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These changes resulted in currents that more closely resembled those obtained from intact muscle fibers. A major difficulty in modeling action potentials is deciding values to use for the numerous other parameters in the model.

Previous estimates of maximal sodium conductance from muscle fibers in vivo vary by up to fivefold. In our initial simulation of action potentials we used a value for specific membrane resistance based on our previous measurements Rich et al.

Although myotonia was prevented by use of a low specific membrane resistance Fig. Choosing values for the voltage dependence of slow inactivation are also problematic since values for Nav1.

The threshold for action potential initiation was very close to the membrane potential, and both myotonia and anode break excitation were observed Fig.

The modeling studies above show that shifts in the voltage dependence of sodium channel activation and fast inactivation are needed to accurately simulate muscle fiber excitability during depolarization of the resting potential.

If such shifts occur in muscle fibers in vivo, they may be of crucial importance for regulating excitability.

Holding potential—induced changes in the voltage dependence of activation and fast inactivation in skeletal muscle have been noted during studies of slow inactivation of sodium currents from rat muscles Simoncini and Stuhmer, ; Ruff et al.

However, the significance of the effect was unclear and because increased extracellular calcium reduced the effect, subsequent studies were performed in high extracellular calcium Simoncini and Stuhmer, ; Ruff et al.

Our modeling studies suggest that this effect might be necessary to appropriately regulate muscle fiber excitability, so we performed further studies using normal extracellular calcium levels.

To study the effects of holding potential on the voltage dependence of inactivation, we held fibers for up to 30 min at various holding potentials.

At each holding potential, we measured the midpoint of the voltage dependence of fast inactivation every 1 to 2 min using the voltage protocol shown in Fig.

We found that changing the holding potential caused a shift in the voltage dependence of fast inactivation Fig. A similar effect was observed for the voltage dependence of activation Fig.

Thus, data from muscle fibers studied in vivo support the prediction from our modeling study that the voltage dependence of sodium channel gating is modulated by resting potential.

The voltage dependence of inactivation and activation at various holding potentials. On the right is the plot of the voltage dependence of activation over the same range of holding potentials.

As the holding potential becomes more depolarized, the voltage dependence of both inactivation and activation shift to more depolarized potentials.

The voltage dependence of shifts in the midpoint of sodium channel gating. Shown on the left is the plot of the midpoint of fast inactivation versus holding potential.

On the right is the plot of the midpoint of activation versus holding potential. For the number of fibers at each holding potential refer to Table II.

Changes in the voltage dependence of the midpoint of activation K m act and fast inactivation K m inact for sodium channels in muscle fibers at various holding potentials V hold.

During the course of experiments, we noticed that different test potential amplitudes Fig. Similarly, changing the prepulse amplitude Fig. This was unexpected since evidence presented above suggested that shifts in the voltage dependence of activation and inactivation take minutes to occur.

Previously, a dependence of inactivation on the test pulse amplitude was observed during tight seal recording from cardiac myocytes, and the authors suggested this might be due to the presence of populations of sodium channels with differences in the voltage dependence of activation and inactivation Kimitsuki et al.

To further characterize these effects in skeletal muscle we determined the relationships between prepulse and test pulse amplitudes and the voltage dependence of activation and inactivation.

As shown in Fig. Similarly, as more depolarized prepulses were used, the measured voltage dependence of activation shifted toward more depolarized potentials Fig.

These results demonstrate that the measured voltage dependence of activation and inactivation is directly related to the amplitude of prepulses and test pulses used during the protocols.

Evidence presented above indicates that changes in voltage dependence require minutes to occur after changes in holding potential. Thus, the results shown in Fig.

The measured voltage dependence of sodium channel inactivation and activation depends on the pulse protocol used.

A Examples of two pulse protocols used to measure the voltage dependence of fast inactivation. In protocol 1, a depolarized test potential is used that activates all sodium channels.

Thus when prepulses are altered to measure the voltage dependence of fast inactivation, the measured voltage dependence of fast inactivation is affected by all Nav1.

In protocol 2, a more hyperpolarized activation pulse is used that only activates channels that gate at hyperpolarized potentials.

In protocol 2, the voltage dependence of inactivation will preferentially measure the voltage dependence of inactivation of sodium channels that have a hyperpolarized voltage dependence of activation.

B Examples of two pulse protocols used to measure the voltage dependence of activation. In protocol 1, a depolarized prepulse preferentially inactivates sodium channels that inactivate at hyperpolarized potentials.

Thus protocol 1 will preferentially measure the voltage dependence of activation of sodium channels that have a depolarized voltage dependence of inactivation.

Protocol 2 will not inactivate any sodium channels so that the voltage dependence of activation includes both sodium channels that inactivate at hyperpolarized potentials and sodium channels that inactivate at depolarized potentials.

Use of more depolarized test pulses yielded more depolarized midpoints of inactivation. As the prepulse becomes increasingly depolarized, the measured voltage dependence of activation shifts toward depolarized potentials.

To study sodium channel gating in skeletal muscle in vivo, we used the loose patch technique to acquire the data presented above.

Loose patch studies have the advantage that they allow study of sodium channel gating in vivo with minimal perturbation of the muscle fiber.

However, loose patch studies in which the holding potential is altered are complicated by changes in the amplitude of the sodium current over time due to changes in the fraction of sodium channels that are slow inactivated.

Since the interior of the muscle fiber is not clamped during loose patch measurement of currents, large inward sodium current can cause depolarization of the fiber and underestimation of the voltage step.

Measures of the voltage dependence of activation, however, could be affected by lack of voltage control of the muscle fiber interior.

To determine whether depolarization of the muscle fiber was shifting our measurements of activation midpoint, we examined the effect of sodium current amplitude on the measured voltage dependence of sodium channel activation.

If an absence of interior voltage control was a contributing factor, we reasoned that patches in which currents were larger should have a more negative voltage dependence of activation.

It thus appears unlikely that lack of voltage control of the muscle fiber interior affected our measures of the voltage dependence of activation.

Holding potential—dependent shifts in sodium channel gating are not due to artifacts of the loose patch technique. There is no correlation between the amplitude of the current and the midpoint of inactivation.

The mean of the data shown is presented in Table II. For data included in Table II the patch with the maximal inward current of 55 nA was discarded.

All recordings were performed in solution containing 12 mM potassium. In this group of patches, the average current was larger in cells held at the more depolarized holding potential.

Thus, the shift in the voltage dependence of activation cannot be accounted for by lack of voltage clamp of the interior of the muscle fiber.

Another potential artifact during loose patch studies involves channels that are under poor voltage control near the edge of the pipette Stuhmer et al.

Sodium channels under poor voltage control near the edge of the pipette can activate during depolarizing steps and thus complicate measures of the voltage dependence of activation.

This is particularly problematic during the most depolarizing voltage steps. This caused slow inactivation of sodium channels along the length of the fiber.

By holding patches at various potentials hyperpolarized to the resting potential, slow inactivation was selectively relieved in channels under good voltage control.

Under these conditions, holding potential—induced shifts in the voltage dependence of activation were still observed Fig.

We report here that altering the holding potential of skeletal muscle membrane for prolonged periods induces a shift in the voltage dependence of sodium channel activation and fast inactivation.

Experiments in which the amplitude of prepulses or test potentials was altered suggest that a distribution may exist in the voltage dependence of gating of Nav1.

Based on these results, we propose that holding potential—induced shifts in the voltage dependence of sodium channel gating are due to shifts in the distribution of voltage dependences of Nav1.

Computer simulation of action potentials suggests that a resting potential—induced shift in the voltage dependence of sodium channel gating is necessary to appropriately regulate muscle excitability.

We used loose patch recording methods to study the voltage dependence of sodium channel gating. One concern is that the shift in the voltage dependence of sodium channel gating that we report is due to an artifact of loose patch recording.

This concern is heightened by the fact that a holding potential—induced shift in the voltage dependence of sodium channel gating has not been widely noted despite numerous studies of sodium channel gating using tight seal patch recording.

One potential explanation for why holding potential—induced shifts in gating have not been more widely reported is that the effect occurs very slowly and thus requires prolonged changes in holding potential.

In most studies of sodium channel gating behavior, a single holding potential is used throughout the experiment so that any changes due to alterations of holding potential will be missed.

Furthermore, many recordings of sodium currents are performed in the whole cell mode or use two-electrode voltage clamp in oocytes.

In both situations, factors that regulate sodium channel gating in vivo may be absent. In the whole cell recording mode, as the intracellular milieu is washed out, shifts in the voltage dependence of Nav1.

When the results of loose patch recording of cardiac sodium currents have been directly compared with the results of tight seal patch recording, it appears that the loose patch technique avoids shifts in sodium channel gating induced by formation of the tight seal Eickhorn et al.

Thus, one advantage of loose patch recording is that it avoids some of the artifacts associated with the formation of a tight seal and thus allows for measurement of sodium channel gating with minimal perturbation of the cell.

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