactivation
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INTRODUCTION |
NORMAL ACTION POTENTIALS in nerve, heart, and skeletal
muscle cells depend critically on a transient influx of
Na+ through voltage-gated
Na+ channels. After membrane
depolarization, Na+ channels open
or "activate" and subsequently rapidly inactivate. Channel
openings are not possible again until channels "recover" from
inactivation, a process requiring several milliseconds after membrane
repolarization. A number of mutations of the
-subunit of skeletal
and cardiac Na+ channels disrupt
inactivation, thereby causing disease. For example, several mutations
of the human skeletal muscle gene (SCN4A) have been associated with
heritable muscle diseases including hyperkalemic periodic paralysis
(HPP), paramyotonia congenita, and
Na+ channel myotonia (18, 33, 40,
42), whereas mutations in cardiac
Na+ channel genes result in a form
of the long Q-T syndrome (7, 28, 48). Functional studies of
muscle-related Na+ channel
mutations have shown that many of these mutations disrupt normal
Na+ channel inactivation, either
alone or in combination with other functional defects (7, 9, 13, 16,
18, 32, 35, 51).
Local anesthetic and antiarrhythmic drugs, such as lidocaine, act by
blocking Na+ flux through
voltage-gated Na+ channels and
have been shown to bind preferentially to inactivated conformation of
the cardiac Na+ channel (3-6,
8, 17, 24, 36, 47, 49). Therefore, local anesthetic agents might be
expected to reconstitute inactivation in
Na+ channels with defective
inactivation (18, 25). Within the "ball and chain" paradigm of
Na+ channel inactivation,
preferential binding of local anesthetics to the inactivated state
appears to stabilize the interaction of the inactivation gate with the
receptor, thereby locking channels into the inactivated conformation
(3-6, 8, 18, 22, 24, 47). This mechanism of local anesthetic
action could explain the clinical use of these agents in patients with
HPP (18, 25).
In the present study, we examined the effects of local anesthetics on
heterologously expressed rat skeletal muscle
Na+ channels containing the equine
HPP (eqHPP) mutation (F1412L), located in the third transmembrane
spanning region (S3) of domain IV (43). The equine mutation is
characterized by episodes of myotonia, weakness, and ultimately flaccid
paralysis in association with elevated serum
K+ levels (39). We and others (11,
23) have shown that the eqHPP mutation disrupts
Na+ channel inactivation.
Specifically, heterologously expressed rat skeletal muscle
Na+ channels containing the F1412L
mutation showed slowed inactivation kinetics, a rightwardly shifted
steady-state inactivation curve, and accelerated rates of recovery from
inactivation compared with wild-type channels (23).
Na+ channels recorded from
skeletal muscle myocytes from horses with eqHPP also showed evidence
for altered modal behavior (11). The objectives of our studies were to
examine whether the local anesthetics could promote inactivation and
whether the application of these agents could, under the appropriate
conditions, restore normal Na+
channel gating properties.
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MATERIALS AND METHODS |
Site-directed mutagenesis.
Site-directed mutagenesis of the rat skeletal muscle
Na+ channel (rSkM1) (46) was
performed to create the F1412L constructs. The mutations were
introduced into a 2.5-kb Sph
I-Kpn I cassette subcloned into
pGEM7f+ (Promega, Madison, WI)
using the oligonucleotide containing the appropriate base substitution.
Phenotypic selection was performed using the Kunkel method (30), and
the mutation was confirmed by dideoxy sequencing (45). The cassette was
subcloned into rSkM1 in pGW1H (British Biolabs, Oxford, UK) for
intranuclear injection of cDNA.
Expression of
Na+ channels in
Xenopus oocytes.
Oocytes were removed from adult female Xenopus
laevis frogs (NASCO, Fort Atkinson, WI; XENOPUS, Ann
Arbor, MI) anesthetized by immersion for 10-25 min in a 0.25%
solution of tricaine (Sigma Chemical, St. Louis, MO) in tap water.
Oocytes were digested with 2 mg/ml collagenase (type 1A, Sigma) in OR-2
containing (in mM) 88 NaCl, 2 KCl, 1 MgCl2, and 5 mM HEPES (pH 7.6).
Oocytes were stored at room temperature in ND96 containing (in mM) 96 NaCl, 2 KCl, 1.8 CaCl2, 1 MgCl2, and 5 HEPES (pH 7.6)
supplemented with 50 µg/ml gentamicin, 5 mM pyruvate, and 0.5 mM
theophylline. The nuclei of oocytes at stages IV-VI were coinjected
with 0.2 ng of
-subunit cDNA and 4.0 ng of
1-subunit cDNA (26). The
20-fold excess of
-subunits compared with
-subunits was used to
minimize the altered gating properties of
Na+ channels
-subunits observed when
expressed in oocytes (4, 12, 27). Injected oocytes were incubated
at 22°C for 24-48 h before recording.
Electrophysiological recording.
Whole cell currents were recorded at room temperature
(21-23°C) using two-electrode voltage-clamp techniques (Oocyte
Clamp OC-725A, Warner Instruments, Hampden, CT) with 3 M KCl in the pipette and a bath solution (ND96). Electrode pipettes were fabricated from 1.2-mm-OD thin-walled borosilicate glass (TW120F-6, World Precision Instruments, Sarasota, FL) pulled on a Sutter puller (model
P-87). Pipette tips were plugged with 1% agarose (in 3 M KCl) and had
a final resistance of 1-4 M
for the voltage-sensing electrode
and <2 M
for the current-passing electrode. Leak subtraction was
accomplished using a P/8 protocol from a holding potential of
120 mV. Currents were filtered at 2 kHz and digitized at 10 kHz
using an IBM-compatible computer, analog-to-digital interface (Warner
model PP-50 Lab1), and custom acquisition software. To minimize
difficulties associated with adequately voltage-clamping oocytes which
expressed large numbers of channels, whole cell recordings were limited
to oocytes expressing <5 µA of peak current. The local anesthetics
lidocaine (Sigma), mexiletine (Boehringer-Ingelheim, Ingelheim,
Germany), and benzocaine (ICN Biomedicals, Mississauga, Canada) were
introduced at the desired concentration in ND96 to the bath by
perfusion with at least 30 ml (bath volume
0.6 ml). After total bath
exchange (requiring <3 min), the drug was allowed to equilibrate with
the oocyte for a minimum of 6 min before recording.
Voltage protocols.
In the absence of drug, steady-state fast-inactivation curves were
constructed by plotting the relative peak current amplitude elicited in
response to "test" depolarizations (to
10 mV for 50 ms) as
a function of conditioning prepulse voltages (ranging from
120
to
10 mV for 50 ms) that were applied immediately before the
test pulse. Previous studies in rSkM1 channels have established that
50-ms prepulses are sufficiently long to allow equilibration between
the closed and inactivated states of the channel (19). Using similar
voltage protocols, we also examined the effects of local anesthetics on
the relative current in test pulses as a function of conditioning
prepulse voltage (from
120 to
10 mV). The prepulse
durations in the presence of drug were either 50 or 500 ms in length.
These measurements allow the assessment of the changes in the voltage
and time dependence of channels entering into the inactivation state
produced by local anesthetics. In these experiments, the voltage pulses
were applied every 10 s to ensure complete recovery from inactivation
between recordings.
Recovery from inactivation was assessed using a two-pulse protocol, in
which (conditioning) depolarizing pulses to
10 mV for either 50 or 500 ms were followed by repolarization to
80 mV (recovery
potential) for a variable duration, and this was immediately followed
by a (test) pulse to
10 mV (50 ms). The test current was
normalized to the conditioning current and plotted as a function of the
duration at the recovery potential. A repetition frequency of 0.1 Hz
was used for the recovery from inactivation protocols to ensure
complete recovery of channels between recordings. Previous studies have
established that 50-ms prepulses are sufficient to allow equilibration
between closed and fast inactivated states of rSkM1 channels (19). In
the presence of polar anesthetics like lidocaine and mexiletine,
equilibration of drug binding to the inactivated channels is dose
dependent but is largely complete within 500 ms in both expressed
cardiac and skeletal muscle Na+
channels with
20 mV prepulses in the presence of 100 µM
lidocaine (47).
Statistics and curve fitting.
The Marquardt-Levenberg algorithm in conjunction with a nonlinear
least-squares procedure was used to fit the functions shown below to
the experimental data. Data from experiments involving voltage
protocols using conditioning prepulses (i.e., relative peak current
measured in a test pulse as a function of conditioning prepulse
voltage) were fit with Boltzmann functions (24)
where
V is the step or conditioning
potential, V1/2
is the voltage midpoint of the function, and
k is the slope factor. In the
remainder of the paper, the curves generated from these fits are
referred to as "steady-state inactivation" curves in the absence of drug (17) and simple "inactivation curves" when drug is
present. This distinction emphasizes the possibility that, with
conditioning prepulses of short duration used in our studies,
incomplete equilibration of drug binding to inactivated channels
occurs.
Monoexponential or biexponential functions were used to fit recovery
from inactivation data and kinetic decay of the whole cell currents
after depolarization. For fits to the recovery from inactivation data,
we used the following biexponential function
where
Afast and
Aslow are the
amplitudes of the fast and slow components of the recovery,
respectively;
fast and
slow are the time constants for
the fast and slow components for recovery, respectively; and
t is the time spent at the recovery
potential (see above). When the recovery data were fit with a
monoexponential, Afast = Aslow = A and
fast =
slow =
. For the recovery
from inactivation, a time delay was commonly observed, particularly with benzocaine. Therefore, when fitting these data, we routinely included a time delay to account for this effect as described previously (37). For these fits, the
F-statistic and
F-distribution (P < 0.05) were used to assess
whether biexponential fits to the data gave significantly superior fits
compared with the monoexponential fits (21).
In the experiments using mexiletine and lidocaine, the amplitude of the
slow component of recovery from inactivation (i.e., Aslow) was fit
with the binding isotherm equation
where
[LA] is the concentration of either lidocaine or
mexiletine, n is the Hill coefficient,
and IC50,LA is the concentration of drug resulting in 50% of the inactivated channels being drug bound.
Combined data are presented as means ± SE. Statistical significance
was determined using an unpaired Student's
t-test (21) and a confidence limit of
95%.
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RESULTS |
Phenotypic alteration of eqHPP channels.
The electrophysiological changes associated with eqHPP channels are
illustrated in Fig.
1A,
showing scaled raw Na+ current
traces recorded from wild-type (WT) and eqHPP channels after step
depolarizations to
10 mV from a holding potential of
120
mV. Notice that WT Na+ currents
decay more rapidly than eqHPP channels:
WT = 0.97 ± 0.08 ms (n = 6) vs.
eqHPP = 2.14 ± 0.07 ms
(n = 15). Slowed current decay in
eqHPP channels compared with WT can originate from either slowed entry
into the inactivated state (2, 23, 32, 34, 38) or voltage shifts in
channel activation (14, 16, 24). Figure
1B shows that there was a significant
rightward shift in the steady-state inactivation curve of eqHPP mutant
channels (solid squares) by ~6 mV relative to WT channels (open
squares)
(V1/2,eqHPP =
47.6 ± 0.6 mV, n = 10 vs.
V1/2,WT =
53.1 ± 0.4 mV, n = 7; P < 0.05), suggesting that the eqHPP
mutation has altered channel inactivation (23). This is further
supported by the observations summarized in Fig.
1C, illustrating that the recovery
from inactivation (after 50-ms conditioning pulse) is accelerated more
than threefold for eqHPP mutant channels compared with WT channels at
80 mV (i.e.,
eqHPP = 2.9 ± 0.4 ms, n = 7 vs.
WT = 10.8 ± 0.9 ms,
n = 7;
P < 0.01).

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Fig. 1.
A: whole cell currents recorded from
wild-type (WT) and equine hyperkalemic periodic paralysis (eqHPP)
mutant Na+ channels expressed in
oocytes. Currents were evoked by a 50-ms voltage step to 10 mV
from a holding potential of 120 mV. Currents have been
normalized to have same peak amplitudes; peak inward current amplitude
was 3.4 µA for eqHPP trace and 2.9 µA for WT trace. eqHPP current
displays a slowed rate of decay relative to decay of WT current
( eqHPP = 2.14 ± 0.07 ms,
n = 15;
WT = 0.97 ± 0.08 ms,
n = 6).
B: steady-state inactivation curves
constructed by plotting relative current magnitude as a function of
50-ms prepulse voltages for WT ( ; n = 7) and eqHPP ( ; n = 10) channels.
Inactivation was induced by 50-ms conditioning prepulses from
100 to 10 mV in 5-mV increments. Extent of inactivation
was then assessed by a step depolarization to 10 mV.
C: recovery from inactivation of WT
( ; n = 9) and eqHPP
mutant ( ; n = 7) was assayed using
a 2-pulse protocol at a recovery potential and holding potential of
80 mV. On average, eqHPP mutant recovers significantly faster
from inactivation ( eqHPP = 2.9 ± 0.4 ms) than WT channel
( WT = 10.8 ± 0.9 ms)
(P < 0.05).
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Effect of local anesthetic on inactivation properties of eqHPP.
Because local anesthetics selectively bind to the inactivated state of
Na+ channels and thereby stabilize
the inactivated state (3-6, 8, 17, 22, 24, 36), we tested the
effects of lidocaine, mexiletine, and benzocaine on the inactivation
properties of eqHPP channels. Figure
2A shows
raw current traces recorded in response to 50-ms step depolarizations
to
10 mV from a holding potential of
80 mV before and
after the application of lidocaine, mexiletine, and benzocaine. At the
indicated dosages, lidocaine, mexiletine, and benzocaine reduced the
peak current of eqHPP channels. This reduction of peak current is
generally referred to as tonic block and probably represents binding of
these drugs to the closed state or possibly rapid block of the open
state (5, 24). Normalization of current traces in the presence of the
drug to current traces in the absence of drug, as illustrated in Fig.
2B, shows that 600 µM lidocaine and
mexiletine do not noticeably accelerate the rate of current decay
(
eqHPP = 2.14 ± 0.07 ms,
n = 15;
lid = 2.29 ± 0.21 ms,
n = 5;
mex = 2.43 ± 0.16 ms, n = 5), whereas 1 mM benzocaine
clearly enhances the rate of current decline
(
ben = 1.11 ± 0.06 ms, n = 5;
P < 0.05). The dependence of the
time constant for whole cell current decline recorded after
depolarizations to
10 mV as a function of the concentration of
drug is illustrated in Fig. 2C.
Lidocaine and mexiletine caused a very modest and statistically
insignificant acceleration in the rate of current decay of the
Na+ current as a function of the
drug concentration (P > 0.05) as previously demonstrated (47). For example, after the application of 600 µM lidocaine, the time constant
for decay decreased from 2.33 ± 0.17 to 1.95 ± 0.19 ms. In contrast, the time constant of
Na+ current reduction decreased
with elevated benzocaine concentrations, asymptotically reaching a
limiting value of 1.05 ± 0.03 ms at 1 mM benzocaine. These
differences between lidocaine and mexiletine compared with benzocaine
likely result from differences in hydrophobicity between these
different local anesthetics (8, 17, 24, 41).

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Fig. 2.
A: reduction of peak current of eqHPP
mutant in presence of local anesthetics. Currents were evoked by 50-ms
step depolarizations to 10 mV from a holding potential of
80 mV. Only first 30 ms are shown.
B: current traces from
A normalized with respect to peak
eqHPP mutant current. Neither 300 µM lidocaine nor 300 µM
mexiletine accelerated rate of current decay of eqHPP mutant, whereas 1 mM benzocaine markedly increased rate of decay in this oocyte.
C: time constants
( h) of current decay of eqHPP
mutant vs. concentration of lidocaine
(left), mexiletine
(middle), and benzocaine
(right). Lidocaine and mexiletine
data were well fit by a linear regression function
[r2
(lidocaine) = 0.95;
r2 (mexiletine) = 0.95] with a slope that was not significantly different from
zero. Benzocaine data were fit to a monoexponential function.
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As a result of the ability of benzocaine to enhance the rate of current
decline, we speculated that, at the appropriate concentration, this
agent might reconstitute WT behavior to eqHPP channel with respect to
other aspects of inactivation. Figure
3A shows
superimposed current traces of WT channels and of HPP channels plus 1 mM benzocaine after depolarization to
10 mV; the current traces
were scaled to match the peak currents. In the presence of 1 mM
benzocaine, the current decay of HPP channels measured at
10 mV
mimic that of WT channels without benzocaine
(
WT = 0.97 ± 0.08 ms,
n = 6 vs.
ben = 1.11 ± 0.06 ms,
n = 5). However, as illustrated in Fig. 3B, the time constants for whole
cell current decay become largely voltage independent in the presence
of 1 mM benzocaine in eqHPP channels (solid triangles) compared with WT
(open circles) or eqHPP (solid squares) channels without drug. As a
result, the relationship between the inactivation time constants and
voltage in eqHPP channels treated with benzocaine cross over the
corresponding relationship in WT channels at around
10 mV.
Therefore, although benzocaine does reverse the slowed inactivation
rate of eqHPP at high concentrations (i.e., 1 mM), it does not
generally confer WT behavior to eqHPP channels at all voltages. In
addition, as discussed more fully below but not shown in Fig. 3, the
peak current magnitude was reduced by ~50% with this dose of
benzocaine (Table 1).

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Fig. 3.
A: superimposed current traces of WT
channel and eqHPP mutant in presence of 1 mM benzocaine normalized with
respect to peak current. Currents were evoked as described in Fig. 2.
Peak current in WT traces was 3.3 µA, and current in
benzocaine-treated eqHPP channels was 2.3 µA. eqHPP mutant in
presence of 1 mM benzocaine closely approximates current decay of WT
channel ( WT = 0.97 ± 0.08 ms, n = 6;
eqHPP,ben = 1.11 ± 0.06 ms,
n = 5).
B: time constants of current decay vs.
voltage of WT channels ( ; n = 6)
and eqHPP mutant channels in absence ( ;
n = 6) and presence of 1 mM benzocaine
( ; n = 5). Notice that little
voltage dependence is observed in time constant of current decay of
eqHPP mutant channels when 1 mM benzocaine is present.
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The rightward shift of the steady-state inactivation curve along the
voltage axis (Fig. 1B) of eqHPP
channels relative to WT channels requires that more positive prepulses
are necessary to inactivate the same proportion of eqHPP channels in
comparison to WT channels (23). Therefore, we examined the effects of
local anesthetics on the relative peak current amplitude recorded
during test depolarizations (
10 mV for 50 ms) as a function of
the conditioning prepulse voltage having a duration of 50 or 500 ms.
The relative peak current amplitudes as a function of prepulse voltage
are plotted in Fig. 4 for WT channels
(solid circles) and eqHPP channels in the absence (solid squares) and
presence of drug (open symbols). The estimated
V1/2 values,
obtained by fitting the data for these experiments to the Boltzmann
equation (see MATERIALS AND METHODS) for these experiments, are summarized in Table
2. With 50-ms conditioning pulses, 600 µM
lidocaine (Fig. 4A), 300 µM
mexiletine (Fig. 4B), and 30 µM
benzocaine (Fig. 4C) shifted the
inactivation curves of eqHPP channels sufficiently leftward to cause
superposition with the WT steady-state inactivation curves. These
results demonstrate that with short conditioning pulses, benzocaine is
far more potent than lidocaine or mexiletine in promoting entry into
the inactivated state. With longer conditioning prepulses (500 ms),
illustrated in Fig. 4, right, 100 µM
lidocaine and mexiletine shifted the dependence of the relative current
on prepulse voltage for eqHPP sufficiently leftward to approximately
overlay the WT curve, whereas 30 µM benzocaine was again similar to
WT (Table 2). Thus increasing the conditioning pulse duration from 50 to 500 ms had little effect on benzocaine binding, consistent with
rapid equilibration of drug binding to the channel (8, 17, 36, 41). In
contrast, prolongation of the conditioning pulse duration in the
presence of lidocaine or mexiletine did markedly enhance drug binding
as expected for the slow binding kinetics displayed by polar local anesthetics (5, 17, 41, 47).

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Fig. 4.
Effect of local anesthetics on voltage dependence of channel
inactivation in response to conditioning prepulses. Relative peak
current observed in WT channels ( ) and eqHPP channels in absence
( ) and presence (open symbols) of lidocaine
(A), mexiletine
(B), and benzocaine
(C) after conditioning prepulses to
various voltages with a duration of either 50 ms
(left) or 500 ms
(right). Solid lines were obtained
from best fits to experimental data using a Boltzmann equation. In
absence of drug, these plots measure steady-state inactivation
properties of WT and eqHPP channels (19). Lidocaine, mexiletine, and
benzocaine shift voltage dependence of these inactivation curves to
more hyperpolarized potentials. With the use of 500-ms conditioning
pulses, 30 µM benzocaine, 100 µM lidocaine, or 100 µM mexiletine
is able to shift inactivation curves measured in eqHPP channels so that
they nearly overlay that recorded in WT channels without drug. rSkM1,
rat skeletal muscle Na+ channel.
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Table 2.
Estimates of V1/2 and k derived from fits to
measured relative peak current amplitude as a function of 500-ms
conditioning prepulse voltages for WT channels and eqHPP channels
in the absence and presence of drug
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Further information about local anesthetic effects on the stability of
the inactivated state of eqHPP channels can be derived from the rate at
which channels recover from inactivation in the presence of drug. The
kinetics of recovery from inactivation in oocytes were assessed using a
two-pulse protocol wherein conditioning pulses to
10 mV for 50 or 500 ms were followed by repolarization to
80 mV for a
variable period before the introduction of a second test depolarization
to
10 mV for 50 ms. Figure 5 summarizes the effects of lidocaine and mexiletine on the kinetics of recovery from
inactivation using 50-ms conditioning pulses in WT channels (solid
circles) and eqHPP channels with (open symbols) and without (solid
squares) lidocaine (left) or
mexiletine (right). On the brief
time scale in Fig. 5, A and
B, it can be seen that the degree of
current recovery in eqHPP channels after 20 ms of repolarization to
80 mV was depressed by both lidocaine and mexiletine in a dose-dependent manner. Furthermore, Fig. 5,
A and
B, demonstrates that during the first
20 ms, the time course of recovery from inactivation of eqHPP in the
presence of lidocaine or mexiletine is reasonably well described by a
monoexponential function (solid lines). Indeed, inclusion of a second
exponential component did not improve the quality of the fit (see
MATERIALS AND METHODS). Despite the
incomplete recovery of current on the time scale displayed in Fig. 5,
A and
B, the time constants estimated from
the monoexponential fits did not change as the drug concentration was
varied from 0 to 600 µM (see Fig. 5,
E and
F). As a result, the time course of
recovery of eqHPP channels with drug application (Fig. 5, open symbols)
never matched the recovery kinetics of WT channels (Fig. 5, solid
circle). Extension of the recovery period as shown in Fig. 5,
C (lidocaine) and
D (mexiletine), reveals that full
recovery from inactivation of eqHPP channels requires longer than 2 s
in the presence of these agents. The full inactivation recovery curves shown in Fig. 5, C and
D, required a biexponential function
to accurately fit the experimental data
(P < 0.05). As summarized in Fig. 5,
E and
F, the fast time constants for
recovery (solid squares) in the presence of either lidocaine (Fig.
5E) or mexiletine (Fig.
5F), estimated from fits to the data
in Fig. 5, C and
D, did not vary with concentration;
that is, the slopes of the relationship between
fast and drug concentration
were not significantly different from zero
(P > 0.20 for lidocaine and
P > 0.32 for mexiletine). Additionally, the fast time constants for recovery were not found to be
statistically different from the time constant for recovery of eqHPP
channels in the absence of drug
(
fast,lid = 3.2 ± 0.2 ms,
n = 15 and
fast,mex = 3.1 ± 0.2 ms,
n = 15 vs.
eqHPP = 2.9 ± 0.4 ms,
n = 7). Lidocaine and mexiletine
application did, however, induce the appearance of a much slower time
constant for recovery from inactivation not observed in the untreated
eqHPP channels. It is apparent from inspection of Fig. 5,
C and
D, that the relative amount of
recovery occurring on a slow time scale (i.e.,
Aslow) increased with elevated drug levels (also see Fig.
6). On the other hand, the
slope of the relationship between the slow time constant (i.e.,
slow) for recovery and the
drug concentration was not significantly different from zero (Fig. 5,
E and
F, solid circles), again indicating
independence of recovery kinetics on drug levels.

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Fig. 5.
A and
B: recovery from inactivation over 20 ms of WT channels ( , n = 9) and of
eqHPP channels in absence ( ) and presence (open symbols) of
lidocaine (A,
n = 3-7) and mexiletine
(B, n = 3-7) using 50-ms conditioning pulses. Data were fit with a
monoexponential function. C and
D: recovery from inactivation/block of
WT and eqHPP mutant channels shown in
A and
B extended over 2 s in absence and
presence of lidocaine (C) and mexiletine (D). Data over
2-s period were best fit with a biexponential function (solid lines).
E and
F: average fast
( fast, ) and slow
( slow, ) time constants for
recovery from inactivation estimated by fitting data in
C and
D to a biexponential function plotted
as a function of lidocaine (E) or
mexiletine (F) concentration. Note
that neither time constant depended on drug concentration.
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Fig. 6.
Recovery from inactivation/block over 2 s of WT channel and eqHPP
mutant in absence and presence of lidocaine
(A) and mexiletine
(B) using a 500-ms conditioning
pulse. Again, data were best fit with a biexponential function (solid
lines). Symbols are as indicated in Fig. 5.
C and
D: magnitude of slow component of
recovery from block of eqHPP mutant, estimated from biexponential fits,
as a function of lidocaine (C) and
mexiletine (D) concentration using
50-ms ( ) or 500-ms ( ) conditioning pulses. Data were fit with a
binding isotherm using a nonlinear least-squares algorithm (see
MATERIALS AND METHODS).
IC50 values and Hill coefficients
(n) for lidocaine and mexiletine
using 50-ms conditioning pulse duration were estimated as follows:
IC50,lid = 230 ± 12 µM, nlid = 1.6 ± 0.2, n = 4; and
IC50,mex = 293 ± 9 µM,
nmex = 1.3 ± 0.1, n = 4. With 500-ms conditioning
pulse durations, values were as follows:
IC50,lid = 44 ± 6 µM,
nlid = 1.5 ± 0.1, n = 4;
IC50,mex = 44 ± 8 µM,
nmex = 1.7 ± 0.1, n = 4.
|
|
By comparison, Fig. 6 summarizes the kinetics of recovery from
inactivation measured in eqHPP channels in the presence of lidocaine
(A) or mexiletine
(B) using conditioning pulses with a
500-ms duration. Inspection of Fig. 6,
A and
B, shows that the magnitude of the
slow and fast time constants of recovery were not notably different
when 500- vs. 50-ms conditioning pulses were used. However, the
amplitude of the slow component for any given concentration of drug was
much greater using 500-ms conditioning pulses compared with 50-ms
pulses. For example, in the presence of 100 µM lidocaine, only 30%
of the overall recovery occurs slowly when 50-ms conditioning pulses
were used (Fig. 5C, open inverted triangles) compared with ~80% with 500-ms conditioning pulses (Fig.
6A, open inverted triangles). Similar
effects of conditioning pulse duration can be seen with mexiletine by
comparing Figs. 5D and
6B. This dependence of the relative
amplitude of the slowly recovering component
(Aslow) on
conditioning pulse duration [i.e., 50 ms (solid squares) and 500 ms (open circles)] for different drug concentrations is plotted
in Fig. 6C for lidocaine and in Fig.
6D for mexiletine application.
Prolonging the conditioning pulse from 50 to 500 ms resulted in about a
fivefold shift in the estimated
IC50 for both lidocaine
[Fig. 6, left,
IC50,lid = 230 µM (50 ms) or 44 µM (500 ms)] and mexiletine [Fig. 6,
right, IC50,mex = 293 µM (50 ms) or
44 µM (500 ms)] binding to eqHPP channels.
The data in Figs. 5 and 6 are consistent with previous studies in
cardiac Na+ channels using
lidocaine (5, 6) and suggest that recovery from inactivation involves
two populations of channels: one drug-free population recovering at
rates indistinguishable from untreated channels and one drug-attached
population whose recovery is nearly 500-fold slower (5, 6). The
differences in the number of drug-bound channels observed as a function
of the conditioning pulse duration is also consistent with the previous
studies using polar local anesthetics like lidocaine and mexiletine (6,
8, 17, 41, 47). The existence of two populations of channels in the
presence of lidocaine and mexiletine underlies the complete lack of
correspondence between the recovery time course of drug-treated eqHPP
channels (open symbols) and either WT (solid circles) or untreated
eqHPP (solid squares) shown in Figs. 5 and 6.
In contrast, Fig. 7 shows that the recovery
properties of benzocaine-treated eqHPP channels using 50-ms
(A) and 500-ms
(B) conditioning pulses are very
different from lidocaine- or mexiletine-treated channels. Figure 7,
A and
B, shows that the fractional recovery from inactivation of eqHPP channels at different benzocaine
concentrations using either 50- or 500-ms conditioning pulse durations
follows a monoexponential time course as expected from the rapid
kinetics of benzocaine binding to
Na+ channels (see
DISCUSSION). Figure 7,
A and
B, further demonstrates that the
kinetics of recovery from inactivation for WT channels closely matches
eqHPP recovery in the presence of 1 mM and 300 µM benzocaine when 50- and 500-ms conditioning pulses were used, respectively. The dependence
of the time constants for recovery from inactivation with increasing
amounts of benzocaine is presented in Fig. 7,
C and
D, which shows that 1 mM benzocaine
causes a nearly three- to sixfold slowing in the rate of recovery from inactivation for eqHPP channels. Although it appears from Fig. 7,
A and
B, that benzocaine, at a concentration
between 300 µM and 1 mM, causes eqHPP channels to mimic the recovery
properties of WT channels, it should be remembered that normalization
of the data has eliminated the effects tonic block by benzocaine as
illustrated in Fig. 2A. The
concentration dependence of tonic block for benzocaine, as well as
lidocaine and mexiletine, are summarized in Table 1.

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|
Fig. 7.
A and
B: recovery from inactivation of WT
channel ( ; n = 4-9) and eqHPP
mutant in absence ( ; n = 7) and
presence of 300 µM ( ; n = 10) and
1 mM ( ; n = 7) benzocaine recorded
for 50-ms (A) and 500-ms
(B) conditioning pulses. Data were
fit with a monoexponential function. Note that time delays were
required to achieve adequate fits in presence of benzocaine (31).
C and
D: plots of recovery time constants of
eqHPP channels as a function of benzocaine concentration with 50-ms
(C) and 500-ms
(D) conditioning pulse durations.
|
|
 |
DISCUSSION |
Our data show that lidocaine, mexiletine, and benzocaine selectively
modified inactivation properties of eqHPP channels. These observations
are consistent with data from many previous studies which have
established that local anesthetics preferentially bind to the
inactivated as well as possibly the open conformation of Na+ channels (1-6, 8, 17, 18,
22, 24, 47) while binding very weakly to the closed conformations (6,
15, 47). We postulated that local anesthetics might be capable of
restoring inactivation to inactivation-deficient
Na+ channels carrying the equine
periodic paralysis mutation (i.e., F1412L). Because local anesthetics
are currently used clinically in the treatment of some forms of muscle
myotonias, we examined whether these agents could confer the normal
inactivation properties of WT channels onto eqHPP channels.
Neither lidocaine, mexiletine, nor benzocaine could fully bestow normal
WT inactivation properties to eqHPP channels under a specific set of
experimental conditions. For example, benzocaine at a concentration of
1 mM slows the rate of recovery from inactivation with 50-ms prepulses
of eqHPP channels, making recovery kinetics nearly identical to WT
channels. Moreover, benzocaine restores WT behavior with respect to the
rate of Na+ current inactivation
in eqHPP channels after depolarization to
10 mV, but this
property was not observed at other membrane potentials, particularly
below
20 mV (see Fig. 3B). At
concentrations of benzocaine above 30 µM, the inactivation curves
(measured as the dependence of peak current amplitude on prepulse
voltage) for eqHPP channels were located leftward of that observed in
WT channels without drug. Therefore, despite benzocaine's ability to
correct the major inactivation defects observed in eqHPP channels, no single dose of this agent could fully restore WT inactivation behavior.
In comparison with benzocaine, lidocaine and mexiletine did not
significantly alter the rate of
Na+ current inactivation even at
doses as high as 600 µM lidocaine (47). With the use of a 500-ms
conditioning pulse protocol, the application of 100 µM lidocaine or
mexiletine causes the eqHPP inactivation curves to overlap with the WT
curve, whereas at lower and higher dosages, the inactivation curves for
eqHPP channels were located rightward and leftward relative to WT
channels, respectively. When examined over a 2-s period, recovery from
inactivation occurred in two distinct phases at all concentrations of
lidocaine and mexiletine studied. The amplitude of the slow component
of recovery depended on drug levels and could be fit using a binding
equation with a Hill coefficient between 1.3 and 1.7 (Fig. 6), which is different from the value of 1 reported previously in cardiac
Na+ channels (5). These
discrepancies might reflect genuine differences in the mechanism of
block by local anesthetics between eqHPP, rSkM1, and cardiac
Na+ channels (37, 47).
Alternatively, this lack of correspondence could arise from differences
in the experimental conditions. For example, Bean et al. (5) used 5-s
conditioning prepulses, which is sufficiently long to induce slow
inactivation (6, 19, 47). Hill coefficients greater than one might also
reflect incomplete equilibration between the drug and channels in our
experiments (47). Regardless, our studies demonstrate that lidocaine
and mexiletine cannot faithfully restore normal inactivation properties in eqHPP channels at any drug concentration.
Differences in the effects of lidocaine and mexiletine vs. benzocaine
probably reflect the faster binding and unbinding rates of benzocaine
compared with lidocaine and mexiletine to the open and inactivated
states of channels (8, 17, 24, 36, 41). With 50-ms conditioning pulses,
a 10-fold lower concentration of benzocaine compared with lidocaine or
mexiletine was required to produce the same degree of shift in the
inactivation curves (i.e., same change in
V1/2). We
initially used conditioning pulses of 50-ms duration to crudely mimic
the late phase of action potential duration observed in skeletal muscle
(20, 29). However, a nearly threefold lower concentration of lidocaine
and mexiletine is required to produce the same change in
V1/2 (Fig. 4)
when 500- vs. 50-ms conditioning pulses are used, reflecting incomplete equilibrium with short prepulses (5, 6, 17, 24, 49). Interestingly,
prolonging the prepulse duration beyond 500 ms (data not shown) did not
result in further measurable shifts in V1/2 as reported
previously (47).
The initial focus of our experiments was to examine the ability of
local anesthetic to modify and correct fast inactivation properties in
eqHPP channels. Although the kinetics of mexiletine and lidocaine
interaction with Na+ channels are
too slow to confer WT kinetics onto eqHPP channels, they are
nevertheless used clinically to abolish the multiple repetitive firing
of skeletal muscle in human myotonia patients (25) at doses well below
those used in our study. Cannon et al. (10) used a mathematical
two-compartment model of skeletal muscle to demonstrate that incomplete
inactivation of subpopulations of (mutant)
Na+ channels within muscle cells
was sufficient to cause repetitive trains of action potentials and,
therefore, myotonia in patients with HPP. Thus preferential blockade of
persistently active Na+ channels
rather than reconstitution of inactivation might be key for clinical
utility. In this regard, previous studies in cardiac channels
containing mutations causing long Q-T syndrome (1) and
inactivation-deficient rSkM1 channels (3) have demonstrated that
lidocaine blocks noninactivating currents with a much higher affinity
than the peak current. On the other hand, many
Na+ channel mutations associated
with skeletal muscle disease (9, 13, 11, 14, 16, 32, 34, 35, 39, 40,
42, 43), including eqHPP (23), and long Q-T syndrome in heart (7, 28,
48) significantly accelerate the rate of recovery from inactivation and
shorten the effective refractory period. This feature of mutant
channels has also been suggested to contribute to repetitive
depolarizations in tissues containing mutant channels (1, 14).
Therefore, slowing the rate of recovery, as we observed with lidocaine,
mexiletine, and benzocaine, could also be an important element of drug
action in disease treatment.
On the basis of the above discussion, fast agents like benzocaine might
be preferred in the treatment of some forms of muscle myotonia by more
rapidly promoting inactivation and blockade of non-inactivated
channels. Slower binding drugs would leave a subpopulation of drug-free
channels with persistent activity and reduced refractoriness (10, 34).
Alternatively, when excessive numbers of
Na+ channels are drug-bound, as
might occur during periods of high muscle activity, sluggish agents
like lidocaine and mexiletine are expected to remain bound to channels
for longer periods thus promoting muscle inexcitability. However, the
applicability of our results to the clinical treatment of muscle
disorders is limited for a number of reasons. First, our investigations
were restricted to the effects of local anesthetics on fast
inactivation. It has been argued previously that impairment of slow
inactivation is necessary for clinical disease (44). Moreover,
use-dependent binding of lidocaine to slow inactivated channels is
potentiated (4). Second, an important consideration guiding the choice of the Na+ channel modifiers in
the treatment of muscle disorders is their relative tissue-specificity
or their ability to discriminate between normal and
inactivation-deficient channels. Indeed, major limitations of using
Na+ channel modifiers in muscle
disease treatment are side effects associated with drug action on
normal channels (25). In this regard, the drug concentrations we used
far exceed the dosages commonly used clinically (25).
It is important to note that two analogs of lidocaine (i.e., mexiletine
and tocainide) are used clinically in the treatment of
Na+ channel-based skeletal muscle
disease like paramyotonia congenita as well as some forms of
hyperkalemic paralysis/paramyotonia but not in the treatment of HPP
(25). This observation is surprising, since the phenotypic changes
occurring with different disease-causing Na+ channel mutations often have
common biophysical properties (9, 13, 11, 23, 32, 34, 51). Our results
show that local anesthetics can differentially modify inactivation
properties in mutant channels associated with periodic paralysis in
horses. Therefore, because disrupted function in eqHPP channels is not unlike that seen in many other Na+
channel mutations (9, 13, 32, 34, 51), these studies may provide some
insights into the mechanism of local anesthetic action in the treatment
of muscle diseases.
 |
ACKNOWLEDGEMENTS |
We thank G. Mandel and W. A. Catterall for kindly providing the
-subunit of the rat skeletal muscle
Na+ channel and the
1-subunit of the rat brain
Na+ channel, respectively.
 |
FOOTNOTES |
This work was supported by the Muscular Dystrophy Association of Canada
and the Medical Research Council of Canada (to P. H. Backx). P. H. Backx holds a Medical Research Council of Canada scholarship award. R. L. Sah was supported by a Muscular Dystrophy of Canada Summer
Studentship. We are grateful for support for equipment provided by the
Alan Tiffin Trust, Toronto, Ontario, Canada.
Address for reprint requests: P. H. Backx, The Toronto Hospital,
General Division, CCRW 3-802, 101 College St., Toronto, Ontario,
Canada M5G 1L7.
Received 26 February 1997; accepted in final form 27 April 1998.
 |
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