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Anesthetic Pharmacology: Research Report

Propofol Blocks Human Skeletal Muscle Sodium Channels in a Voltage-Dependent Manner

Haeseler, Gertrud, MD*,; Störmer, Martina*,; Bufler, Johannes, MD†,; Dengler, Reinhard, MD†,; Hecker, Hartmut, MD‡,; Piepenbrock, Siegfried, MD*,; Leuwer, Martin, MD§

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doi: 10.1097/00000539-200105000-00021
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Abstract

Propofol provides some degree of “muscle relaxation” in a wide variety of clinical conditions in the absence of neuromuscular blocking drugs. Propofol reduces pathologically increased muscle tone (1), prevents drug-induced muscle rigidity (2), and provides jaw relaxation for conventional tracheal intubation (3). Adequate jaw relaxation requires larger propofol doses than suppression of airway reflexes assessed during facilitated endotracheal intubation (3). However, whether these effects are peripheral in origin is not known. The fact that propofol (and not thiopental) significantly reduces a succinylcholine-induced increase of masseter muscle tone (2) provides strong evidence for the hypothesis that a peripheral effect is involved in propofol-induced muscle relaxation. Intermittent IV propofol administration to a patient with severe tetanus reduced muscle tone after each bolus and also decreased spontaneous electromyography recordings. Neuromuscular transmission, assessed by evoked electromyography, was not affected (1). The hypothesis that a presumed peripheral effect does not involve impairment of neuromuscular transmission was strengthened by experiments on propofol-induced reduction in diaphragmatic contractility assessed with high- and low-frequency phrenic nerve stimulation in anesthetized dogs (4). Propofol-induced reduction of muscle contractility after direct stimulation has been shown for the isolated rat diaphragm in vitro and the rat gastrocnemius muscle in vivo(5). However, further conclusions about possible target sites could not be derived from that study. Although propofol affects voltage-gated sodium channels of other tissues (6,7), no reports on drug effects on the skeletal muscle isoform have been published. Sodium channels of the sarcolemma are responsible for the initiation and propagation of the action potential and, therefore, play a crucial role in muscle excitability. We investigated the effects of propofol on heterologously expressed skeletal muscle sodium channels as one possible target contributing to the reduction in muscle tone achieved clinically by the drug.

Methods

The gene SCN4A encodes the α-subunit of the principal voltage-gated sodium channel expressed in adult human skeletal muscle (8). A full-length construct of its complementary DNA, designated hSkM1, was made in the expression vector pRc/CMV (Invitrogen, San Diego, CA) (9). To obtain cells expressing α-subunits of human skeletal muscle sodium channels, the plasmid pRc/CMV-hSkM1 was transfected into human embryonic kidney (HEK 293) cells (American Tissue Culture Collection CRL 1573), by using calcium phosphate precipitation (10). Permanent expression was achieved by selection for resistance to the aminoglycoside antibiotic geneticin G418 (Life Technology, Eggenstein, Germany) (9). Transfected cells were a gift from Prof. Lehmann-Horn, Ulm, Germany. Successful channel expression was verified electrophysiologically. The clone has been used in several investigations (9,11).

Purified 2,6-diisopropylphenol (propofol) was a gift from Zeneca GmbH (Plankstadt, Germany). Propofol was prepared as 1-M stock solution in ethanol, light-protected, and stored in glass vessels at −20°C. The stock solution was dissolved directly in bath solution immediately before the experiments. Propofol-containing vials were vigorously vortexed for 60 min. The solution was applied via a glass-polytetrafluoroethylene perfusion system and a stainless steel superfusion pipette. The bath solution contained [mM] NaCl 140, MgCl2 1, KCl 4, CaCl2 2, HEPES 5, dextrose 5. Patch electrodes contained [mM] CsCl2 130, MgCl2 2, EGTA 5, HEPES 10. All solutions were adjusted to 290 mOsm by the addition of mannitol and to pH 7.4 by the addition of Cs(OH)2.

Standard whole-cell voltage-clamp experiments (12) were performed at 20°C. Each experiment consisted of test recordings with the drug present at only one concentration, and of drug-free control recordings before and after the test. Each cell was exposed to one test concentration only. At least four experiments were performed at each concentration. The amount of the diluent ethanol corresponding to the respective propofol concentration was added to the control solution up to a maximal ethanol concentration of 8.7 mM corresponding to a propofol concentration of 500 μM. At this concentration, ethanol had no effect on the peak current amplitude; the peak current was 2.2 ± 0.1 nA in the controls and 2.3 ± 0.1 nA in the presence of 8.7 mM of ethanol at −150-mV holding potential (n = 3).

For data acquisition and further analysis, we used the EPC9 digitally controlled amplifier in combination with Pulse and Pulse Fit software (HEKA Electronics, Lambrecht, Germany). The EPC9 provides automatic subtraction of capacitive and leakage currents by means of a prepulse protocol. The data were filtered at 10 kHz and digitized at 20 μs per point. Input resistance of the patch pipettes was at 1.8–2.5 MΩ, cells’ capacitances ranged between 9–15 picofarad; residual series resistance (after 50% compensation) was 1.2–2.5 MΩ. Experiments with an increase in series resistance were rejected. To minimize time-dependent shifts in the voltage dependence of steady-state inactivation (13), all test experiments were performed within 5 min of patch rupture. Under these experimental conditions, time-dependent hyperpolarizing shifts in control conditions were <−2 mV (11). Voltage-activated currents were studied by applying different voltage-clamp protocols described in the Results section or in the appropriate figure legends.

Drug effects on the peak current amplitude were assessed at a holding potential close to the normal resting potential in physiological conditions (−70 mV) (14), at hyperpolarized membrane potentials (−100 and −150 mV), or when a 2.5-s prepulse to −35 mV was introduced before the test pulse to induce slow inactivation (15). The residual sodium current (INa+) in the presence of drug (with respect to the current amplitude in control solution) was plotted against the applied concentration of the drug [C]. All data were presented as means ± sd. Fits of the Hill equation INa+Symbol to the data yielded the concentration for half-maximum channel blockade (IC50) and the Hill coefficient nH.

Symbol
Symbol

Statistical analysis of the concentration-response plots was performed to reveal differences related to the applied pulse protocol. Curve fitting and variable estimation of the Hill curves was performed by using the program Non-linear Least Squares Regression of S-PLUS (S-PLUS 2000 Professional Release 1®; MathSoft, Inc., Cambridge, MA). The differences between the variable values (IC50 and nH) of two independent data sets were entered into the common model as shift variables ΔIC50 and ΔnH, activated for all data of the second data set. The corresponding (asymptotic) t-value was used to test the null hypothesis of no variable difference. A one-sample t-test was applied to analyze significance of blocking effects with respect to the starting value. To avoid problems related to multiple hypotheses testing, we applied the method of multiple comparisons with a priori ordered hypotheses (16). The test is based on the assumption that, if the null hypothesis is rejected, there is a positive monotonic relation between concentration and effect. As a consequence, the hypotheses to be tested could be ordered in advance, starting with the largest concentration. If the test results differed significantly from the control data, the effects of the next, smaller concentration were evaluated. The evaluation was stopped as soon as the first insignificant result was obtained. The advantage of this procedure, compared with other approaches to multiple testing, is that the level of type-I error is kept at α = 0.05 for each statistical test. The null hypothesis was rejected when the P value was <0.05.

Results

A total of 60 cells were included in the study. Average currents in the control experiments after depolarization from −100 mV to 0 mV were 4.03 ± 2.2 nA. Maximal inward currents elicited by 10-ms pulses from −150 mV, −100 mV, or −70 mV to 0 mV were reversibly suppressed by propofol in a concentration-dependent manner. Normalized currents (with respect to control) derived from at least six different experiments for each drug concentration were averaged to establish concentration-response plots (see Fig. 1). Significant blocking effects were detected at concentrations ≥10 μM at −70-mV and ≥50 μM at −100- and −150-mV holding potential (P < 0.001). Hill fits to the data yielded IC50 values of 23 μM at −70-mV, 87 μM at −100-mV, and 94 μM at −150-mV holding potential. The degree of suppression was significantly different at −70 mV compared with −100 and −150 mV (P < 0.001). Calculated values for the Hill coefficients nH were 1.1 (−70 mV), 2.7 (−100 mV), and 3.0 (−150 mV). The difference in stoichiometry of drug binding at −70-mV holding potential compared with −100 and −150 mV was significant (P < 0.001).

Figure 1
Figure 1:
Concentration-dependent reduction in test-pulse current with respect to control (Itest/Icontrol, mean ± sd). The data were derived from at least six different experiments for each concentration tested. Depolarizing pulses to 0 mV (10-ms duration) were started from −150, −100, or −70 mV. Solid lines are Hill fits to the data. The concentration-response plots at −100 and −150 mV are nearly superimposable (empty triangles and filled rhombi), whereas blocking potency of propofol was significantly increased when depolarizations were started from −70 mV (filled squares).

After propofol application at concentrations <150 μM, currents reached 89% ± 22% of control values after 2 min of washout.

The increase in blocking potency at a holding potential of −70 mV, where part of the channels are inactivated, compared with −150 mV, where all channels are expected to be in the resting state, suggests that the amount of block achieved by propofol depends on the membrane potential and is increased with increased fraction of inactivated channels with respect to resting channels. Voltage dependence of block and affinity for the inactivated state was further assessed by applying a double-pulse protocol. After brief depolarizations, Na+ channels enter a fast-inactivated state, from which they cannot readily reopen. Currents elicited by test pulses (Itest) starting from varying prepulse potentials (from −150 to −5 mV), normalized to the current elicited at the most hyperpolarized prepotential (−150 mV), represent the relative fraction of channels that have not been inactivated during the 50-ms inactivating prepulse. Boltzmann fits to the resulting current-voltage plots yield the membrane potential at half-maximum channel availability (V0.5) and the slope factor k: I/Imax = (1 + exp([Vtest − V0.5]/k)1. In control conditions, the variables of the Boltzmann fits reflect the voltage dependence of the distribution between resting and fast-inactivated channels.

Summarized control data (n = 45) showed that half of the channels were unavailable at −57.5 ± 4.3 mV because of fast inactivation. The slope factor k was 7.4 ± 1.1. With exposure to propofol, V0.5 was shifted considerably in the direction of more negative prepulse potentials; the degree of alteration showed concentration dependence. The slope factors k remained unchanged in the presence of drug (see Fig. 2). The drug-induced hyperpolarizing shifts reflect an additional reduction of channel availability induced by propofol in the voltage range of channel inactivation compared with −150 mV. Drug binding apparently reached steady state during the 50-ms prepulse at each holding potential, because a prolongation of the prepulse duration to 100 ms did not enhance propofol effects. For comparison, the negative shift in the midpoints induced by 100 μM of propofol was −20.5 ± 3.5 mV when the prepulse duration was 50 ms and −20.0 ± 4.3 mV when the prepulse duration was 100 ms (n = 3).

Figure 2
Figure 2:
Propofol effects on fast-inactivated channels assessed by shifts in the steady-state availability curve. A, Steady-state availability curves assessed by a two-pulse protocol in the absence (control, circles, and washout, rhombi) and presence of 50 μM of propofol (triangles). Each symbol represents the mean fractional current (n > 4 for each concentration) elicited by a 4-ms test pulse to 0 mV, after a 50-ms inactivating prepulse from −150 mV to the indicated prepulse potential. Currents were normalized to maximal value (in each series at −150 mV prepotential); solid lines represent the best Boltzmann [Inorm = {1 + exp(]V]test − V0.5]/k)} 1] fit to the data yielding the membrane potential at half-maximum channel availability (V0.5) and the slope factor k. Error bars are standard deviations. In the presence of propofol, currents were normalized either to maximal value in the presence of drug (filled triangles) or to maximal value in the controls (empty triangles). Propofol shifted the midpoints of the curves in the direction of more negative prepulse potentials, reflecting an additional reduction in channel availability at depolarized prepotentials (compared with the block achieved at −150 mV). B, Concentration dependence of drug-induced negative shifts in the midpoints (ΔV0.5 [mV], mean ± sd) of the steady-state availability plots relative to the starting values. The solid line is a least-squares fits to the equation ΔV0.5 = k · ln{(1 + [C]/IC50 150) × (1 + [C]/Kd) 1}.

To estimate the dissociation constant (Kd) of propofol for the fast-inactivated state of the channel, we used a model developed by Bean et al. (17) for the example of lidocaine effects on Purkinje fibers. The model is based on the assumption that the higher amount of channel block achieved with consecutive membrane depolarization, revealed by the drug-induced hyperpolarizing shift, is determined by the apportionment of channels between resting and fast-inactivated states as well as the different binding affinities for the two channel states. The concentration dependence of the shift in the midpoint was well described by the equation ΔV0.5 = k · ln{(1 + [C]/IC50150) × (1 + [C]/Kd)1} where ΔV0.5 is the shift in the midpoint in each concentration of propofol (mean, n > 4), k is the mean value for the slope factor derived from Boltzmann fits to the current-voltage plots (see above), [C] the applied concentration of propofol, IC50150 the concentration for half-maximum effect derived from the concentration-response plots at −150-mV membrane potential described above, and Kd the dissociation constant for propofol from the inactivated state (see Fig. 2B). The estimated value of Kd was 4.6 μM.

Drug binding to resting or fast-inactivated channels does not reflect all aspects of drug effects, especially in pathological conditions, in which membrane potential might be depolarized for longer periods. Prolonged depolarization induces a kinetically distinct slow-inactivated state that requires much longer periods for recovery (over 1 s) (18). To examine drug binding to slow-inactivated channels, we applied a protocol that induces a reproducible fraction of slow-inactivated channels in control conditions (15). To induce slow inactivation, we stepped up the holding potential from −100 to −35 mV for 2.5 s. The membrane potential was then returned to −100 mV for 10 ms, allowing recovery from fast inactivation. The availability of channels that were not slow-inactivated was then assessed by a 40-ms test pulse to 0 mV. In control solution, the inactivating prepulse caused a reduction of the current elicited by the test pulse by 32.7% ± 2% because of slow inactivation during the long conditioning prepulse, from which channels did not recover during the 10-ms repolarization. Propofol blocked the peak current amplitude in the presence of 32% slow inactivation (with respect to the current amplitude obtained with the same protocol in control solution) at concentrations ≥5 μM (P < 0.001) and block was almost complete at concentrations close to the IC50 for rest block at −100 mV (see Fig. 3). Hill fits to the concentration-response plots revealed a significant increase in potency reducing the IC50 to 22.2 μM when 2.5-s inactivating pulses were applied before the test pulses (P < 0.001).

Figure 3
Figure 3:
A, Concentration-dependent reduction in test pulse current with (filled squares) or without (empty triangles) at 2.5-s inactivating prepulse (n > 5, mean ± sd). Currents were normalized to the current elicited with the same protocol in the controls. The solid lines are Hill fits to the data. The 2.5-s prepulse enhanced sensitivity to propofol. In the presence of slow inactivation, 5 μM of propofol produced significant effects and block was almost complete at concentrations close to the IC50 for rest block. B, Representative current traces elicited by test pulses to 0 mV, after 2.5-s inactivating prepulses in the absence and presence of 10 μM of propofol. With this protocol, the peak current amplitude during the test pulse is reversibly suppressed by 10 μM of propofol.

Discussion

The main finding of our study is that propofol blocks voltage-gated skeletal muscle sodium channels in a concentration-dependent manner in resting, fast-inactivated, and slow-inactivated states. The blocking potency of propofol strongly depends on the kinetic state of the channel. The affinity of propofol for the resting state assessed at an extremely hyperpolarized holding potential (−150 mV) was more than one order of magnitude below the estimated affinity for the fast-inactivated state. As a consequence, blocking effects of propofol are determined by the fraction of inactivated channels, which in turn depends on the respective holding potential. When the holding potential was −70 mV, close to the resting potential of frog skeletal muscle at an external potassium concentration of 5 mM (14), the IC50 value of the blocking effect was 23 μM of propofol, the threshold concentration for significant effects being 10 μM.

In native tissues, drug-free sodium channels can occupy at least two inactivated conformations that are kinetically distinct (18). Brief depolarizations induce fast inactivation, from which recovery at hyperpolarized membrane potentials is rapid (<10 ms). Prolonged depolarizations induce slow inactivation. Slow-inactivated channels require long repriming periods at hyperpolarized membrane potentials to regain availability (18) and may modulate excitability in response to slow shifts in the resting potential (19). When channels were depolarized for a longer period inducing about 30% slow inactivation, propofol achieved significant block of the residual current at concentrations as small as 5 μM. This finding might gain particular importance in pathological conditions, such as myotonia, hypoxia, or ischemia, in which the muscle membrane is depolarized for longer periods (20).

The Hill coefficient derived from the concentration-response plots obtained with different protocols was about 1 in the presence of either fast or slow inactivation and >2, when blocking effects were assessed on resting channels. A Hill coefficient >1 (1.9 ± 0.1) was equally reported for concentration-dependent effects of propofol on central nervous sodium channels when depolarizations were started from −120 mV (6). These Hill coefficient estimates suggest that binding itself shows cooperativity [higher affinity for the second binding site than for the first (21)] at hyperpolarized membrane potentials, indicating that propofol might stabilize a conformational state reached from the resting state that favors binding of more drug molecules. However, this point requires further investigation on channel gating in the presence of propofol, analogous to the experiments performed for lidocaine, by using a conformational marker for channel inactivation (22).

Channel gating kinetics in voltage-operated sodium channels from different tissues are similar concerning activation and inactivation time constants, but they differ markedly in the voltage dependence of these processes (23). Differences in the voltage dependence of inactivation between channel isoforms determine sensitivity to blocking drugs with different binding affinities for resting or inactivated channel states at a given membrane potential (24,25). Propofol equally blocks sodium currents from other tissues. The IC50 values found for the neuronal sodium channels were slightly smaller than those found for the skeletal muscle sodium channels in our preparation (6). However, it should be noted that, in the latter case, each concentration-response plot was obtained from the same cell, whereas we exposed each cell only to one concentration of propofol. These differences in the experimental protocol might account for the small differences in potency seen between the two isoforms. In analogy to the results we obtained for the skeletal muscle channels, the half-maximum blocking concentrations found for propofol effects on cardiac sodium channels were voltage dependent, ranging from 240 μM at hyperpolarized membrane potentials (−140 mV) to 15 μM at −90 mV; the Hill coefficient was kept at 1.5 in both cases (7,26). The difference in blocking potency at −90 mV/−100 mV between the cardiac and the skeletal muscle sodium channel (15 μM at −90 mV compared with 87 μM at −100 mV in our study) is mainly explained by the difference in steady-state voltage dependence of inactivation between the isoforms. The membrane potential of half-maximum channel inactivation was −57 mV for the skeletal muscle and −87 mV for the cardiac isoform (7); thus, at −90 mV, approximately half of the cardiac sodium channels are in the inactivated state, showing higher affinity for propofol, whereas most skeletal muscle sodium channels are in the resting state at −100 mV.

When considering a possible clinical relevance of observations obtained at the molecular level, several complicating factors have to be taken into account. Despite the lack of the β-subunit, the suitability of our preparation for pharmacological studies has been confirmed by several investigators. The α-subunit is the primary, pore-forming subunit of the channel and functions as an ion channel when expressed alone (27). α-Subunits of human skeletal muscle sodium channels show normal gating characteristics (with respect to experiments in native tissue) and retain sensitivity to tetrodotoxin and μ-conotoxin when expressed in a mammalian cell line (28). In addition, the receptor sites for all pharmacological agents interfering with voltage-gated sodium channels are located on the α-subunit (27). However, other factors that might modify channel function and pharmacological effects in vivo, such as coexpression of the β-subunit, posttranslational modification, or phosphorylation of channel proteins (27), are beyond the scope of that model. In addition, free plasma concentrations of propofol may not be identical to the effect-site concentrations in the membrane reached in vivo. Plasma concentrations of propofol during anesthesia required to suppress somatic or hemodynamic responses to different surgical stimuli in 50% of patients range between 13 and 19 μg/mL (70–106 μM) (29). As the protein-bound fraction of propofol is 98%(30), the free propofol concentrations are much smaller than the plasma concentrations. In aqueous solution, the propofol concentration producing loss of righting reflex in Rana pipiens tadpoles (a standard for estimation of anesthetic potency), ranged between 1 and 10 μM (31).

Thus, in conditions in which a normal resting potential of muscle is maintained, a relevant degree of muscle relaxation because of blockade of muscle sodium channels should only be expected at relatively large tissue concentrations of propofol. These results are consistent with a clinical study showing that spontaneous dystonic movements during induction of anesthesia with propofol, attributed to a transient stimulation of deep brain structures, are reduced in magnitude and duration at larger induction doses of propofol (5 mg/kg), although depth of anesthesia monitored with electroencephalogram was equal. The authors postulated an additional, yet unexplained, “muscle relaxing” effect achieved by larger tissue concentrations of propofol (32).

In pathological conditions, such as myotonia, hypoxia or ischemia, in which the normal resting potential cannot be maintained and the muscle membrane is more depolarized, increasing the fraction of inactivated channels (20), sensitivity to propofol should be increased. Thus, under pathological conditions, relevant blocking effects of propofol are likely to occur at small propofol concentrations. In addition, the high oil solubility of the drug [octanol/water partition coefficient of 4300 (31)] suggests that tissue concentrations, especially after long-term therapy, might exceed free plasma concentrations by far. This aspect gains importance when propofol is used for long-term sedation in intensive care units (33).

In conclusion, the results of these in vitro experiments show that voltage-gated sodium channels of the sarcolemma are targeted and blocked in a concentration-dependent manner by propofol. In conditions of normal membrane potentials, a relevant contribution of skeletal muscle sodium channel blockade to the reduction in muscle tone achieved by propofol in vivo might be expected at larger concentrations (10 μM). Because of the increase in blocking potency with consecutive membrane depolarization, pronounced effects might be expected in pathological conditions, in which the normal resting potential cannot be maintained.

We are indebted to Prof. Lehmann-Horn, Ulm, for providing us with transfected cells, and to Prof. Bernd Urban, Bonn, and Prof. Werner Vogel, Gieβen, for helpful discussions. We thank Birgitt Nentwig, Department of Anesthesiology, Hannover, for taking care of the cell culture, Dr. Horst Rückoldt and Dr. Burkhard Vangerow, Department of Anesthesiology, Hannover, for help with software problems, and Jobst Kilian, Department of Neurology, Hannover, for technical support.

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