Chapter 42 Skeletal Muscle Excitability

The skeletal muscle fibers are long, being formed by the fusion of myoblast cells and thereby producing a multinucleated long myotube or fiber. The fast-rising AP propagates at a velocity of about 5 m/s. Each skeletal muscle twitch fiber is normally closely controlled by the motor innervation, there being one or two motor end-plates (neuromuscular junctions) located near the midregion of each fiber. Excitation spreads in both directions from the neuromuscular junction.

The twitch fibers undergo developmental changes similar to those in cardiac muscle and neurons. In early development, there are few or no fast Na⁺ channels and the AP upstroke is slow and produced by an inward current through slow Ca²⁺ channels. The AP duration is also long because the delayed rectifier K⁺ conductance is not fully developed. During subsequent development, fast Na⁺ channels are gained, the RP increases (becomes more negative) and the AP shortens to a brief spike.

The skeletal muscle AP spike is immediately followed by a large and prominent early depolarizing afterpotential that slowly decays over 10–20 ms. This early afterpotential is caused in part by the persistence and slow decay of the delayed rectifier K⁺ conductance (that was turned on by the Na⁺ influx-caused depolarization), which has a Na⁺:K⁺ selectivity or P_Na/P_K ratio higher (e.g. 1/30) than that of the resting membrane (e.g. 1/100). This delayed rectifier K⁺ conductance holds E_m for a time more depolarized than the normal RP.

After (and during) a tetanic burst (train) of AP spikes, a large prominent late depolarizing afterpotential is produced. This late afterpotential is caused by K⁺ accumulation in the T-tubules that acts to depolarize them due to the decrease in E_K. Thereby, the surface sarcolemma is depolarized passively. In addition, a slow component of the delayed rectifier K⁺ conductance (less selective for K⁺ than the resting conductance) may persist during the train.

The AP invades into the T-tubules and propagates inward at a slower velocity of about 7–10 cm/s (Gonzalez-Serratos et al., 1978). At this velocity, it would take about 1.0–1.5 ms to propagate to the center of a myofiber having a radius of 30–40 μm. This serves to bring excitation deep into the fiber interior quickly. The depolarization of the T-tubules activates slow (L-type) Ca²⁺ channels located in them and this serves as a critical step in excitation–contraction (EC) coupling. The chapter on EC coupling provides a detailed discussion of a mechanism that involves the Ca²⁺ channels acting as voltage sensors that are coupled to and open the Ca²⁺ release channels in the TC-SR (surface facing the T-tubule).

Some skeletal muscles also contain a fraction of fibers that are non-twitch slow muscle fibers, which normally do not fire APs. They are multiply innervated by the motor neuron, with numerous motor end-plates spaced about 1 mm apart along the entire length of the fiber. Graded contraction of each fiber is produced by varying the frequency of axon APs that increase the amplitude of the end-plate potentials (EPPs) by temporal summation. The membrane potential change produced by the summed EPPs is carried passively into the T-tubules to bring about contraction.

II Introduction

The normal contraction of vertebrate twitch-type skeletal muscle fibers is always preceded by an AP. The AP depolarizes the sarcolemma beyond the membrane potential (E_m) level at which contraction is triggered, i.e. the mechanical threshold (see the subsequent chapter on E-C coupling). This is the first step in the chain of events triggered by the initial excitatory process in the sarcolemma, linking it to the final mechanical response. This chain of events is known as excitation–contraction (E-C) coupling. In this chapter, we will study the sarcolemmal electrophysiological properties that are the basis of the AP generation. AP generation and excitability in neurons were covered in an earlier chapter. Most of the general electrophysiological principles discussed there also apply to skeletal muscle fibers and so are only briefly reviewed and summarized in this chapter. In most respects, the electrogenesis of the APs in nerve axons and skeletal muscle fibers is quite similar. Both are long fibers and have very brief and fast-rising APs whose inward current is carried by Na⁺ ions through fast Na⁺ channels. Skeletal muscle fibers, however, have the added complexity of an extensive internal transverse (T) tubular system formed by a periodic invagination of the surface cell membrane (Fig. 42.1), forming an orderly three-dimensional array of tubules that propagate excitation from the cell surface into the deep interior of the fiber for purposes of E-C coupling. In addition, electrogenesis in skeletal muscle fibers differs from nerve fibers in that (1) different types of afterpotentials are produced, (2) propagation is continuous (i.e. saltatory propagation does not occur) and (3) the membrane has a high Cl⁻ conductance.

FIGURE 42.1 Intracellular and extracellular ion distributions in vertebrate skeletal muscle fibers. Also shown are the polarity and magnitude of the resting potential (RP). Arrows indicate direction of the net electrochemical gradient. The Na⁺-K⁺ pump and Ca²⁺-Na⁺ exchange carrier are located in the cell surface and in the T-tubule membranes. A calmodulin-dependent Ca²⁺-ATPase and Ca²⁺ pump, similar to that in the sarcoplasmic reticulum (SR), is located in the cell surface and T-tubule membranes.

The skeletal muscle AP is considerably different from the AP of cardiac muscle cells which have a very long duration with a pronounced plateau and a substantially lower rate of rise and propagation velocity. The myocardial cells are short and there is a slight delay in propagation at each cell-to-cell junction. Like skeletal muscle fibers, myocardial cells have a fast I_Na responsible for the rapid upstroke of the AP, but the delayed rectifier K⁺ conductance is turned on slowly and there is a substantial inward I_Ca during the entire plateau.

The APs of smooth muscle cells (SMCs) also are markedly different from those of skeletal muscle fibers, in that the RP (takeoff potential) is lower (more depolarized), the rate of rise of the AP is much slower, the AP overshoot is much less and the AP duration (APD) is considerably longer. Propagation velocity is much slower and the SMCs are short and small in diameter. The inward current for the APs in SMCs is primarily a slow Ca²⁺ current carried through L-type Ca²⁺ channels, but some cells do possess some functioning fast Na⁺ channels.

III General Overview of Electrogenesis of the Action Potential

The ion distributions and ion pumps and exchangers found in skeletal muscle fibers are similar to those of other types of cells, as described in the chapters on the RP and nerve APs (see Fig. 42.1). The APs in vertebrate skeletal muscle twitch fibers consist of a spike followed by a depolarizing (“negative”) afterpotential (Figs. 42.2 and 42.3). A large fast inward Na⁺ current, passing through voltage-dependent fast Na⁺ channels, is responsible for electrogenesis of the spike depolarization, which rises rapidly (500–700 V/s). Subsequently, a small slow inward Ca²⁺ current, passing through kinetically slow channels, may be involved in E-C coupling. Outward currents passing through K⁺ channels and Cl⁻ channels are responsible for repolarization of the AP.

FIGURE 42.2 Representation of the electrochemical driving forces for Na⁺, Ca²⁺, K⁺ and Cl⁻ at rest (left diagram) and during the AP in a skeletal muscle fiber (right diagram). Equilibrium potentials for each ion (e.g. E_Na) are positioned vertically according to their magnitude and sign; they were calculated from the Nernst equation for a given set of extracellular and intracellular ion concentrations. Measured RP is assumed to be −80 mV. Electrochemical driving force for an ion is the difference between its equilibrium potential (E_i) and the membrane potential (E_m), i.e. (E_m−E_i). Thus, at rest, the driving force for Na⁺ is the difference between E_Na and the resting E_m; if E_Na is +60 mV and resting E_m is −80 mV, the driving force is 140 mV. The driving force is then the algebraic sum of the diffusion force and the electrical force and is represented by the length of the arrows in the diagram. Driving force for Ca²⁺ (about 210 mV) is even greater than that for Na⁺, whereas that for K⁺ is much less (about 18 mV). Direction of the arrows indicates the direction of the net electrochemical driving force, namely, the direction for K⁺ is outward, whereas that for Na⁺ and Ca²⁺ is inward. If Cl⁻ is passively distributed, then for a cell sitting a long time at rest, E_Cl = E_m and there is no net driving force. The driving forces change during the AP, as depicted. The equations for the different ionic currents are given in the upper right-hand portion of the figure. (Adapted from Sperelakis, 1979.)

FIGURE 42.3 (A) Action potential (AP) recorded with an intracellular microelectrode in a skeletal muscle fiber of frog semitendinosus muscle bathed in normal frog Ringer’s solution. Note the prominent depolarizing afterpotential. Shock artifact is at left of spike. (Modified fromSperelakis et al., 1973.) (B) Diagrammatic representation of the relative conductance changes for Na⁺ and K⁺ during an AP. The rising phase of the AP is caused by an increase in g_Na, which brings the E_m toward E_Na. The falling phase of the AP is due to the rise in g_K, the decrease in g_Na, and to an outward Cl⁻ current. The depolarizing afterpotential is explained in part by the fact that the delayed rectifier K⁺ channel is less selective for K⁺ (30:1 over Na⁺) than is the resting channel (100:1) and, in part, by the contribution of the AP traveling down the T-tubular network.

The skeletal muscle cell membrane has at least two types of voltage-dependent K⁺ channels (Fig. 42.4). One type allows K⁺ ions to pass more readily inward than outward, the so-called inward-going rectifier. This channel is responsible for anomalous rectification (i.e. decrease in g_K with depolarization). There is a quick decrease in K⁺ conductance on depolarization and increase in K⁺ conductance with repolarization. The second type of K⁺ channel is similar to the usual K⁺ channel found in nerve membrane (e.g. squid giant axon), the so-called delayed rectifier. Its conductance turns on more slowly than g_Na on depolarization. This channel allows K⁺ to pass readily outward down the electrochemical gradient for K⁺. The activation of this channel produces the large increase in total g_K that helps to terminate the AP (see Fig. 42.3).

FIGURE 42.4 Electrical equivalent circuits for a skeletal muscle fiber cell membrane at rest (A and B) and during excitation (C). (A) Membrane as a parallel resistance-capacitance circuit, the membrane resistance (R_m) being in parallel with the membrane capacitance (C_m). RP (E_m) is represented by an 80 mV battery in series with the membrane resistance, the negative pole facing inward. (B) Membrane resistance is divided into four component parts, one for each of the four major ions of importance: K⁺, Cl⁻, Na⁺ and Ca²⁺. Resistances for these ions (R_K, R_Cl, R_Na and R_Ca) are parallel to one another and represent totally separate and independent pathways for permeation of each ion through the resting membrane. These ion resistances are depicted as their reciprocals, namely, ion conductances (g_K , g_Cl , g_Na and g_Ca). Equilibrium potential for each ion (e.g. E_K), determined solely by the ion distribution in the steady-state and calculated from the Nernst equation, is shown in series with the conductance path for that ion. RP of −80 mV is determined by the equilibrium potentials and by the relative conductances. (C) Equivalent circuit is further expanded to illustrate that, for the voltage-dependent conductances, there are at least two separate K⁺-conductance pathways (labeled here g_K1 and g_K). In series with the K⁺ conductances are rectifiers pointing in the direction of least resistance to current flow. There is one Na⁺ conductance pathway, the kinetically fast Na⁺ conductance (g_Na). In addition, there is a kinetically slow pathway that allows Ca²⁺ to pass through. Arrows drawn through the resistors indicate that the conductances are variable, depending on membrane potential and time. (Adapted from Sperelakis, 1979.)

The AP amplitude is about 120 mV, from an RP of −80 mV in mammalian myofibers (−90 mV in amphibian) to a peak overshoot potential of about +40 mV (see Figs. 42.2 and 42.3). The duration of the AP (at 50% repolarization, or APD₅₀) ranges between 3 and 6 ms, depending on the species and temperature. The threshold potential (V_th) for triggering of the fast Na⁺ channel conductance is about −65 to −55 mV; thus, a critical depolarization of about 25 mV is required to reach V_th. The turn-on of the fast g_Na (fast I_Na) is very rapid (within 0.2 ms) and E_m is brought rapidly toward E_Na (see Figs. 42.2 and 42.3). There is an explosive (positive exponential initially) increase in g_Na, caused by a positive feedback relationship between g_Na and E_m.

From the current versus voltage (I/V) curves, the maximum inward fast Na⁺ current occurs at an E_m of about −20 mV. The current decreases at more depolarized E_m levels because of the diminution in electrochemical driving force as the membrane is further depolarized, even though the conductance remains high. At the reversal potential (E_rev) for the current, the current goes to zero; I_Na then reverses direction with greater depolarization.

As E_m depolarizes, it crosses V_th for slow Ca²⁺ channels (also called L-type Ca²⁺ channels or dihydropyridine receptors), which is about −35 mV. These Ca²⁺ channels are primarily located in the transverse tubules. Turn-on of the Ca²⁺ conductance (g_Ca) and I_Ca is relatively slow and the peak I_Ca is considerably smaller than the peak I_Na. This Ca²⁺ influx is involved in E-C coupling.

The molecular rearrangements involved in activation of the slow Ca²⁺ channels are directly coupled to opening of Ca²⁺-release channels (or ryanodine receptors) in the sarcoplasmic reticulum (SR) membrane. However, the open probability of these channels is very low. Therefore, the resulting increase in Ca²⁺ conductance (g_Ca) and the peak I_Ca is relatively slow and considerably smaller than the peak I_Na. This Ca²⁺ influx is small during a single AP, but can contribute to E-C coupling during repetitive AP firing.

IV Ion Channel Activation and Inactivation

As discussed in the chapter on nerve excitability (see Chapter 19), the fast Na⁺ channels (and the slow Ca²⁺ channels) have a double gating mechanism; one gate is the activation gate (A-gate) and the second gate is the inactivation gate (I-gate). For a channel to be conducting, both the A-gate and I-gate must be open; if either one is closed, the channel is non-conducting. The A-gate is closed at the resting E_m and opens rapidly on depolarization, whereas the I-gate is open at the resting E_m and closes slowly on depolarization. In the Hodgkin–Huxley (1952) analysis, the opening of the A-gate requires simultaneous occupation of three negatively-charged sites by three positively-charged m⁺ particles. Therefore:

(42.1)

where m is the activation variable, h is the inactivation variable and is the maximum conductance. A small gating current (I_g) has been measured that corresponds to the movement of the charged m particles (or rotation of an equivalent dipole). The outward I_g leads into the inward I_Na.

The fast I_Na lasts only for 1–2 ms because of the spontaneous voltage inactivation of the fast Na⁺ channels, i.e. they inactivate quickly, even when the membrane remains depolarized. Inactivation is produced by the voltage-dependent closing of the I-gate. The voltage dependence of inactivation is given by the h_∞ versus E_m curve. The Na⁺ conductance (g_Na) at any time is equal to the maximal value () times m³ h. Therefore, when h = 0, g_Na = 0, and when h = 1.0, g_Na = (if m = 1.0). At the normal RP, h_∞ is nearly 1.0 and diminishes with depolarization, becoming nearly zero at about −30 mV. The maximal rate of rise of the AP (max dV/dt) is directly proportional to the net inward current or I_Na, which is directly proportional to g_Na and can be expressed as:

(42.2)

Therefore, a decrease in h_∞ causes decrease in max dV/dt. Thus, depolarization by any means (e.g. elevated [K+]_o or applied current pulses) decreases max dV/dt, and excitability disappears at about −50 mV.

The slow Ca²⁺ channels behave much the same way as the fast Na⁺ channels with respect to activation and inactivation, with one main difference being the voltage range over which the slow channels activate and inactivate. Slow channels inactivate between −45 mV and −10 mV, compared to −70 and −30 mV for the fast Na⁺ channels. Another major difference is that the slow Ca²⁺ conductance inactivates much more slowly than the fast Na⁺ conductance; i.e. they have a long inactivation time constant (τ_inact). (The h variable for the slow channel is sometimes referred to as the f variable and the m variable as the d variable.) Because slow Ca²⁺ channels are located in the T-tubular system, their function is affected by tubular Ca²⁺ depletion caused by the Ca²⁺ ions that flow into the myoplasm. The recovery process for the slow Ca²⁺ channels is slow compared to 1–2 ms for fast Na⁺ channels.

The K⁺ channel (delayed rectifier) may have only an activation gate, because it does not inactivate quickly. In the Hodgkin–Huxley analysis of squid giant axon, the A-gate opens when four positively-charged n⁺ particles simultaneously occupy four favorable positions (negatively-charged sites). If n is the probability that one site is occupied, then n⁴ is the probability that all four sites are occupied. Therefore,

(42.3)

The fourth power to which n must be raised causes a delay (sigmoidal foot) in turn-on of the K⁺ conductance.

V Slow Delayed Rectifier K⁺ Current

Two types of K⁺ delayed rectifier currents occur in skeletal muscle. A slow I_K was first described by Adrian and co-workers (1970a,b) in voltage-clamped frog sartorius fibers. The slow component of outward I_K reached a maximum in about 3 s and declined with a time constant of about 0.5 s. In voltage-clamped frog toe muscle, Lynch (1978) observed that most fibers had both a slow component and a fast component of the outward I_K (threshold of −55 mV). The voltage dependences of both K⁺ currents were shifted equally in the depolarizing direction by elevated [Ca²⁺]_o or [H⁺]_o, presumably due to altering the net negative outer surface charge of the membrane, thereby hyperpolarizing. Acidosis also increased the rate of turn-on of the slow delayed rectifier. The fast delayed current was relatively selectively blocked by TEA or by a sulfhydryl reagent, whereas the slow delayed current was selectively depressed by a histidine reagent. It was estimated that about 25% of the delayed rectifier channels are in the T-tubular membrane. The functional significance of the slow I_K is unknown, although it may be partly responsible for the late depolarizing afterpotential (see Section VIII).

VI Mechanisms of Repolarization

The skeletal muscle AP is terminated by three processes: turn-on of g_K, turn-off of g_Na and influx of Cl⁻ ions. The turn-on of the V-dependent K⁺ conductance (g_K) (the delayed rectifier) (see Fig. 42.3) acts to bring E_m towards E_K (about −98 mV), since the membrane potential at any time is determined primarily by the ratio of g_Na/g_K. This type of g_K channel is activated by depolarization and turned off by repolarization. Therefore, this g_K channel is self-limiting, in that it turns itself off as the membrane is repolarized by its action.

In addition to the g_K turn-on, turn-off of g_Na occurs (see Fig. 42.3) (contributing to repolarization) for two reasons: (1) spontaneous inactivation of fast Na⁺ channels that had been activated, i.e. closing of their I-gate (inactivation τ of 1–3 ms) and (2) reversible shifting of activated channels directly back to the resting state (deactivation), because of the rapid repolarization that is occurring due to the g_K increase (Fig. 42.5). Theoretically, it would be possible to have an AP that would repolarize (but more slowly) even if there were no g_K mechanism, because the g_Na channels would spontaneously inactivate and so the g_Na /g_K ratio and E_m would be more slowly restored to their original resting values.

FIGURE 42.5 Illustration of the hypothetical states of the fast Na⁺ channel. The three states patterned after the Hodgkin–Huxley view were modified to reflect the fact that there is evidence for three closed states. As depicted, in the most closed state (C₃), all three m gates (or particles) are in the closed configuration. In the mid-closed state (C₂), two m gates are closed and one is open. In the least closed state (C₁), one gate is closed and two are open. In the resting state, the activation gate (A) is closed and the inactivation gate (I) is open: m = 0, h = 1. Depolarization to the threshold activates the channel to the active state, the A-gate opening rapidly and the I-gate still being open: m = 1, h = 1. The activated channel spontaneously inactivates to the inactive state due to closure of the I-gate: m = I, h = 0. The recovery process on repolarization returns the channel from the inactive state back to the resting state, thus making the channel again available for reactivation. Na⁺ ion is depicted as being bound to the outer mouth of the channel and poised for entry down its electrochemical gradient when both gates are open. The reaction between the resting state and the active state is readily reversible and there is some reversibility of the other reactions. The fast Na⁺ channel is blocked by tetrotodoxin (TTX) binding to the outer mouth and plugging it.

In addition, there is an important third factor involved in repolarization of the AP in skeletal muscle: the Cl⁻ current (see Fig. 42.2). The Cl⁻ permeability (P_Cl) and conductance (g_Cl) are very high in skeletal muscle (and are not strongly V-dependent). In fact, P_Cl of the surface membrane is much higher than P_K , the P_Cl/P_K ratio being about 3–7. As discussed in the chapter on RP (see Chapter 9), the Cl⁻ ion is passively distributed, or nearly so, and thus cannot determine the RP. However, net Cl⁻ movements inwards (hyperpolarizing) or outward (depolarizing) do affect E_m transiently until re-equilibration occurs and there is no further net movement. At the RP, there is no net Cl⁻ current (I_Cl), since there is no electrochemical driving force for Cl⁻ (since E_m = E_Cl). However, during the AP depolarization, there is a larger and larger driving force for outward I_Cl (i.e. Cl⁻ influx), since I_Cl = g_Cl (E_m−E_Cl). In other words, the large electric field that was keeping Cl⁻ out (i.e. [Cl⁻]_i << [Cl⁻]_o) diminishes during the AP and so Cl⁻ ion enters the fiber. This Cl⁻ entry is hyperpolarizing and so tends to repolarize the membrane more quickly than would otherwise occur. That is, AP repolarization is sharpened by the Cl⁻ mechanism. (Note that influx of the negatively-charged Cl⁻ ion is an outward Cl⁻ current, which is repolarizing.)

To illustrate further some of the preceding points on the role of Cl⁻, when skeletal muscle fibers are placed into Cl⁻-free Ringer solution (e.g. methanesulfonate substitution), depolarization and spontaneous APs and twitches occur for a few minutes until most or all of the [Cl⁻]_i is washed out. After equilibration, the resting E_m returns to the original value ca. −90 mV for frog skeletal muscle and −80 mV for mammalian, clearly indicating that Cl⁻ does not determine the RP and that net Cl⁻ efflux produces depolarization. Re-addition of Cl⁻ to the bath produces a rapid large hyperpolarization, e.g. to −120 mV, due to net Cl⁻ influx; the E_m then slowly returns to the original value (e.g. −90 mV) as Cl⁻ re-equilibrates, i.e. redistributes itself passively. These same effects occur in cardiac muscle, smooth muscle and nerve, but to a lesser extent, because in these tissues P_Cl is much lower (e.g. P_Cl/P_K ratio is only about 0.5 in vascular smooth muscle).

The importance of the Cl⁻ current in repolarization in skeletal muscle fibers is illustrated by one type of myotonia in which an abnormally low P_Cl causes repetitive APs to occur. Because g_Cl is abnormally low, total membrane conductance G_m is also low. From the relationship between membrane current (I_m) and G_m (I_m /G_m = E_m), it can be deduced that only a smaller outward depolarizing membrane current I_m is necessary to reach threshold E_th for an AP. Since g_Cl is abnormally low, the membrane resistance R_m will be abnormally high, making the space constant λ larger than normal. Because g_Cl is abnormally low, the Cl⁻ influx during AP repolarization is much less than normal and so the repolarization process is slowed, thus increasing the duration of the AP. As a consequence of the above, when depolarization occurs, the AP threshold is easily and quickly reached and the generated APs spread easily along the sarcolemma. These factors make the whole system unstable and oscillations trigger repetitive discharge of APs. That is, the muscle fibers lose their tight control by the motor neurons and so contraction becomes partly involuntary. For example, persons with myotonia find it difficult to release a handshake or to remove their hand from a drinking glass. There are several causes of myotonia, including genetic abnormalities in ion channels, as well as drug-induced conditions. Any agent that greatly lowers P_Cl or g_Cl will have the same effect. It has been shown that simply decreasing g_Cl causes repetitive firing in equivalent circuit models of skeletal muscle fibers. In addition, K⁺ ions tend to accumulate in the lumen of the T-tubules under normal conditions (see Section VIII). This accumulation is exaggerated with the prolonged APs and so tends partially to depolarize the fibers and increase their excitability. Some forms of myotonia are produced by abnormal fast Na⁺ channels; namely, a small fraction of these channels do not inactivate as quickly as usual (i.e. their I-gates do not close normally) and so causes a prolonged small depolarization after the AP and, consequently, a repetitive discharge.

In myotonia, AP repolarization is slowed and the duration of the AP is increased. As AP duration increases, more Na⁺ channels have time to return to the resting conformation by deactivation or recovery from inactivation (a process which has a time constant of 2–3 ms). This creates a window of instability during AP repolarization. The membrane potential remains depolarized above threshold, allowing some Na⁺ channels to reopen and trigger another AP. The skeletal muscles have a large safety factor with respect to the number of Na⁺ channels in the membrane and as few as 3–5% in the open state can trigger an AP. This instability in membrane repolarization is further enhanced by the high membrane resistance. Without the normal large g_Cl, only a smaller than normal outward depolarizing current is required to reach threshold. Because of the high membrane resistance R_m, the space constant λ is longer than normal. Consequently, the APs propagate at a faster velocity. These factors act synergistically to make the whole system unstable. Consequently, when depolarization occurs, the AP threshold is easily and quickly reached, the generated APs spread fast along the sarcolemma and the membrane is more excitable and susceptible to repetitive discharge of APs.

The high g_Cl in skeletal muscle fibers is due to a large number of voltage-dependent gated Cl⁻ channels, which are outwardly rectifying. The major Cl⁻ channel of skeletal muscle is the ClC-1 channel (Steinmeyer et al., 1991), a member of the ClC family of Cl⁻ channels and Cl⁻/H⁺ antiporters (Zifarellli and Pusch, 2007). These Cl⁻ channels are located both on the surface sarcolemma and T-rubule membrane in frogs and mammals. Denervation of mammalian fibers causes g_Cl to decrease almost to zero.

VII ATP-Dependent K⁺ Channels

K_ATP channels are among the most abundantly expressed K⁺ channels in the skeletal muscle sarcolemma, reaching densities of 10 channels per μm² of surface membrane (Spruce et al., 1987), comparable to that of K⁺ delayed rectifier channels (Standen et al., 1985). K_ATP channel currents can be recorded from both surface (Spruce et al., 1985) and transverse tubular membranes (Heiny et al., 1983). However, their physiological role in skeletal muscle is not well understood (reviewed in Flagg et al., 2010). The functional K_ATP channel is an octomeric protein composed of four pore-forming Kir subunits (Kir 6.1 and Kir 6.2) and four regulatory SUR (sulfonylurea receptor) subunits assembled in a 4:4 stoichiometry. Functioning of the K_ATP channel requires the proper coupling between Kir and SUR subunits. More details on the subunits is given in the Appendix to this chapter.

ATP-dependent K⁺ (K_ATP) channels are characterized by the inhibition of channel openings by ATP (Yokoshiki et al., 1998). In addition to being ligand sensitive, these K_ATP channels are voltage-dependent; the open state probability increases with depolarization. Most studies of K_ATP channels have been performed using patch-clamp of isolated inside-out patches of skeletal muscle (Spruce et al., 1987). The unitary conductance of this channel varies with [K⁺]_o, ranging from 15 pS in 2.5 mM [K⁺]_o to 42 pS in 60 mM. K_ATP channels are closed when the ATP concentration in the intracellular myoplasm is in the range of 1.0 mM or higher, which is the normal physiological concentration. A decrease in ATP concentration or depolarization activates the channel. The half-maximum inhibition of this channel opening by ATP, measured at a constant [K⁺]_o/[K⁺]_i, is 0.135 mM at pH 7.2 (Fig. 42.6). Hydrolysis of ATP (into ADP + P_i) is not required for ATP to close K_ATP channels.

FIGURE 42.6 Effect of ATP (open circles), ADP (filled circles) and AMP (open triangles) on closing the ATP-regulated K⁺ channels. Ordinate: open-state probability (P_o) of the channel relative to its value in nucleotide-free solution (P_open control). (Reproduced with permission from Spruce et al., 1987.)

Although ATP is the most specific ligand to close K_ATP channels, there are other ligands that modulate these channels. The metabolites produced during contraction, like ADP, Mg²⁺ and H⁺ also modulate the channels. The two adenine nucleotides, ADP and AMP (in the absence of ATP), also can block K_ATP channels in a dose-dependent manner. However, they are less effective than ATP (see Fig. 42.6). ATP analogs like GTP, ITP, XTP, CTP and UTP also have reduced effectiveness (about tenfold) in closing these channels. ADP shifts the ATP dose–response curve, raising the half-inhibition concentration, consistent with competition between ATP and ADP for the nucleotide binding site on the channel (Spruce et al., 1987; Vivaudou et al., 1991).

A decrease in pH at the cytoplasmic surface (pH_i) reduces the degree of K_ATP channel inhibition caused by ATP. The ATP concentration for half-inhibition is 2.5 times and 15 times greater at pH_i 6.8 and 6.3, respectively, as compared to that at pH_i 7.2. Thus, during acidosis, a smaller decrease in ATP concentration will lead to a larger opening of K_ATP channels. It was proposed that proton binding to the channel prevents ATP binding (Davies et al., 1992). Mg²⁺ has a similar effect as protons. An increase of cytosolic Mg²⁺ reduces the ability of ATP to close K_ATP channels (Vivaudou et al., 1991). Mg²⁺ may bind to the channels and plug them (Woll et al., 1989). In addition, the inhibitory effect may be partly due to the ability of Mg²⁺ to bind to ATP.

K_ATP channels have been studied in excised patches of surface membrane blebs from muscles of frogs and mammals. These channels are, therefore, likely to be present in the sarcolemma. It is not known whether they are present in the T-tubular system. As stated earlier, the K_ATP channel density has been estimated to be as high as 10 channels per μm² of surface membrane (Spruce et al., 1985), a density comparable to that for K⁺ delayed rectifier channels.

The physiological role played by K_ATP channels in skeletal muscle is not clear. Under physiological conditions, the intracellular ATP concentration is about 5 mM at rest. Therefore, at the resting ATP level, almost all of the K_ATP channels should be inactive (Spruce et al., 1987; Davies et al., 1992). Even during repetitive contractions that lead to muscle fatigue, the ATP concentration is maintained at near normal levels by the action of creatine kinase and creatine phosphate (Carlson and Siger, 1960; Nassar-Gentina et al., 1978).

It has been proposed that K_ATP channels may be associated with the decrease in force development underlying muscle fatigue. A drug (SR44866) that opens K_ATP channels (in frog skeletal muscles) also reduces the AP duration, the early afterpotential and the peak twitch force (without affecting the RP) in intact muscles (Sauviat et al., 1991). The high K⁺ permeability found in muscle fibers that have undergone a permanent contracture (rigor) produced by repetitive stimulation when poisoned with cyanide and iodocetate (Fink and Lüttgau, 1976), may be caused by the decreased ATP concentration opening K_ATP channels. In metabolically-exhausted frog semitendinosus muscle fibers, the addition of tolbutamide and glyburide, two K_ATP channel antagonists, significantly reduces the K⁺ efflux rate (Castle and Haylett, 1987). Fatigued muscle fibers can contract when Ca²⁺ is released directly from the intracellular SR stores (Gonzalez-Serratos et al., 1978; Garcia et al., 1991). In intact animals, some of the skeletal muscle fatigue results from synaptic fatigue, including at the neuromuscular junction.

As stated previously, a decrease in pH_i (e.g. from lactic acid production) reduces the inhibitory effect of ATP on K_ATP channels. During exercise, with fatigue development, pH_i may decrease by about one unit (Renaud, 1989). A small decrease in ATP concentration is accompanied by an increase in ADP, H⁺ and Mg²⁺ concentrations and the overall combination of these chemical changes may then lead to the activation of K_ATP channels. As K_ATP channels open, they contribute to the increased K⁺ efflux found during repetitive muscle contraction. Because of the restricted diffusion out of the T-tubular system and the closeness of the intercellular fiber spacing, extracellular K⁺ concentration increases, especially inside the tubular system. This may cause a decrease in cell excitability, which may be reflected as decreased force development. This mechanism may protect skeletal muscle cells from large ATP depletions that would have deleterious effects.

VIII Electrogenesis of Depolarizing Afterpotentials

As mentioned previously, the AP spike in skeletal muscle fibers is followed by a prominent depolarizing afterpotential (also called a negative afterpotential based on the old terminology from external recording) (see Fig. 42.3). In addition to this early depolarizing afterpotential (i.e. emerging from the spike downstroke), there is a late depolarizing afterpotential that follows a tetanic train of spikes (e.g. 10 spikes). The electrogenesis of the early and late afterpotentials is different. The early afterpotential is due to a membrane conductance change, whereas the late afterpotential is due primarily to K⁺ accumulation in the T-tubules.

The early depolarizing afterpotential of frog skeletal fibers is about 25 mV in amplitude immediately after the spike component and gradually decays to the RP in 10–20 ms. This afterpotential results from the fact that the delayed rectifier K⁺ channel that opens during depolarization to terminate the spike is less selective for K⁺ (ca. 30:1, K⁺:Na⁺) than is the K⁺ channel in the resting membrane (ca. 100:1) (Adrian et al.,1970a). Therefore, the constant-field equation predicts that the membrane should be partly depolarized when E_M is dominated by this K⁺ conductance that is turned on during the AP. Thus, the early depolarizing afterpotential is partly due to the persistence and slow decay of this less-selective K⁺ conductance. Adrian and Peachey (1973) were able to reconstruct the time course of the AP and the early depolarizating afterpotential by giving values to the access resistance of the T-tubular system, presence of Na⁺ and K⁺ tubular membrane currents and velocity of the tubular AP. The early depolarizing afterpotential reflects, in part, the tubular AP, as evidenced by the disappearance of the early depolarizing afterpotential in muscles in which the T-tubular system has been disrupted and disconnected from the surface membrane by the glycerol osmotic shock method¹ (Eisenberg and Gage, 1969).

The late depolarizing afterpotential of frog skeletal fibers may result from accumulation of K⁺ ions in the T-tubules (Adrian and Freygang, 1962). During the AP depolarization and turn-on of g_K (delayed rectifier), there is a large driving force for K⁺ efflux from the myoplasm coupled with a large K⁺ conductance, resulting in a large outward K⁺ current [I_K = g_K (E_m−E_K)] across all surfaces of the fiber, namely across the surface sarcolemma and T-tubule walls. The K⁺ efflux at the fiber surface membrane can rapidly diffuse away and mix with the relatively large interstitial fluid (ISF) volume, whereas the K⁺ efflux into the T-tubules (TT) is trapped in this restricted diffusion space. The resulting high [K⁺]_TT decreases E_K across the T-tubule membrane and thereby depolarizes this membrane. Because of cable properties, part of this depolarization is transmitted to the surface sarcolemma and is recorded by an intracellular microelectrode. The K⁺ accumulation in the T-tubules can only be dissipated relatively slowly by diffusion out of the mouth of the T-tubules and by active pumping back into the myoplasm (across the T-tubule wall) by the Na⁺-K⁺ pump sites located in the T-tubular membrane. Thus, the decay of the late depolarizing afterpotential will be a function of these two processes.

The amplitude and duration of the late depolarizing afterpotential is a function of the number of spikes in the train and their frequency. That is, the greater the spike activity, the greater its amplitude and duration. If the train consists of 20 spikes at a frequency of 50/s, a typical value for the amplitude of the late depolarizing afterpotential in frog fibers is about 20 mV. When the diameter of the T-tubules is increased by placing the fibers in hypertonic solutions 2, the amplitude of the late afterpotential decreases as expected because of the greater dilution of the K⁺ ions accumulating in the T-tubule lumen. When the T-tubular system is disrupted and disconnected from the surface membrane by the glycerol osmotic shock method, the late afterpotentials disappear together with the early afterpotentials.

An alternative explanation for the late depolarizing afterpotential is that it may be due to the slow delayed rectifier g_K change described above (Adrian et al., 1970b). The equilibrium potential for the slow I_K is −83 mV and the sign (direction) of the late afterpotential reverses when the fiber is depolarized beyond −80 mV (e.g. to −70 mV). Hence, the late afterpotential could arise from the slow relaxation of a component of the K⁺ conductance increase, which is less selective for K⁺ than the K⁺ channels open in resting membrane. In this view, the electrogenesis of the late afterpotential would be similar to that for the early afterpotential.

All depolarizing afterpotentials, regardless of whether early or late, have physiological importance because they alter excitability and the propagation velocity of the fiber. A depolarizing afterpotential should enhance excitability (lower threshold) to a subsequent AP. This is because the critical depolarization required to reach the threshold potential would be decreased. A large late depolarizing afterpotential, such as that due to K⁺ accumulation in the T-tubules, can, under certain pathological conditions, trigger repetitive APs. The effect of depolarizing afterpotentials on velocity of propagation involves two opposing factors: (1) the decrease in critical depolarization required; and (2) the decrease in maximal rate of rise of the AP (max dV/dt), which is a function of the takeoff potential (h_∞ versus E_m curve). Therefore, what factor dominates will depend on the degree of depolarization and the shape of the h_∞ curve. When frog skeletal fibers are depolarized slightly by elevating [K⁺]_o, only a decrease in propagation velocity is observed (Sperelakis et al., 1970).

IX Ca²⁺-Dependent Slow Action Potentials

Slow APs are recorded under conditions in which the fast Na⁺ current is blocked by Na⁺-deficient solution, tetrodotoxin (TTX) or voltage inactivation of the fast Na⁺ channels in elevated [K⁺]_o. Under these conditions, the only carrier of inward current available to produce an AP is Ca²⁺ ion. Spontaneously-occurring slow APs were first observed in frog sartorius fibers equilibrated in Cl⁻-free solution containing TTX (Sperelakis et al., 1967). Upon addition of Ba²⁺ ion (e.g. 0.5 mM), which is a potent blocker of K⁺ channels and P_K, the fibers partially depolarize and spontaneously discharge slowly-rising (e.g. 1–10 V/s), overshooting APs of long duration (e.g. several seconds), having a prominent plateau component (resembling a cardiac AP in shape). An abrupt repolarization terminates the slow AP. Ba²⁺ depolarizes rapidly in Cl⁻-free solution, because the voltage-clamping effect of the Cl⁻ distribution (E_Cl), due to the large P_Cl , is circumvented. Cl⁻-free solution raises the resistance of the cell membrane about sevenfold.

In a frog skeletal muscle fiber, using two intracellular microelectrodes, one for applying current intracellularly and the other for recording voltage a short distance away in the same fiber ([K⁺]_o of 25 mM to depolarize the fiber to about −45 mV and thereby inactivate the fast Na⁺ channels and [Na⁺]_o reduced to zero so that there could be no inward Na⁺ current), application of small hyperpolarizing current pulses during the slow AP indicated that membrane resistance increases progressively during the plateau (Kerr and Sperelakis, 1982). The rate of rise, overshoot and duration of the slow APs are a function of [Ca²⁺]_o (Beaty and Stefani, 1976; Vogel et al., 1978; Kerr and Sperelakis, 1982). For example, the AP duration at 50% amplitude (APD₅₀) was generally 2–8 s. The amplitude of the slow AP plotted against log [Ca²⁺]_o gave a straight line with a slope of 28 mV/decade, which is close to the theoretical 29 mV/decade (at 21°C) from the Nernst relationship for a situation in which only Ca²⁺ ion carried the inward current. The slow APs were depressed and blocked by the Ca²⁺-antagonistic and slow-channel-blocking drugs, verapamil and bepridil, with an ED₅₀ of about 5×10⁻⁸ M. The slow AP arises from the T-tubular system of the fiber (Vogel et al., 1978; Kerr and Sperelakis, 1982), based on their disappearance when the T-tubules were disrupted and disconnected from the surface membrane by the glycerol osmotic shock method. The normal fast APs are not affected by the glycerol treatment. These results indicate that the slow Ca²⁺ channels giving rise to the slow APs are located primarily in the tubular system.

Isotope flux measurements have shown that there is a net Ca²⁺ influx during contractions of phasic skeletal muscle fibers, suggesting that an influx from the extracellular space may initiate the contraction (Bianchi and Shanes 1959). Additionally, voltage-clamped muscle fibers have slow inward Ca²⁺ currents (I_Ca) (Stanfield, 1977; Sanchez and Stefani, 1978). Elevation of [Ca²⁺]_o increased I_Ca, and I_Ca was depressed by the slow Ca²⁺ channel blockers D-600, nifedipine and Ni²⁺ (Stanfield, 1977; Sanchez and Stefani, 1978; Almers et al., 1981). Detubulation by the glycerol osmotic shock method abolishes I_Ca (Nicola-Siri et al., 1980; Potreau and Raymond, 1980). These results support the conclusion that I_Ca produces the slow APs. The various conformational states that the Ca²⁺ slow channels undergo during excitation are depicted in Fig. 42.7. These states are similar to those of the fast Na⁺ channels (see Fig. 42.5), except there are only two closed states.

FIGURE 42.7 Illustration of the four hypothetical states of the slow Ca²⁺ channel. There is evidence for two closed states. As depicted, in the most closed state (C₂), both d gates (or particles) are in the closed configuration. In the least closed state (C₁), one gate is closed and one is open. In the resting membrane, the activation gate (A) is closed and the inactivation gate (I) is open: d = 0, f = 1. Depolarization to the threshold activates the channel to the active state, the A-gate opening rapidly and the I-gate still being open: d = 1, f = l. The activated channel spontaneously inactivates to the inactive state due to closure of the I-gate: d = 1, f = 0. The recovery process on repolarization returns the channel from the inactive state back to the resting state, thus making the channel again available for reactivation. Ca²⁺ ion is depicted as being bound to the outer mouth of the channel and poised for entry down its electrochemical gradient when both gates are open. The reaction between the resting state and the active state is readily reversible and there is some reversibility in the other reactions. The slow channels behave similarly to the fast channels, except that their gates appear to move more slowly on a population basis; i.e. the slow channels activate and recover more slowly. (Although the gates of any individual slow channel may move quickly, the stochastic behavior of the population of channels is such that their summed conductance changes slowly.) The slow channel gates operate over a different voltage range than the fast channels (i.e. less negative, more depolarized). TTX does not block the slow channels, but drugs such as nifedipine do block by binding to the channel.

Do slow inward calcium currents (I_CaS), play a role in E-C coupling? A substantial contraction, of between 20 and 50% of the normal twitch tension, accompanies the slow APs (Vogel et al., 1978), suggesting that the Ca²⁺ channels in the tubular system may play a role during E-C coupling. However, skeletal muscle fibers contract for several minutes after [Ca²⁺]_o is lowered to 10⁻⁸ M (Armstrong et al., 1972) and the Ca²⁺-channel blocker diltiazem did not depress twitch or tetanic force development (Gonzalez-Serratos et al., 1982). These results suggest that I_Ca may play no role in E-C coupling in normal amphibian muscles. Nevertheless, in dysgenic mice, in which contraction of skeletal muscles is weak, the Ca²⁺ channels in the T-tubules are few or absent.

Ca²⁺ influx during the slow AP could trigger the release of more Ca²⁺ from the nearby TC-SR via the Ca²⁺-trigger Ca²⁺-release mechanism (Fabiato, 1982). Because the time course of the slow AP is much longer than that of a twitch contraction, it was suggested that the inward Ca²⁺ current may play a role in K⁺ contracture, in tetanic contraction, or in long-term regulation of contraction, perhaps by increasing the Ca²⁺ concentration in the SR and thereby increasing the amount of internal Ca²⁺ available for release on subsequent activation (Nicola-Siri et al., 1980). [Ca²⁺]_SR does increase following tetanic stimulation (Gonzalez-Serratos et al., 1982).

Slow APs were also recorded from mouse skeletal muscle fibers equilibrated in a solution that was Cl⁻-free, low Na⁺ (10 mM) and high K⁺ (20 mM) (Kerr and Sperelakis, 1982). As with frog muscle, the slow APs were abolished after detubulation and blocked by verapamil, bepridil, Mn²⁺ and La³⁺. Their rate of rise, amplitude and duration increased as a function of [Ca²⁺]_o, with max dV/dt being about 0.5 V/s in 8 mM.

During the first 5 days in culture, embryonic skeletal muscle cells from Xenopus laevis need extracellular Ca²⁺ to contract when stimulated (in contrast to adult muscles). Thus, an inflow of Ca²⁺ from the extracellular space may be required in embryonic cells as the means to produce contraction. In whole-cell voltage-clamp studies, I_Ca currents have been observed in embryonic and neonatal skeletal muscle cells in culture (Moody-Corbett et al., 1989; Cognard et al., 1992; Gonzalez-Serratos et al., 1996; Cordoba-Rodriguez et al., 1997). The current density increased from 1.7 to 3.3 and to 7.9 pA/pF at 1, 5 and 15 days in culture, respectively. These results indicate that the T-tubules and SR are poorly developed or not functional in early stages of muscle development and that, in early development, I_Ca may be an important mechanism to trigger contraction.

X Developmental Changes in Membrane Properties

The cell membranes of most excitable cells apparently pass through similar stages of differentiation during development. For example, young (2- to 3-day-old) embryonic chick hearts (tubular) have few or no functional fast Na⁺ channels, but have a high density of slow channels (both Na⁺ and Ca²⁺) and fire slowly-rising TTX-insensitive APs. Fast Na⁺ channels then appear and progressively increase in number, reaching the maximal (adult) level at late embryonic development (e.g. day 20). The P_Na/P_K ratio is high in young hearts, due to a low P_K , and accounts for the low RP and automaticity in nearly all the cells.

Skeletal muscle fibers and neurons also undergo developmental changes in membrane electrical properties (e.g. Spector and Prives, 1977; Spitzer, 1979) (see Chapter 25). In general, fast Na⁺ channels are absent in the young, less differentiated cells, but they do possess excitability because of a large number of slow channels. The AP is TTX-insensitive, slowly rising and of long duration (resembling a slow AP in cardiac muscle). Later during development, fast Na⁺ channels make their first appearance and the fast Na⁺ channels and slow channels coexist. During that period, TTX does not abolish the APs, but reduces max dV/dt (i.e. slow APs remain). At a later stage, the slow channels in the sarcolemma are lost (or greatly reduced in number) and the fast Na⁺ channels progressively increase in density. The APs become fast rising and of short duration and are completely abolished by TTX. As discussed previously, some functional slow Ca²⁺ channels remain in the T-tubular system.

XI Electrogenic Na⁺-K⁺ Pump Stimulation

The Na⁺, K⁺-ATPase pump is electrogenic in skeletal muscle fibers (both mammalian and amphibian). The pump produces a net outward current, because three Na⁺ ions are pumped out to every two K⁺ ions pumped in. The electrogenic pump potential contribution to the RP (see Chapter 11 to 9) is very large, about 12–16 mV, in rat skeletal muscle fibers (Sellin and Sperelakis, 1978). The net pump current can be stimulated by increasing the number of pump sites per unit area of cell membrane or by increasing the turnover rate of each pump site. β-Adrenergic agonists (e.g. isoproterenol) rapidly hyperpolarize skeletal muscle fibers by 7–9 mV within 5 min. Insulin has a similar effect, but hyperpolarizes more slowly (e.g. peak reached by 10 min) and to a smaller degree (e.g. 5–7 mV) (Iannaccone et al., 1989). Since cAMP also hyperpolarizes, the action of β-agonists is believed to be mediated by elevation of cAMP and phosphorylation of the Na⁺-K⁺ pump (or an associated regulatory protein) by protein kinase A (PKA). The action of insulin is thought to be mediated by the incorporation of spare membrane from an internal pool, which contains Na⁺-K⁺ pumps, into the cell membrane.

The pump current (I_p) can be directly measured in single fibers (cultured skeletal myotubes rounded by use of colchicine, a microtubule disrupter) by doing whole-cell voltage clamp under conditions in which all the ionic conductances are blocked. When this is done, the pump current can be measured at different voltages and normalized for unit membrane capacitance (hence membrane area). Values of about 1 pA/pF or 1.0 μA/cm² were obtained, with a reversal potential (or zero current) of about −140 mV (Li and Sperelakis, 1994).

When [K⁺]_o is lowered below the normal physiological level, e.g. from 4.5 mM to about 0.1 mM, a large depolarization occurs in mammalian skeletal muscle fibers (Fig. 42.8). This depolarization is caused, in part, by inhibition of the Na⁺-K⁺ pump current. The K_m value for [K⁺]_o for the Na⁺, K⁺-ATPase is about 2 mM and the relationship between Na⁺, K⁺-ATPase activity and [K⁺]_o is very steep. Therefore, inhibition of the Na⁺-K⁺ pump occurs.

FIGURE 42.8 The mean resting membrane potential (E_m) of normal mouse skeletal muscle plotted as a function of the extracellular K⁺ concentration ([K⁺]_o) on a logarithmic scale. The straight line drawn through the data points for 20 mM [K⁺]_o and above has a slope of 50 mV/decade. Extrapolation of this line to zero potential gives the intracellular K⁺ concentration ([K⁺]_i) of 185 mM. The dashed line gives the calculated E_K values (slope of 61 mV/decade). Note the “fold-over” of the E_m curve at [K⁺]_o, levels below 1 mM, presumably due to inhibition of the electrogenic pump potential (V_p) and to a decrease in P_K and g_K at low [K⁺]_o levels. (Reproduced from Sellin and Sperelakis, 1978.)

XII Slow Fibers

One type of skeletal muscle fibers, known as slow fibers, subserves tonic functions, including posture. Slow fibers should not be confused with “slow twitch fibers”. The true slow fibers do not fire APs, whereas all types of twitch fibers do. The slow fibers are usually smaller in diameter than twitch fibers and they exhibit a less distinct myofibrillar arrangement (so-called “felden” structure). Slow fibers have been found in a number of vertebrate muscles, e.g. in the frog rectus abdominus muscle, frog ileofibularis muscle and mammalian extraocular muscles. It is probable that careful searching will reveal some slow fibers in other mammalian muscles as well.

The slow fibers have multiple innervation by a series of motor end-plates (spaced about 1 mm apart), all from a single motor neuron. As with twitch fibers, acetylcholine (ACh) is the synaptic transmitter. The force of contraction of the slow fibers is controlled by graded end-plate potentials (EPPs). That is, an increase in frequency of impulses in the motoneuron produces a larger EPP (by temporal summation) and this, in turn, produces a greater contraction in the vicinity of the end-plate. Since the end-plates are spaced closely together – at a distance of about one length constant – the entire fiber becomes nearly uniformly depolarized, even though there are no propagated APs. Therefore, the entire length of the slow fiber contracts almost uniformly.

The slow fibers do possess T-tubules which abut at the triadic junctions with the terminal cisternae of the SR (TC-SR). Therefore, the T-tubules may act as passive conduits in the slow fibers to bring the depolarization (produced in the surface membrane by the EPP) deep into the fiber interior. Thus, depolarization of the T-tubule occurs by their cable properties. This depolarization, in turn, could bring about the influx of Ca²⁺ by activation of voltage-dependent slow Ca²⁺ channels located in the T-tubules.

APs normally cannot be induced to occur in vertebrate slow fibers under a variety of experimental conditions. However, denervation of frog slow fibers does allow an AP-generating mechanism to appear (Miledi et al., 1971). APs can be induced in slow fibers of invertebrates (e.g. crustacean skeletal muscles) (Fatt and Ginsborg, 1958). Similarly, in the neurogenic horseshoe crab (Limulus) heart, which normally is activated by summating excitatory postsynaptic potentials, propagating (ca. 5 cm/s) and overshooting spontaneous APs can be rapidly induced by Ba²⁺ (0.1–10 mM) (Rulon et al., 1971). These slowly-rising (ca. 1.0 V/s) APs are resistant to TTX and these voltage-dependent slow channels can pass Ba²⁺, Sr²⁺ and Ca²⁺.

XIII Conduction of the Action Potential

When the EPP, generated at the neuromuscular junction, reaches threshold for eliciting an AP in the vertebrate twitch skeletal muscle fiber, an AP is propagated down the muscle fiber in both directions from the end-plate. (In some muscle fibers, there is a second end-plate innervated by a motoneuron exiting the spinal cord at another level.) The AP is overshooting (to about +40 mV) and propagates at a constant velocity of about 5 m/s over the surface sarcolemma. Propagation occurs by means of the local-circuit currents that accompany the impulse, as discussed in Chapter 19. The reader is referred to that chapter for details on the radial (transmembrane) currents and the longitudinal (axial) currents. The external longitudinal currents can use the entire ISF space (since current takes the path of least resistance), allowing the electromyogram (EMG) to be recorded from the skin overlying an activated skeletal muscle. The amplitude of the EMG potentials becomes larger when more fibers within the muscle are activated (fiber summation), because of summation of the IR voltage drops produced by each fiber activated simultaneously. The frequency of the EMG potentials reflects the frequency and asynchrony of activation of the muscle.

The skeletal muscle fibers are formed by myoblast cells that have fused end to end to become long multinucleated myotubes and then cylindrical fibers later in development. They behave as semi-infinite cables. That is, an AP can propagate from one end of the fiber to the other, uniformly and unimpeded. The space constant or length constant (λ) of the fiber cable is about 1.5 mm for frog sartorius fibers (Sperelakis et al., 1967) and about 0.76 mm for the rat EDL muscle (Sellin and Sperelakis, 1978). The length constant is the distance over which a voltage applied at one region would decay to 1/e (1/2.717 = 0.368) or 36.8% of the initial value. That is, in a passive cable, voltage decays exponentially with a certain length constant as given by:

(42.4)

where V_x is the voltage at the distance x and V_o is the voltage at the origin (x = 0). λ is given by:

(42.5)

Assuming that r_o (the outside longitudinal resistance) is negligibly small compared to r_i (this would be true for a superficial fiber in a bundle immersed in a large bath):

(42.6)

where r_m (Ω·cm) and r_i (Ω/cm) are the membrane resistance and the internal longitudinal resistance normalized for unit length of fiber, R_m (Ω·cm²) is the membrane resistance normalized for both fiber radius and length, R_i (Ω·cm) is the resistivity of the myoplasm (normalized for length and cross-sectional area), and a (cm) is the fiber radius. R_m is often loosely called membrane resistivity, but this is not accurate because for true membrane resistivity (ρ_m) there must be correction for membrane thickness δ:

(42.7)

The factors that determine active velocity of propagation (θ_α) include the intensity of the local-circuit current, threshold potential and the passive cable properties, λ and τ_m. As discussed previously, the greater the rate of rise of the AP, the greater the intensity of the local-circuit current, hence the greater the θ_α. In addition to its dependence on the density of the fast Na⁺ channels (determinant of the maximum Na⁺ conductance, ), the kinetic properties of the channel gating and the threshold potential (V_th), max dV/dt is determined also by the RP (or takeoff potential) (related to the h_∞ versus E_m curve), as discussed previously (Fig. 42.9). In addition, because cooling decreases Na channel activation (Q₁₀ ent 3), max dV/dt and θ_α are slowed accordingly. R_m is increased by cooling, the Q₁₀ for R_K in frog sartorius fibers being about 2.8 (ion diffusion in free solution has a Q₁₀ of about 1.2) (Sperelakis, 1969). For a description of passive conduction, see Appendix III to this chapter.

FIGURE 42.9 Graphic representation of the maximal rate of rise of the APl (max dV/dt) as a function of resting E_m or takeoff potential. Max dv/dt is a measure of the inward current intensity (membrane capacitance being constant), which is dependent on the number of channels available for activation; h is the inactivation factor of Hodgkin–Huxley as g_Na = where g_Na is the Na⁺ conductance, is the maximal conductance and m and h are variables; h_∞ represents h at t = ∞ or steady state (practically, after 20 ms). The fast Na⁺ channels begin to inactivate at about −75 mV and nearly complete inactivation occurs at about −30 mV (h_∞ low). Therefore, max dV/dt decreases because h_∞ decreases.

XIV Excitation Delivery to Fiber Interior by Conduction into the T-Tubular System

The experiments of Huxley and Taylor (1958) were the first to provide evidence that there was some structure, located at the level of the Z-lines in frog skeletal muscle fibers, which is involved in E-C coupling. This structure allows relatively fast conduction of the excitatory process (AP) from the surface membrane to the center of the muscle cells. These investigators applied current pulses at different points along the length of the sarcomeres in isolated fibers and found that when the microelectrode tip was opposite the Z-line, graded contractions of the two half-sarcomeres occurred. The greater the current, the greater was the inward spread of the contraction. In addition, they discovered that there were sensitive spots located around the perimeter of the fiber at the Z-line level; i.e. the membrane was not uniformly sensitive. At about the same time, it was discovered by electron microscopy that transverse (T-) tubules were located at the level of the Z-lines in amphibian skeletal muscle (and at the level of the A-I junctions of the sarcomere in mammalian skeletal muscle). Thus, the T-tubules probably represent the morphological conduit for the findings of Huxley and Taylor. The morphological arrangements of the sarcotubular system of skeletal muscle fibers are illustrated in Fig. 42.10.

FIGURE 42.10 Sarcotubular system of skeletal muscle fibers from tibialis anterior muscle of mouse (A,C) and iliotibialis muscle of lizard (B). (A) Longitudinal section showing the sarcomere structure of several myofibrils: A-band, I-band, Z-line. The network sarcoplasmic reticulum (N-SR), also known as the longitudinal SR, appears as a torn sleeve surrounding the surface of each myofibril. The N-SR is continuous with the junctional SR (J-SR) that abuts close to the transverse tubules (TT). The TT membranes are invaginations of the cell surface membrane at the level of the A-I junctions in mammalians (or at the level of the Z-line in lizards and amphibians). The J-SR and TT form the complex coupling known as a triad (∗). The N-SR is continuous across the I-band, but this is obscured in this section by the presence of paired mitochondria (Mit) over the I-bands. The TT and SR are both selectively filled with osmium tetroxide precipitate, causing their profiles to be more electron opaque than the other structures. Scale bar at lower right represents 1.0 μm. (B) Higher magnification of a triad to show more detail. As shown, the triad consists of a single T-tubule sandwiched between two cisternae of the J-SR. Scale bar = 0.1 μm. (C) High magnification of a triadic junction to illustrate the array of regularly-spaced junctional processes or SR foot processes (several indicated by arrowheads) that project between the TT membrane and the J-SR membrane. There are dense granules within the lumen of the J-SR cisternae (∗). Scale bar = 0.1 μm. (Electron micrographs provided courtesy of Dr Mike Forbes, University of Virginia.)

Diffusion of some substance from the surface membrane into the skeletal muscle fiber interior is much too slow to account for the relatively short latent period of about 1–3 ms between the beginning of the AP and the beginning of contraction. That is, the diameter of the fibers (mean value of about 70 μm in frog sartorius fibers) is much too large for a diffusion mechanism from the fiber surface to be involved. Diffusion time (for 95% equilibration) increases by the square of the distance and would require about 2.5 s for a small molecule freely diffusing across a cell radius of 50 μm; estimates for Ca²⁺ diffusion time are considerably longer than this (Podolsky and Costantin, 1964). Therefore, the T-tubular system serves as an electrical conduit to bring excitation deep into the fiber interior rapidly and thereby reduces the required diffusion distance to an average value of about 0.7 μm (Sperelakis and Rubio, 1971). It was reported that disruption of the T-tubule system (by a glycerol osmotic-shock method) uncouples contraction from excitation (Eisenberg and Gage, 1969).

Estimates of the length constant of the T-tubules (λ_TT), assuming the T-tubule membrane has about the same resistivity (R_m) as the surface membrane give values of about 50 μm. Because the resistivity of the T-tubule membrane of frog muscle is probably higher than that of the surface sarcolemma because of lower g_Cl (Hodgkin and Horowicz, 1959; Adrian and Freygang, 1962; Sperelakis and Schneider, 1968), this would give a longer value. Therefore, it is theoretically possible for the T-tubules to serve as passive conduits to bring the depolarization from the surface membrane (during its AP) into the fiber interior. (This mechanism does operate when small depolarizing voltage changes [below the AP threshold] occur on the surface sarcolemma, and conversely any voltage change originating in the T-tubules can be conducted passively to the surface sarcolemma.)

However, direct microscopic observations of the degree of myofibril activation across the width of the fiber caused by raised [K⁺]_o depolarization suggested otherwise (Gonzales-Serratos, 1975). Also, during the foot of the AP, only a small outer ring of the tubular network depolarizes beyond the mechanical threshold (Hodgkin and Nakajima, 1972). These results imply that in order for all the myofibrils in the cross-section of a fiber to be activated, which had been demonstrated previously (Gonzalez-Serratos, 1971), there must be a T-tubular AP (TT-AP).

There is evidence that the T-tubules actually do fire APs, i.e. they actively propagate impulses inward and so bring large depolarization deep into the fiber interior. The evidence for this includes the observation of a threshold for sudden initiation of localized contraction (Costantin and Podolsky, 1967; Costantin and Taylor, 1973). The TT-AP is sensitive to TTX and is Na⁺ dependent and, therefore, is apparently similar in nature to the surface membrane AP. By use of high-speed cinemicrography to measure sequential activation of the myofibrils in a radial direction, Gonzalez-Serratos (1971) estimated the propagation velocity of the TT-AP (θ_TT) to be about 10 cm/s, with a Q₁₀ of 2.2 (which is similar to the Q₁₀ of the surface membrane for AP conduction). This velocity is sufficient to account for the short latent period before contraction begins.

Early evidence for the existence of active propagation in the T-tubules came from a number of indirect measurements (reviewed in Caputo, 1978). The speed and Q₁₀ (2.2) of the spread of mechanical activation were greater than expected for passive conduction (Gonzalez-serratos 1971). Moreover, twitch tension was reduced by TTX or a Na⁺-deficient medium (Costantin, 1970). Depolarization by elevated [K⁺]_o, which inactivates Na⁺ channels, fails to activate contraction in deeper myofibrils (Gonzales-Serratos, 1975). These results suggested that in order for all the myofibrils in the cross-section of a fiber to be activated, there must be fast Na⁺ channels in the T-tubules and a T-tubular AP (TT-AP). Subsequently, tubular Na⁺ currents have been measured directly (Hille and Campbell, 1976) and direct experimental confirmation of propagating APs in the T-tubules have been demonstrated in amphibian (Nakajima and Gilai, 1980) and mammalian muscle fibers (DiFranco et al., 2005). These direct recordings of the propagating AP in the T-tubules were achieved using potential-sensitive dyes, since the T-tubule membranes are not accessible to conventional microelectrode methods.

In muscles placed in low [Na⁺]_o and stimulated briefly at high frequency, the normal tetanic tension rapidly falls, simultaneous with the central myofibrils becoming inactive. These results are due to Na⁺ depletion in the T-tubule network, particularly in the deeper parts far from the orifice at the fiber surface (Bezanilla et al., 1972). It is thought that the Na⁺ influx (the inward fast Na⁺ current) with each AP in the T-tubule produces a progressive decline in [Na⁺]_TT, which slows propagation velocity down the T-tubules and eventually leads to loss of excitability when [Na]_TT drops below some critical level (e.g. 30 mM). Na⁺ depletion should occur more rapidly deep in the T-tubule network because there would be less diffusion of Na⁺ in from the mouth of the T-tubule to replenish the Na⁺ loss. Active Na⁺-K⁺ pumping in the T-tubules may not occur fast enough to keep up with the Na⁺ loss into the fiber myoplasm.

There are also voltage-dependent slow Ca²⁺ channels in the T-tubule membrane and slow APs that arise from the T-tubule can be recorded under appropriate conditions (Sperelakis et al., 1967; Vogel et al., 1978). The evidence for the existence of this type of channel and some of its properties was discussed above. The Ca²⁺ influx into the myoplasm through these Ca²⁺ channels could play a role in E-C coupling. For a discussion of the relationship between the T-tubules and the terminal cisternae of the SR, see Appendix IV.

Appendix

AI More Information on Cl⁻ Channels

In frog, there are several subtypes of Cl⁻ channels that have single-channel conductances ranging between 40 and 70 pS and each channel may exhibit several subconductance states. But there is usually a main gate that opens or closes the entire channel. In fetal mammalian fibers, Cl⁻ channels with conductances of about 40, 60 and 300 pS have been observed. Myoballs cultured from muscle biopsies of patients having one form of myotonia had a reduced (ca. 50%) single-channel conductance for the Cl⁻ channel, which would contribute to the myotonia (Fahlke et al., 1993). In primary cultures of rat skeletal muscle, the fast Cl⁻ channel showed a behavior consistent with six closed states and two open states (Weiss and Magleby, 1992). The Cl⁻ channel in myoblasts and myotubes of the L6 cell line derived from rat skeletal muscle had a high conductance of about 330 pS (Hurnak and Zachar, 1992). Voltage-gated Cl⁻ channels have also been found in the SR membrane of skeletal muscle.

Some Cl⁻ channels described for other tissues include: (1) Ca²⁺-dependent Cl⁻ channels; (2) stretch-activated Cl⁻ channels; and (3) cyclic AMP-stimulated Cl⁻ channels. The receptor-operated Cl⁻ channels apparently have a G-protein (e.g. G_s or G_i) as intermediate for coupling.

The voltage-dependent Cl⁻ channels can be blocked relatively selectively by several methods, including acidosis and use of compounds such as the stilbene derivatives (DIDS and SITS) and 9-anthracene carboxylic (9-AC) acid. The Cl⁻ channels in frog skeletal muscle are relatively insensitive to 9-AC acid, whereas those in adult mammalian muscle are highly sensitive. The anion selectivity sequence for some voltage-dependent Cl⁻ channels is I⁻ > Br⁻ > Cl⁻ > F⁻.

AII More Information on K_ATP Channels

It has been proposed that K_ATP channels may be associated with the decrease in force development underlying muscle fatigue. A drug (SR44866) that opens K_ATP channels (in sarcolemmal membrane patches from frog skeletal muscles) also reduces the AP duration, the early afterpotential and the peak twitch force (without affecting the RP) in intact frog muscles (Sauviat et al., 1991). The high K⁺ permeability found in muscle fibers that have undergone a permanent contracture (rigor) produced by repetitive stimulation while metabolically-poisoned with cyanide and iodocetate (Fink and Lüttgau, 1976), may be caused by the decreased ATP concentration opening K_ATP channels. In metabolically-exhausted frog semitendinosus muscle fibers, the addition of tolbutamide and glyburide, two K_ATP channel antagonists, significantly reduce the K⁺ efflux rate (Castle and Haylett, 1987). As said previously, even under fatigue induced by prolonged repetitive stimulation, the decrease in intracellular ATP is small (Nassar-Gentina et al., 1978) and fatigued muscle fibers can contract when Ca²⁺ is released directly from the intracellular SR stores (Gonzalez-Serratos et al., 1978; Garcia et al., 1991). In intact animals, some of the skeletal muscle fatigue results from synaptic fatigue, including fatigue at the neuromuscular junction.

The Kir6.x channel is a typical inward-rectifier type K⁺ channel protein. It has two transmembrane helices that form the pore and K⁺ selectivity filter. The cytoplasmic NH₂ and COOH termini interact to form an ATP binding site. A key property of Kir channels is inhibition of channel opening by ATP (Spruce et al., 1987). Each Kir subunit can bind one molecule of ATP. Inhibition by ATP is not a consequence of phosphorylation or ATP hydrolysis, but of direct binding to intracellular domains on the Kir channel (Kakei et al., 1985). That is, ATP is not consumed in this action.

SUR is a regulatory protein that is linked to the C-terminus of Kir. SUR is an ATP-binding cassette (ABC) protein, which itself (unlike other ABC proteins) has no intrinsic transport function, but associates with Kir6.x K⁺ channels to form the functional K_ATP channel. SUR serves as a regulatory subunit which fine-tunes the activity of Kir6.x in response to changes in cell metabolism. Each SUR subunit contains two nucleotide-binding folds which contain binding sites for Mg²⁺-adenosine nucleotides. Cytosolic free Mg²⁺ is high in resting skeletal muscles (ca 1 mM). In the absence of Mg²⁺, nucleotides (such as ADP) act on SUR to inhibit K_ATP activity. Therefore, nucleotides can regulate K_ATP channels through interactions with both Kir and SUR.

As with other members of the Kir family, the K_ATP channel exhibits inward rectification (ir). That is, the open channels pass inward current with increasing hyperpolarization, but outward K⁺ current does not flow when the membrane is depolarized by more than about 10 mV above the E_K potential. This property of inward rectification is caused by cytoplasmic ions such as Mg²⁺ and polyamines plugging the pore pathway upon depolarization and thereby obstructing the outward flow of K⁺ (Woll et al., 1989).

The half-maximum inhibition of opening of K_ATP channels by ATP in skeletal muscle (measured at a constant [K⁺]_o/[K⁺]_i) is 0.135 mM at pH 7.2 (see Fig. 42.6). Intracellular [ATP] is about 5 mM in resting muscle (Dawson et al., 1978, 1980) and it is maintained at near basal levels even during repetitive contractions that lead to muscle fatigue (Carlson and Siger, 1960; Nassar-Gentina et al., 1978). Therefore, the K_ATP channels are largely inactive (or masked) under normal physiological conditions.

In searching for a possible role of the K_ATP channel under other conditions, a modest role was demonstrated in maintaining membrane polarization under sustained muscle use and fatigue. Although ATP is the controller of K_ATP opening, K_ATP activity can be modulated by a number of metabolites through interactions with Kir, SUR or both. These include: K⁺, H⁺ and phospholipids (such as PIP₂), all of which are known to increase during muscle contraction.

Extracellular K⁺ concentration ([K]_o) rises dramatically in the muscle extracellular space during repetitive AP activity and can reach 15–50 mM in the transverse tubules (Almers, 1980; Clausen, 2008). The unitary conductance of the K_ATP channel increases with [K⁺]_o, from 15 pS in 2.5 mM to 42 pS in 60 mM_. Therefore, at higher [K_o], K_ATP channels are more conductive at the same ATP concentration.

Kir channels are also regulated by pH at the cytoplasmic surface (pH_i) (reviewed in Jiang et al., 2002; Qu et al., 2000). Intracellular pH may drop by up to one unit during intense muscle use (Renaud, 1989). A decrease in pH_i reduces the inhibitory effect of ATP on K_ATP channels, leading to an increase in the probability of K_ATP channel opening (Davies et al., 1992). The ATP concentration required for half-inhibition increases by 2.5 times and 15 times, respectively, at pH_i 6.8 and 6.3, compared to that at pH_i 7.2. That is, a decrease in pH_i increases the ATP concentration required for half-inhibition of channel activity. During acidosis, a small decrease in ATP concentration leads to a greater opening of K_ATP channels.

The decrease in pH_i during contraction is matched by an increase in cytosolic free Mg²⁺. A rise in cytosolic Mg²⁺ reduces the ability of ATP to close K_ATP channels (Vivaudou et al., 1991). This is most likely related to the ability of Mg²⁺ to bind to ATP and to the stimulatation of K_ATP channels via the SUR regulatory subunit. Although the increases in [K]_o_, pH_i and [Mg²⁺]_i may act collectively and synergistically to activate K_ATP channels during repetitive muscle contraction, this effect is modest at normal muscle [ATP]_i.

On the other hand, K_ATP channels may become activated when the muscle is stressed or metabolically compromised (Hussain et al., 1994). Adenosine, which is released from metabolically-compromised muscle, stimulates K_ATP (Bartlett-Jolley and Davies, 1997) and, as said previously, intracellular acidosis is a potent activator of K_ATP channels. Although the skeletal muscles of genetically-altered mice which lack Kir (Kir6.2 knockout mice) develop fatigue more rapidly than control muscles (Cifelli et al., 2007), the more rapid onset of fatigue is not associated with K_ATP channel activation (Boudreault et al., 2010). These findings indicate that K_ATP channels are not essential for maintaining force during fatigue.

The activation of K_ATP channels after fatigue has developed helps preserve a polarized membrane potential, keeping it near E_K. This function prevents voltage-dependent Ca²⁺ entry that can occur following fatigue and cause fiber injury. Consistent with this idea, both Kir6.2 knockout mice and SUR knockout mice show extensive muscle fiber damage when subjected to exercise training (Thabet et al., 2005; Stoller et al., 2007). Therefore, K_ATP channels in muscle serve a myoprotective role to prevent fiber damage following intense fatigue or during metabolic exhaustion. In this respect, the role of K_ATP channels in skeletal muscles is similar to that in other cell types, i.e. K_ATP channels serve as molecular sensors of cellular metabolism, linking metabolism to membrane excitability.

AIII Further Evidence that the T-Tubules Fire Na⁺-Dependent APS

Further evidence of a Na⁺-dependent TT-AP came from observations in which muscles placed in low [Na⁺]_o and stimulated briefly at high frequency, initially develop normal tetanic tension that rapidly falls and steadily decreases to a lower sustained tension; simultaneously the central myofibrils becoming inactive. These results are due to Na⁺ depletion in the T-tubule network, particularly in the deeper parts far from the orifice at the fiber surface. Na⁺ depletion occurs because g_Na increases during the TT-AP generation, causing an inward Na⁺ flow into the myoplasm (Bezanilla et al., 1972). The fatigue occurs more rapidly in fibers pre-equilibrated in low [Na⁺]_o (e.g. 60 mM). There is a progressive decline in [Na⁺]_TT, which slows propagation velocity down the T-tubules and eventually leads to loss of excitability when [Na]_TT drops below some critical level (e.g. 30 mM). Na⁺ depletion should occur more rapidly deep in the T-tubule network because there would be less diffusion of Na⁺ in from the mouth of the T-tubule to replenish the Na⁺ loss. Active Na⁺-K⁺ pumping in the T-tubules may not occur fast enough to keep up with the Na⁺ loss into the fiber myoplasm.

There is evidence that the T-tubules actually do fire APs, i.e. they actively propagate impulses inward and so bring large depolarization deep into the fiber interior. The evidence for this includes the observation of a threshold for sudden initiation of localized contraction (Costantin and Podolsky, 1967; Costantin and Taylor, 1973). The TT-AP is sensitive to TTX and is Na⁺ dependent and, therefore, is apparently similar in nature to the surface membrane AP. By use of high-speed cinemicrography to measure sequential activation of the myofibrils in a radial direction, Gonzalez-Serratos (1971) estimated the propagation velocity of the TT-AP (θ_TT) to be about 10 cm/s. The Q₁₀ of 2.2 is similar to that of the surface membrane AP. Although this velocity is about 16 times slower than propagation down the fiber longitudinally (about 1.6 m/s), it is sufficient to account for the short latent period before contraction begins. Additional data supporting the presence of APs in the T-tubules has been obtained more recently using V-sensitive dyes.

AIV Propagation Velocity in a Passive Cable

In a passive cable, such as in a resting skeletal muscle fiber, the passive propagation velocity (θ_ρ) is directly proportional to the length constant and inversely proportional to the time constant:

(42A.1)

(42A.2a)

(42A.2b)

Therefore, propagation velocity is directly proportional to the square root of the fiber radius (a) and inversely proportional to membrane capacitance (C_m) and to the square root of R_i and the square root of R_m. For example, propagation velocity is greater in large-diameter muscle fibers. The length constant for sinusoidally-varying applied currents (λ_ac) is shorter than λ_dc, depending on the ac frequency.

The relationship between propagation velocity and membrane current density (I_m) is given by:

(42A.3)

where d²V/dt² is the second time derivative of the AP. As indicated, membrane current is proportional to d²V/dt², whereas the longitudinal current (I_l) or the capacitative current (I_c) is proportional to dV/dt:

(42A.4)

AV Evidence for T-Tubule Communication with the SR across the Triadic Junction under Some Conditions

This Appendix section is included to let the student know that there are data that do not fit with currently accepted hypotheses.

Ca²⁺ for contraction in skeletal muscle is primarily released from the TC-SR (Winegrad, 1968) and there is an internal cycling of Ca²⁺ ion. Changes in [Ca²⁺]_o of the bathing solution take a relatively long time (e.g. 30 min) before exerting a large effect on the force of contraction. In contrast, in cardiac muscle, the effect of lowered [Ca²⁺]_o is significant within a few seconds, indicating that the primary determinant of the force of contraction is the Ca²⁺ influx across the sarcolemma through the slow Ca²⁺ channels. Therefore, in skeletal muscle, excitation propagates actively down the T-tubules and Ca²⁺ is released from the TC-SR, but it is controversial as to how the signal is transferred from the T-tubule to the TC-SR across the triadic junction.

Electron-opaque tracer molecules, like horseradish peroxidase (HRP) (ca. 60 Å diameter), enter into the T-tubules and from there can enter into some of the TC-SR of frog skeletal muscle (Rubio and Sperelakis, 1972; Kulczycky and Mainwood, 1972) (Fig. 42A.1A,B). Exposure of the fibers to hypertonic solutions facilitates the entry of HRP into the TC-SR, so that nearly 100% of the TC-SR become filled (Fig. 42.A.1C,D). Thus, there may be a functional connection between the SR and the extracellular space (Sperelakis et al., 1973). If so, there may be lumen-to-lumen continuity between the T-tubules and TC-SR during excitation, allowing the AP in the T-tubules to invade directly into the TC-SR to depolarize and bring about the release of Ca²⁺. The depolarization of the TC-SR could activate voltage-dependent Ca²⁺ channels, allowing Ca²⁺ influx into the myoplasm down an electrochemical gradient.

FIGURE 42A.1 Evidence that large molecules of horseradish peroxidase (HP) can enter into the terminal cisternae (TC) of the SR via the transverse tubules (TT) of frog sartorius fibers. Electron micrographs of longitudinal sections. (A,B) Fiber was exposed to HP under isosmotic conditions. (A) Section through several myofibrils showing presence of HP activity (as a dense electron-opaque material) in the TT and in some of the TC at triadic junctions. In amphibian muscle, the TT occur at the level of the Z-lines (Z) of the sarcomeres. Arrows point to two branches of the TT running longitudinally. (B) Higher magnification of two triads, one with both cisternae filled with HP and the other with only one cisterna filled. (C,D) Fiber was exposed to HP under hypertonic condition (3 X isotonic, using NaCl), showing that almost all cisternae were filled with peroxidase. (C) Section at low magnification. (D) Portion of same section as in Part C shown at higher magnification. The surface vesicles (Ves) also became filled with HP. (Modified from Figs. 2 and 4 of Rubio, R. and Sperelakis, N. (1972). Z. Zellforsch. 124, 57–72.)

If the longitudinal SR (L-SR) were electrically isolated from the TC-SR by a substantial resistance (e.g. zippering between the two SR compartments, described later), this would account for the fiber capacitance measured being relatively low (Mathias et al., 1980). The effect of this would be to remove the very large membrane surface area of the L-SR and hence greatly reduce the capacitance that would be measured.

It has been suggested that the SR is depolarized during the release of Ca²⁺ in E-C coupling. For example, optical signals (e.g. birefringence and fluorescence changes) can be recorded from the SR membranes during contraction (e.g. Baylor and Oetliker, 1975; Bezanilla and Horowicz, 1975). In addition, Natori (1965) demonstrated that propagation of contraction (1–3 cm/s) triggered by electrical stimulation can occur in muscle fiber regions that had been denuded (skinned) of their sarcolemma, the propagation of excitation presumably occurring by means of the SR membranes.

It was demonstrated that E-C uncoupling could be produced by exposing frog skeletal muscle fibers to Mn²⁺ (1 mM) or La³⁺ (1 mM) while in hypertonic solution (to facilitate entry of the blockers into the TC-SR) (Sperelakis et al., 1973). After the fibers were returned to normal Ringer solution, normal fast APs could be elicited, but there were no contractions accompanying them; i.e. a “permanent” E-C uncoupling was produced. These results were interpreted as suggesting that Mn²⁺ and La³⁺ entered into the lumen of the TC-SR and blocked the Ca²⁺ channels. A similar exposure of frog sartorius fibers to Mn²⁺, La³⁺ or to Ca²⁺-free solution blocked the caffeine-induced contracture as well (Rubio and Sperelakis, 1972). Thus, from these physiological and ultrastructural studies, it was suggested that the lumen of the SR is continuous with that of the T-tubule under conditions of hypertonicity and that substances can enter into the TC-SR to exert an effect on Ca²⁺ release into the myoplasm.

Compartmental analysis of skeletal muscle has also suggested that the SR is open to the ISF. (In contrast, in cardiac muscle, there is no evidence that the SR is open to the ISF [Rubio and Sperelakis, 1972].) For example, Conway (1957), Harris (1963) and Keynes and Steinhardt (1968) concluded that Na⁺ inside frog skeletal muscle fibers is distributed in two separate compartments. Harris (1963) suggested that the Na⁺, K⁺ and Cl⁻ concentrations in one compartment (presumably the SR) were about equal to those of the ISF. Rogus and Zierler (1973) concluded that the Na⁺ concentration in the SR of rat skeletal muscle approximates that of the ISF. The volume of the SR compartment was 14.3% of fiber volume and, in hypertonic solution, the SR volume increased and the washout of the SR compartment was faster. Tasker et al. (1959) also had reported a large sucrose space of 26.5% for frog sartorius fibers.

Other researchers (Birks and Davey, 1969) have demonstrated that the volume changes of the SR of skeletal muscle in hypertonic (sucrose) and hypotonic solutions were always opposite of those occurring within the myoplasmic compartment. They concluded that sucrose must enter into the SR, pulling in water osmotically from the myoplasm, to produce the marked swelling of the SR that occurred in hypertonic solutions. Vinogradova (1968) concluded from the distribution of non-penetrating sugars in frog sartorius muscle that the SR compartment is continuous with the ISF; the inulin space was 19.0% and increased in hypertonic solution and decreased in hypotonic solution and in glycerol-treated fibers (for disruption of the T-tubules).

The total [3H]-sucrose space of frog sartorius muscles was found to be 18.0% in isotonic solution and 22.6% in twofold hypertonic solution (Sperelakis et al., 1978). The relative SR volume (including the small T-tubule volume) was 12.4% and 17.0% of fiber volume, respectively. This value for SR volume of frog skeletal muscle is close to that measured by ultrastructural techniques (Peachey, 1965; Mobley and Eisenberg, 1975). Evidence that the TC-SR and L-SR may not be freely connected to one another under resting conditions comes from the observations that: (1) the L-SR did not fill with HRP, whereas the TC-SR did (Rubio and Sperelakis, 1972); and (2) there is a zippering of the membranes connecting these two components of the SR in mouse and frog skeletal muscle (Howell, 1974; Wallace and Sommer, 1975; Forbes and Sperelakis, 1979).

In ⁴⁵Ca washout experiments on frog muscles, Kirby et al. (1975) found three compartments, similar to the three sucrose compartments described previously, except the half-times were about two- to threefold shorter. They suggested that the first compartment was the ISF space, the second was the T-tubule plus the TC-SR and the third was the L-SR. Bianchi and Bolton (1974) also found a transient increase in ⁴⁵Ca efflux and a marked loss of muscle Ca²⁺ from frog sartorius muscles exposed to hypertonic solutions (twice isotonicity) and suggested that hypertonicity produces transient communication between the TC-SR and the T-tubules, thus allowing their Ca²⁺ to be lost to the ISF. In addition, it has been reported in a human muscle disease, polymyositis, that the T-tubules are spatially continuous with the SR, as visualized with lanthanum tracer, and that enzymes leak from the TC-SR into the T-tubules and ISF (Chou et al., 1980).

Frog skeletal muscle fibers have an osmotically inactive volume of about 32% when placed into Ringer solution made hypertonic with sucrose or other non-penetrating solutes; i.e. fiber diameter does not shrink to the theoretical value expected if it were a perfect osmometer (Sperelakis and Schneider, 1968; Sperelakis et al., 1970). For example, in twofold hypertonic solution, there should be a decrease in fiber volume to one-half and fiber radius to 0.707 of the original value. The observed change is to only 0.81 of the original diameter. Because the SR volume increases in hypertonic solution (Huxley et al., 1963; Sperelakis and Schneider, 1968; Birks and Davey, 1969), it is likely that the osmotic inactive volume is due to the SR. The swollen SR would prevent the fiber volume from decreasing to one-half in twofold hypertonic solution, even if the volume of the myoplasm proper were to decrease to one-half.

In cardiac muscle, an osmotically-inactive volume is not present (Sperelakis and Rubio, 1971), electron-opaque tracers do not enter the SR (Sperelakis et al., 1974) and the SR volume does not increase with hypertonicity (Sperelakis and Rubio, 1971).

AVI Invertebrate Striated Muscle Fibers

This Appendix was prepared by Hugo Gonzalez-Serratos and Hector Rasgado-Flores and was edited by Nicholas Sperelakis.

Introduction

In 1902, Overton observed that frog skeletal muscle cells lost their excitability when the extracellular Na⁺ concentration ([Na⁺]_o) was decreased below 10% of normal. This was the first indication that excitability of vertebrate skeletal muscle fibers was Na⁺-dependent. Using intracellular microelectrodes, Nastuck and Hodgkin demonstrated, in 1950, that skeletal muscle fibers of the frog have a clear relationship between the amplitude of the AP and [Na⁺]_o. They concluded that vertebrate striated skeletal muscle APs are generated by mechanisms similar to those in squid giant nerve axons. In an attempt to test this Na⁺ hypothesis in striated muscle fibers of the crustacean, Fatt and Katz (1953) bathed crayfish muscles with different external Na⁺ concentrations. However, against their prediction, they found that there was no effect on the amplitude of the APs. That is, striated invertebrate crayfish muscles were excitable and capable of developing full APs in the absence of external Na⁺ ions. Thus, they concluded that invertebrate striated muscle fibers did not have an Na-dependent excitability. Instead, the APs were generated by currents using other ions. The APs of invertebrate striated muscle fibers, compared to vertebrates, are longer in duration and some of them even have a plateau (similar to cardiac muscle) (Fig. 42A.2).

FIGURE 42A.2 Different action potentials in invertebrate striated muscle fibers of different crab species. (A) Portunus (fromAtwood, 1965 ). (B) Carcinus maenas (fromFatt and Katz, 1953 ). (C) Portunus deporatus (fromFatt and Katz, 1953 ). Voltage calibration bars:40 mV.

Calcium Hypothesis

What are the ionic currents that generate the APs in invertebrate striated muscle fibers? With zero extracellular Na⁺, the invertebrate muscles were capable of generating APs as long as there were Ca²⁺ ions in the bathing solution. It was concluded that these fibers produced Ca²⁺-dependent APs (Fatt and Ginsborg, 1958). These APs persisted when Sr²⁺ or Ba²⁺ (which have some physicochemical properties similar to Ca²⁺) replaced the Ca²⁺. This led to the calcium hypothesis. Hagiwara and Naka (1964) induced APs in the giant barnacle striated muscles fibers (Balanus nubilus) in which they changed the intracellular Ca²⁺ concentration by injecting the fibers with Ca²⁺chelating substances. As the internal free Ca²⁺ concentration was decreased progressively, the AP amplitude increased progressively. At an internal Ca²⁺ concentration of 8×10⁻⁸ M, the AP became all-or-none (Hagiwara and Nakajima, 1966). Thus, they confirmed the generation of Ca²⁺ APs in invertebrate striated muscle fibers.

Three types of experiments have proven the Ca²⁺ hypothesis for invertebrate striated muscle cells: (1) the Ca²⁺-dependence of the AP amplitude; (2) the lack of APs in zero Ca²⁺ bathing solution; and (3) the entry of Ca²⁺ ions during the APs.

Ca²⁺ dependence of the APs. Fatt and Ginsborg in 1958 and Pilgrim and Wiersma in 1963 demonstrated that the striated myofibers of the crayfish (Orconectes virilis) and lobster (Homarus americanus) were capable of generating propagated all-or-none APs. These APs did not disappear when the muscles were immersed in bathing solutions without Na⁺, K⁺ or Mg²⁺. However, the amplitude of the APs depended on the extracellular Ca²⁺ concentration [Ca²⁺]_o. The AP amplitude increased with a slope of 25 mV per decade of [Ca ²⁺]_o. These results strongly supported the proposal that the APs of invertebrate striated skeletal muscle fibers are generated by inward Ca²⁺ currents. In the crayfish (Astacus fluvilis), the potentiating effect of [Ca²⁺]_o was strongly enhanced in the presence of quaternary ammonium ions like tetraethylammonium (TEA, an inhibitor of outward I_K) (Fig. 42A.3). The maximum rate of depolarization of the APs increased from 8 V/s to 15 V/s when [Ca²⁺]_o was changed from 4 to 16 mM. In the giant muscle fibers from the barnacle (Balanus nubilus), Hoyle and Smith (1963) and Hagiwara and Naka (1964) found that the usual graded responses were converted to all-or-none responses when the gradient between extracellular and intracellular [Ca²⁺] increased by injecting the cells with Ca²⁺-chelating substances like EDTA and K₂SO₄. The amplitudes of the APs were a function of the extracellular and intracellular [Ca²⁺] ratios.

FIGURE 42A.3 Potentiation of invertebrate striated muscle of the crayfish (Astacus fluvialitis) with TEA. Upper trace: action potential. Lower trace: applied current. (From Fatt and Ginsborg, 1958.)

Calcium influx. Hagiwara and Naka (1964) found that, during prolonged stimulations that produced repetitive APs, there was a substantial amount of radioactive Ca²⁺ influx into barnacle giant muscle. They estimated that the intracellular Ca²⁺ concentration increased by an average of 75 pmole/cm²/impulse and calculated that, during each AP, there was an influx of 6 pmoles of Ca²⁺/cm². This is a very high Ca²⁺ influx compared with the values reported for vertebrate skeletal muscle fibers (Bianchi and Schanes, 1959).

Prolonged depolarization (elicited with high concentrations of extracellular K⁺) above the mechanical threshold leads to sustained mechanical activation (contractures). When K⁺-contractures were elicited in a bathing solution without Ca²⁺ ions, the K⁺-contractures disappeared after a short time, even though the membrane remained depolarized. This indicates that, during prolonged depolarization of invertebrate striated muscles, there is a substantial inflow of Ca²⁺ that brings about the mechanical activation.

Potentiation of APs.Fatt and Ginsborg (1958) observed that when all the extracellular Ca²⁺ was removed and substituted with equimolar concentrations of SrCl₂ or BaCl₂, the crayfish striated muscles not only produced all-or-none APs, but they were of higher amplitude and longer duration (Fig. 42A.4). The RPs were unaffected. These results were confirmed by Werman and Grundfest (1961) in lobster muscle fibers and by Werman et al. (1961) in the muscle of the grasshopper (Romalea microptera). Thus, inward Sr²⁺and Ba²⁺ currents are also capable, like Ca²⁺ ions, of generating APs. The increased AP durations were the result of increased durations of the plateau. The control muscle fibers had an average duration of only 3–6 ms. The duration of the Ba²⁺ APs was 15–100 ms, compared with 400–12 000 ms for the Sr²⁺ APs (Fig. 42A.4). The AP overshoot was about 20 mV larger with Ba²⁺ compared with Sr²⁺ in striated muscles from crayfish (Astacus fluvialis) and lobster (Homarus americanus) (Fatt and Ginsburg, 1958; Werman and Grundfest, 1961). The RP of −90 mV did not change with Sr or Ba ions.

FIGURE 42A.4 Crayfish striated muscles fibers. Action potentials of fibers bathed with strontium (left panel) and barium (right panel). (From Fatt and Ginsborg, 1958.)

Ionic currents. Hagiwara and Naka (1964), by using a voltage-clamp method, showed that the currents developed during the AP were composed of early inward Ca²⁺ currents, since they varied in amplitude with extracellular Ca²⁺ concentration and were not affected by TTX. It was concluded that, in invertebrate striated muscle fibers, the propagated AP was the consequence of inward Ca²⁺ current. Hen ent ek et al. (1969) measured Sr²⁺ current in the crayfish (Astacus fluviaitilis) using a sucrose-gap voltage-clamp. They found that the currents consisted of early inward current followed by outward current. The Ca²⁺ currents were smaller than the Ba²⁺ currents (Fig. 42A.5). The late outward current was decreased by the K⁺-current blocker TEA (Fig. 42A.6).

FIGURE 42A.5 Membrane currents under voltage clamp from crayfish striated muscle (Astacus Fluviatilis) bathed with extracellular strontium chloride isotonic solution. The clamped membrane potentials are shown in mV. Records below the zero line are inward currents, above are outward currents.

FIGURE 42A.6 Effect of TEA on membrane currents from the crayfish striated muscle (Astacus Fluviatilis) under voltage clamp. Top records, bathed with normal extracellular solution. Bottom records, after addition of 50 mM of TEA to the bathing solution. Calibrations: 0.15 mA/cm² and 5 ms. (From Hen ent ek et al., 1969.)

The AP in invertebrate striated muscles, like in vertebrate striated muscles, fulfills two main roles. One role is to propagate a depolarization wave all along the length of the entire muscle fiber. Another role is to increase the myoplasmic Ca²⁺ to levels that induce the binding of myosin with actin to bring about activation of the contractile proteins (Fig. 42A.7). The mechanical activation develops either shortening, force, or both (producing muscle work).

FIGURE 42A.7 Myoplasmic calcium transients recorded with the luminescent ptotein aequorin during stimulations of barnacle (Balanus nubilis) striated muscle. Traces: 1: membrane potentials; 2: light output, myoplasma calcium transients; 3: isometric forces; 4: stimulating electrical pulses. On each set of recording traces, the recordings from top to bottom correspond to each other. Vertical calibrations: 20 mV, 3.8×10⁻⁹ lumens, 5 g. Horizontal calibration: 100 ms. (From Ashley and Ridgway, 1968.)

Regulation of Intracellular Ca²⁺

Vertebrate skeletal muscle fibers bathed in Ringer’s solution containing 1 mM [Ca²⁺] had an average Ca²⁺ influx at rest of 0.26 pmoles/cm²/s and, during stimulation, the Ca²⁺ influx increased to 0.73 pmoles/cm²/impulse (Curtis,1966). Blocking the inward Ca²⁺ current has no effect on the contraction or E-C coupling mechanism (Gonzalez-Serratos et al., 1982). Thus, the main role of inward Ca²⁺ current in vertebrate skeletal muscles fibers is to maintain long-term adequate amounts of intracellular Ca²⁺ (mainly in the SR²⁺).

In contrast, invertebrate striated muscle fibers have an average Ca²⁺ influx of 75 pmoles/cm²/impulse (Hagiwara and Naka, 1964). Since the SR is sparse, there must be another mechanism to decrease the free myoplasmic Ca²⁺ after activation. Otherwise, these muscles would develop permanent and powerful contractures and suffer deleterious effects. The mechanism by which invertebrate striated muscles decrease and regulate the free intracellular Ca²⁺ concentration is via the sarcolemmal Ca²⁺/H⁺ exchanger mediated by Ca²⁺-ATPase (DeSantiago et al., 2007) and the Na⁺/Ca²⁺ exchanger (Rasgado-Flores et al., 1991). The Na⁺/Ca²⁺ exchanger mediates the coupling of the movement of external Na⁺ ions down their electrochemical gradient, with the movement of internal Ca²⁺ ions (in the opposite direction) across the sarcolemma. In giant striated muscle of the barnacle (Balanus nubile) (like most excitable cells), the Na⁺/Ca²⁺ exchanger has a stoichiometry of three Na⁺ ions exchanged per each Ca²⁺ ion (Rasgado-Flores et al., 1989).

Innervation of Invertebrate Muscle Fibers

One important characteristic of the innervation of non-vertebrate striated muscle fibers is that the nerve terminals on the muscle fibers form terminal buttons spread all along the muscle fiber. This is in contrast with vertebrate muscle where a motor nerve ends in one button localized near the middle of the fiber. The branching of invertebrate motor nerves leads to multiple nerve terminals that are distributed along the length and around the perimeter of the muscle fibers. There are some differences as a consequence of the differences in the topological distribution of the nerve terminals. The excitatory responses to motor nerve APs can trigger in the muscle fiber either local APs or full-size APs all along the muscle fiber. The transmission of the excitatory process leads to multiple postsynaptic potentials. The regulation of force development during contractions of vertebrate muscles is under central nervous system control. In contrast, in invertebrate muscles, the regulation of the degree of contractions is done peripherally and depends on the number of either excitatory or inhibitory nerve terminals that are activated.

Another feature of invertebrate neuromuscular junctions is that they are structurally simpler than vertebrate motor end-plates. The motor nerves of the crustaceans have short nerve terminals that have periodical swellings in series. The swellings make contact with the surface membrane of the muscle fiber. The nerve terminals are very close together. The response of the invertebrate neuromuscular junction to a nerve AP is called the junctional potential (JP). Evoked JPs are due to a large and synchronized release of neurotransmitters triggered by the nerve terminal AP. The JPs of invertebrates are either excitatory (EJPs) or inhibitory (IJPs). The excitatory neurotransmitter is glutamate. The inhibitory neurotransmitter is GABA (γ-aminobutyric acid). The release of these transmitters will lead either to excitation and mechanical activation of the muscle fiber or inhibition of the excitation and thus producing relaxation. The RP can be hyperpolarized due to the IJPs. Therefore, the amount of current necessary to reach the mechanical threshold voltage (for E-C coupling) can also increase as a consequence of the IJPs hyperpolarizing the membrane.

The electrical responses (postsynaptic potentials) at the neuromuscular junctions are, in most cases, small and graded. But when the small responses are summated, the muscle membrane reaches threshold that generates a full AP that propagates along the muscle fiber. Another important feature of invertebrate muscle innervations is that a single myofiber can be innervated by more than one motor neuron, i.e. polyneural innervation. The polyneural innervation leads to complex electrical and mechanical responses. The topological distribution of nerve junctions and the graded activation permit the existence of fast and slow contractions, as well as excitation and inhibition (often within a single myofiber).

When an EJP reaches the mechanical threshold, a local contraction is produced. When several EJPs are elicited in synchrony, they generate a graded contraction that can show summation. This summation causes a large contraction. IJPs cause hyperpolarization, moving the membrane potential away from the mechanical threshold, thereby causing relaxation. The multiple innervations characteristics of invertebrate muscles cause one muscle fiber to produce EJPs while its neighbor may produce IJPs. The same muscle fiber can show an excitatory response that develops force and an inhibitory response that reduces the force developed (Dudel and Kuffler, 1961). From one to nine excitatory nerves may innervate a single muscle fiber, while the number of inhibitory nerves can be up to three (Kennedy et al., 1966). The size of EJPs (initially of low magnitude) increases progressively during repetitive stimulation and this is called facilitation (Hoyle and Wiersma, 1958). In contrast, the large EJPs decrease in size during the repetitive activation due to transmitter depletion or fatigue, a process known as fatigue or anti-facilitation (Hoyle and Burrows, 1973). Another feature of invertebrate muscle fibers is that the faster fibers display higher E-C thresholds. Therefore, they require EJPs of larger magnitude to reach the E-C threshold. They also develop stronger twitches.

There is relevant information about crayfish muscle stretch receptors given in a previous chapter.

BIBLIOGRAPHY

1. Adrian RH, Freygang WH. The potassium and chloride conductance of frog muscle membrane. J Physiol (London). 1962;163:61–103.

2. Adrian RH, Peachey LD. Reconstruction of the action potential of frog sartorius muscle. J Physiol (London). 1973;235:103–131.

3. Adrian RH, Chandler WK, Hodgkin AL. Voltage clamp experiments in striated muscle fibers. J Physiol (London). 1970a;208:607–644.

4. Adrian RH, Chandler WK, Hodgkin AL. Slow changes in potassium permeability in skeletal muscle. J Physiol (London). 1970b;208:645–668.

5. Almers W, Fink R, Palade PT. Calcium depletion in frog muscle tubules: the decline of calcium depletion in frog muscle tubules: the decline of calcium current under maintained depolarization. J Physiol (London). 1981;312:177–207.

6. Armstrong CM, Bezanilla FM, Horowicz P. Twitches in the presence of ethylene glycol bis(β-aminoethyl ether)-N, N′tetraacetic acid. Biochem Biophys Acta. 1972;267:605–608.

7. Ashley CC, Ridgway EB. Simultaneous recording of membrane potential, calcium transient and tension in single muscle fibers. Nature. 1968;219:1168–1169.

8. Atwood HL. Characteristics of fibres in the extensor muscle of a crab. Comp Biochem Physiol. 1965;14:205–207.

9. Baylor SM, Oetliker H. Birefingence experiments on isolated skeletal muscle fibres suggest a possible signal from the sarcoplasmic reticulum. Nature. 1975;253:97–101.

10. Beaty GN, Stefani I. Calcium dependent electrical activity in twitch muscle fibers of the frog. Proc R Soc London (Biol). 1976;194:141–150.

11. Bezanilla F, Horowicz P. Fluorescence intensity changes associated with contractile activation in frog muscle stained with Nile Blue A. J Physiol (London). 1975;246:709–735.

12. Bezanilla F, Caputo C, Gonzalez-Serratos H, Venosa RA. Sodium dependence of the inward spread of activation in isolated twitch muscle fibres of the frog. J Physiol (London). 1972;223:507–523.

13. Bianchi CP, Bolton TC. Effect of hypertonic solutions and glycerol treatment on calcium and magnesium movements of frog skeletal muscle. J Pharmacol Exp Ther. 1974;188 536–522.

14. Bianchi CP, Shanes AM. Calcium influx in skeletal muscle at rest, during activity, and during potassium contracture. J Gen Physiol. 1959;42:803–815.

15. Birks RI, Davey DF. Osmotic responses demonstrating the extracellular character of sarcoplasmic reticulum. J Physiol (London). 1969;21:171–188.

16. Boudreault L, Cifelli C, Bourassa F, Scott K, Renaud JM. Fatigue preconditioning increases fatigue resistance in mouse flexor digitorum brevis muscle with non-functioning K(ATP) channels. J Physiol. 2010;588:4549–4562.

17. Caputo C. Excitation and contraction processes in muscle. Annu Rev Biophys Bioeng. 1978;7:63–83.

18. Carlson ED, Siger A. The mechanochemistry of muscular contraction I The isometric twitch. J Gen Physiol. 1960;44:33–60.

19. Castle NA, Haylett DG. Effect of channel blockers on potassium efflux from metabolically exhausted frog skeletal muscle. J Physiol. 1987;383:31–43.

20. Chou SM, Nonaka I, Voice GF. Anastomoses of transverse tubules with terminal cisternae in polymyositis. Arch Neurol. 1980;37:257–266.

21. Cifelli C, Bourassa F, Gariepy L, Banas K, Benkhalti M, Renaud JM. KAPT channel deficiency in mouse flexor digitorum brevis causes fiber damage and impairs Ca²⁺ release and force development during fatigue in vitro. J Physiol. 2007;582:843–857.

22. Clausen T. Clearance of extracellular K⁺ during muscle contraction–roles of membrane transport and diffusion. J Gen Physiol. 2008;131:473–481.

23. Cognard C, Rivet-Bastide M, Constantin B, Raymond G. Progressive predominance of ‘skeletal’ versus ‘cardiac’ types of excitation-contraction coupling during in vitro skeletal myogenesis. Pflügers Arch. 1992;422:207–209.

24. Conway EJ. Nature and significance of concentration relations of potassium and sodium ions in skeletal muscle. Physiol Rev. 1957;37:84–132.

25. Cordoba-Rodriguez R, Gonzalez-Serratos H, Matteson DR, Rozycka M. Ica is important in E-C coupling in developing cultured embryonic amphibian skeletal muscle cells. Biophys J. 1997;72:A119 H5.

26. Cordoba-Rodriguez R, Matteson DR, Gonzalez-Serratos H. Embryonic E-C coupling in cultured skeletal muscle cells. Biophys J. 1996;70:A390.

27. Costantin LL. The role of sodium current in the radial spread of contraction in frog muscle fibers. J Gen Physiol. 1970;55:703–715.

28. Costantin LL, Podolsky RJ. Depolarization of the internal membrane system in the activation of frog skeletal muscle. J Gen Physiol. 1967;50:1101–1124.

29. Costantin LL, Taylor SR. Graded activation in frog muscle fibers. J Gen Physiol. 1973;61:424–443.

30. Curtis RA. Ca fluxes in single twitch muscle fibers. J Gen Physiol. 1966;50:225–267.

31. Davies NW, Standen NB, Stanfield PR. The effect of intracellular pH on ATP-dependent potassium channels of frog skeletal muscle. J Physiol. 1992;445:549–568.

32. Dawson MJ, Gadian DG, Wilkie DR. Muscular fatigue investigated by phosphorus nuclear magnetic resonance. Nature. 1978;274:861–866.

33. Dawson MJ, Gadian DG, Wilkie DR. Mechanical relaxation rate and metabolism studied in fatiguing muscle by phosphorus nuclear magnetic resonance. J Physiol. 1980;299:465–484.

34. DeSantiago J, Batlle D, Khilnani M, et al. Ca²⁺/H⁺ exchange via the plasma membrane Ca²⁺ ATPase in skeletal muscle. Front Biosci. 2007;12:4641–4660.

35. DiFranco M, Capote J, Vergara JL. Optical imaging and functional characterization of the transverse tubular system of mammalian muscle fibers using the potentiometric indicator di-8-ANEPPS. J Memb Biol. 2005;208:141–153.

36. Dudel J, Kuffler SW. Presynaptic inhibition at the crayfish neuromuscular junction. J Physiol. 1961;155:543–562.

37. Eisenberg RS, Gage PW. Ionic conductances of the surface and transverse tubular membranes of frog sartorius fibers. J Gen Physiol. 1969;53:279–297.

38. Fabiato A. Mechanism of calcium-induced release of calcium from the sarcoplasmic reticulum of skinned cardiac cells studied with potential-sensitive dyes. In: Ohnishi ST, Endo M, eds. The Mechanism of Gated Calcium Transport Across Biological Membranes. New York: Academic Press; 1982;:237–255.

39. Fahlke C, Zachar E, Rudel R. Chloride channels with reduced single-channel conductance in recessive myotonia congenita. Neuron. 1993;10:225–232.

40. Fatt P, Ginsborg BL. The ionic requirements for the production of action potentials in crustacean muscle fibers. J Physiol (London). 1958;142:156–543.

41. Fatt P, Katz B. The electrical properties of crustacean muscle. J Physiol. 1953;120:171–204.

42. Fink R, Lüttgau HC. An evaluation of the membrance constants and the potassium conductance in metabolically exhausted muscle fibers. J Physiol. 1976;263:215–238.

43. Flagg TP, Enkvetchakul D, Koster JC, Nichols CG. Muscle KATP channels: Recent insights to energy sensing and myoprotection. Physiol Rev. 2010;90:799–829.

44. Flucher BE, Takekura H, Franzini-Armstrong C. Development of the excitation-contraction coupling apparatus in skeletal muscle: association of sarcoplasmic reticulum and transverse tubules with myofibrils. Dev Biol. 1993;160:135–147.

45. Forbes MS, Sperelakis N. Ruthenium red staining of skeletal and cardiac muscles. Z Zellforsch Cell Tissue Res. 1979;200:367–382.

46. Garcia Mdel C, Gonzalez-Serratos H, Morgan JP, Perreault CL, Rozycka M. Differential activation of myofibrils during fatigue in phasic skeletal muscle cells. J Musc Res Cell Motil. 1991;12:412–424.

47. Gonzalez-Serratos H. Inward spread of activation in vertebrate muscle fibers. J Physiol (London). 1971;212:777–799.

48. Gonzalez-Serratos H. Graded activation of myofibrils and the effect of diameter on tension development during contractures in isolated skeletal muscle fibers. J Physiol (London). 1975;253:321–339.

49. Gonzalez-Serratos H, Cordoba-Rodriguez R, Matteson DR, Rozycka M. Role of calcium currents in excitation-contraction coupling in developing cultured embryonic amphibian skeletal muscle cells. J Physiol, 1996;:494P.

50. Gonzalez-Serratos H, Somlyo AV, McClellan G, Shuman H, Borrero LM, Somlyo AP. Composition of vacuoles and sarcoplasmic reticulum in fatigued muscle: electron probe analysis. Proc Natl Acad Sci USA. 1978;75:1329–1333.

51. Gonzalez-Serratos H, Valle-Aguilera R, Lathrop DA, Garcia Mdel. Slow inward calcium currents have no obvious role in muscle excitation-contraction coupling. Nature. 1982;298:292–294.

52. Hagiwara S, Naka K. The initiation of spike potential in barnacle-muscle fibers under low intracellular Ca⁺⁺. J Gen Physiol. 1964;48:141–162.

53. Hagiwara S, Nakajima S. Difference in Na and Ca spikes as examined by application of tetrodotoxin, procaine, and manganese ions. J Gen Physiol. 1966;49:793–806.

54. Hagiwara S, Chichibu S, Naka K. The effect of various ions on resting and spike potentials of barnacle muscle fibers. J Gen Physiol. 1964;43:163–179.

55. Harris EJ. Distribution and movement of muscle chloride. J Physiol (London). 1963;166:87–109.

56. Heiny JA, Ashcroft FM, Vergara J. T-system optical signals associated with inward rectification in skeletal muscle. Nature. 1983;301:164–166.

57. Henek M, Nonner W, Stämpfli R. Voltage clamp of a small muscle membrane area by means of a circular sucrose gap arrangement. Pflügers Arch. 1969;313:71–79.

58. Hille B, Campbell DT. An improved vaseline gap voltage clamp for skeletal muscle fibers. J Gen Physiol. 1976;67:265–293.

59. Hodgkin AL, Horowicz P. The influence of potassium and chloride ions on the membrane potential of single muscle fibers. J Physiol (London). 1959;148:127–160.

60. Hodgkin AL, Huxley AF. Currents carried by sodium and potassium ions through the membrane of the giant axon of Loligo. J Physiol (London). 1952;116:449–472.

61. Hodgkin AL, Nakajima S. Analysis of the membrance capacity in frog muscle. J Physiol (London). 1972;221:121–136.

62. Howell JN. Intracellular binding of ruthenium red in frog skeletal muscle. J Cell Biol. 1974;62:242–247.

63. Hoyle G, Burrows M. Correlated physiological and ultrastructural studies on specialized muscles 3 neuromuscular physiology of the power-stroke muscle of the swimming leg of portunus sanguinolentus. J Exp Zool. 1973;185:83–95.

64. Hoyle G, Smith T. Neuromuscular physiology of giant muscle fibres of a barnacle, Balanus nubilus Darwin. Comp Biochem Physiol. 1963;10:219–314.

65. Hoyle G, Wiersma CAG. Excitation at neuromuscular junctions in Crustacea. J Physiol. 1958;143 493–425.

66. Hurnak O, Zachar J. Maxi chloride channels in L6 myoblasts. Gen Physiol Biophys. 1992;11:389–400.

67. Hussain M, Wareham AC, Head SI. Mechanism of action of a K⁺ channel activator BRL 38227 on ATP-sensitive K⁺ channels in mouse skeletal muscle fibres. J Physiol., 478 Pt. 1994;3:523–532.

68. Huxley AF, Taylor RE. Local activation of striated muscle fibres. J Physiol (London). 1958;144:426–441.

69. Huxley HE, Page S, Wilkie DR. Appendix An electron microscopic study of muscle in hypertonic solutions. J Physiol (London). 1963;169:312–329.

70. Iannaccone ST, Li K-X, Sperelakis N, Lathrop DA. Insulin-induced hyperpolarization in mammalian skeletal muscle. Am J Physiol. 1989;256:C368–C374.

71. Jiang C, Qu Z, Xu H. Gating of inward rectifier K(+) channels by proton-mediated interactions of intracellular protein domains. Trends Cardiovasc Med. 2002;12:5–13.

72. Kakei M, Noma A, Shibasaki T. Properties of adenosine-triphosphate-regulated potassium channels in guinea-pig ventricular cells. J Physiol. 1985;363:441–462.

73. Kennedy D, Evoy WH, Fields. HL. The unit bases of some crustacean reflexes. Symp Soc Exp Biol. 1966;20:75–109.

74. Kerr LM, Sperelakis N. Effects of the calcium antagonists verapamil and bepridil (CERM-1978) on Ca²⁺-dependent slow action potentials in frog skeletal muscle. J Pharmacol Exp Ther. 1982;222:80–86.

75. Keynes RD, Steinhardt RA. The components of the sodium efflux in frog muscle. J Physiol (London). 1968;198:581–599.

76. Khan AR. Influence of ethanol and acetaldehyde on electromechanical coupling of skeletal muscle fibres. Acta Physiol Scand. 1981;111:425–430.

77. Kirby AC, Lindley BD, Picken JR. Calcium content and exchange in frog skeletal muscle. J Physiol (London). 1975;253:37–52.

78. Kulczycky S, Mainwood GW. Evidence for a functional connection between the sarcoplasmic reticulum and the extracellular space in frog sartorius muscle. Can J Physiol Pharmacol. 1972;50:87–98.

79. Li K-X, Sperelakis N. Electrogenic Na-K pump current in rat skeletal myoballs. J Cell Physiol. 1994;159:181–186.

80. Lueck JD, Rossi AE, Thornton CA, Campbell KP, Dirksen RT. Sarcolemmal-restricted localization of functional ClC-1 channels in mouse skeletal muscle. J Gen Physiol. 2010;136:597–613.

81. Lynch III C. Kinetic and biochemical separation of potassium currents in frog striated muscle PhD thesis. New York: University of Rochester; 1978.

82. Mathias RT, Levis RA, Eisenberg RS. Electrical models of excitation-contraction coupling and charge movement in skeletal muscle. J Gen Physiol. 1980;76:1–31.

83. Miledi R, Parker RI, Schalow G. Measurement of calcium transients in frog muscle by the use of arseno III. Proc R Soc London (Biol). 1977;198:201–210.

84. Miledi R, Stefani E, Steinbach AB. Induction of the action potential mechanism in slow muscle fibres of the frog. J Physiol (London). 1971;217:737–754.

85. Mobley BA, Eisenberg BR. Sizes of components in frog skeletal muscle measured by methods of stereology. J Gen Physiol. 1975;66:31–45.

86. Moody-Corbett F, Gilbert R, Akbarali H, Hall J. Calcium current in embryonic Xenopus muscle cells in culture. Can J Physiol Pharmacol. 1989;67:1259–1264.

87. Nakajima S, Gilai A. Action potentials of isolated single muscle fibers recorded by potential-sensitive dyes. J Gen Physiol. 1980;76:729–750.

88. Nassar-Gentina V, Passonneau JV, Vergara JL, Rapoport SI. Metabolic correlates of fatigue and of recovery from fatigue in single frog muscle fibers. Gen Physiol. 1978;72:593–606.

89. Nastuk WL, Hodgkin AL. The electrical activity of single muscle fibres. J Cell Comp Physiol. 1950;35:39–74.

90. Natori R. Propagated contractions in isolated sarcolemma-free bundle of myofibrils. Jikeidai Med J. 1965;12:214–221.

91. Nicola-Siri L, Sanchez JA, Stefani E. Effect of glycerol treatment on calcium current of frog skeletal muscle. J Physiol (London). 1980;305:87–96.

92. Ortega A, Gonzalez-Serratos H, Lepock J. Effect of organic calcium channel blocker D-600 on sarcoplasmic reticulum calcium uptake in skeletal muscle. Am J Physiol. 1997;272:C310–C317.

93. Overton E. Beitrage zur allgemeinen muskel- und nervenphysiologie I Abh I Ueber die osmotischen figenschatten der muskeln. Pflugers Arch. 1902;92:115–280.

94. Peachey LD. The sarcoplasmic reticulum and transverse tubules of the frog’s sartorius. J Cell Biol. 1965;25:209–231.

95. Peachey LD, Huxley AF. Transverse tubules in crab muscles. J Cell Biol. 1964;23:70A.

96. Pilgrim RIC, Wiersma CAG. Observations on the skeletal and somatic musculature of the abdomen and thorax of Procambaru clarkii (Girard) with notes on the thorax of Panulirus interruptus and Astacus. J Morphol. 1963;113:453–487.

97. Podolsky RJ, Costantin LL. Regulation by calcium of the contraction and relaxation of muscle fibers. Fed Proc. 1964;23:933–939.

98. Potreau D, Raymond G. Calcium-dependent electrical activity and contraction of voltage-clamped frog single muscle fibers. J Physiol (London). 1980;307:9–22.

99. Qu Z, Yang Z, Cui N, et al. Gating of inward rectifier K+ channels by proton-mediated interactions of N- and C-terminal domains. J Biol Chem. 2000;275:31573–31580.

100. Rasgado-Flores H, DeSantiago J, Espinosa-Tanguma R. Stoichiometry and regulation of the Na-Ca exchanger in barnacle muscle cells. Ann NY Acad Sci. 1991;639:22–33.

101. Rasgado-Flores H, Santiago EM, Blaustein MP. Kinetics and stoichiometry of coupled Na efflux and Ca influx (Na/Ca exchange) in barnacle muscle cells. J Gen Physiol. 1989;93:1219–1241.

102. Renaud JM. The effect of lactate on intracellular pH and force recovery of fatigued sartorius muscles of frog, Rana pipiens. J Physiol. 1989;416:31–47.

103. Rogus E, Zierler KL. Sodium and water contents of sarcoplasm and sarcoplasmic reticulum in rat skeletal muscle: Effects of anisotonic media, ouabain, and external sodium. J Physiol (London). 1973;233:227–270.

104. Rubio R, Sperelakis N. Penetration of horseradish peroxidase into the terminal cisternae of frog skeletal muscle fibers and blockade of caffeine contracture by Ca⁺⁺ depletion. Z Zellforsch. 1972;124:57–71.

105. Rulon R, Hermsmeyer K, Sperelakis N. Regenerative action potentials induced in the neurogenic heart of Limulus polyphemus. Comp Biochem Physiol. 1971;39A:333–335.

106. Sanchez JA, Stefani E. Inward calcium current in twitch muscle fibers of the frog. J Physiol (London). 1978;283:197–209.

107. Sauviat M-P, Ecault E, Faivre J-F, Finlay I. Activation of ATP-sensitive K channels by a K channel opener (SR 44866) and the effect upon electrical and mechanical activity of frog skeletal muscle. Pflügers Arch. 1991;418:261–265.

108. Sellin LC, Sperelakis N. Decreased potassium permeability in dystrophic mouse skeletal muscle. Exp Neurol. 1978;62:609–617.

109. Spector I, Prives JM. Development of electrophysiological and biochemical membrane properties during differentiation of embryonic skeletal muscle in culture. Proc Natl Acad Sci USA. 1977;74:5166–5170.

110. Sperelakis N. Changes in conductance of frog sartorius fibers produced by CO₂, ReO₄, and temperature. Am J Physiol. 1969;217:1069–1075.

111. Sperelakis N. Origin of the cardiac resting potential. In: Berne R, Sperelakis N, eds. Handbook of Physiology, the Cardiovascular System, Vol 1: the Heart. Bethesda: American Physiological Society; 1979;:187–267.

112. Sperelakis N, Gonzalez-Serratos H. Skeletal muscle action potentials. In: Sperelakis N, ed. Cell Physiology Sourcebook. San Diego: Academic Press; 2001;:865–886.

113. Sperelakis N, Mayer G, Macdonald R. Velocity of propagation in vertebrate cardiac muscles as functions of tonicity and [K+]. Am J Physiol. 1970;219:952–963.

114. Sperelakis N, Rubio R. Ultrastructural changes produced by hypertonicity in cat cardiac muscle. J Mol Cell Cardiol. 1971;3:139–156.

115. Sperelakis N, Schneider MF. Membrane ion conductances of frog sartorius fibers as a function of tonicity. Am J Physiol. 1968;215:723–729.

116. Sperelakis N, Forbes MS, Rubio R. The tubular systems of myocardial cells: ultrastructure and possible function. In: Dhalla NS, Rona G, eds. Recent Advances in Studies on Cardiac Structure and Metabolism, Myocardial Biology, Vol 4. Baltimore: University Park Press; 1974;:163–194.

117. Sperelakis N, Schneider MF, Harris EJ. Decreased K⁺ conductance produced by Ba⁺⁺ in frog sartorius fibers. J Gen Physiol. 1967;50:1565–1583.

118. Sperelakis N, Shigenobu K, Rubio R. ³H-Sucrose compartments in frog skeletal muscle relative to sarcoplasmic reticulum. Am J Physiol. 1978;234:C181–C190.

119. Sperelakis N, Valle R, Orozco C, Martinez-Palomo A, Rubio R. Electromechanical uncoupling of frog skeletal muscle by possible change in sarcoplasmic reticular content. Am J Physiol. 1973;225:793–800.

120. Spitzer NC. Ion channels in development. Annu Rev Neurosci. 1979;2:363–397.

121. Spruce AE, Standen NB, Stanfield PR. Voltage-dependent ATP-sensitive potassium channels of skeletal muscle membrane. Nature. 1985;316:736–738.

122. Spruce AE, Standen NB, Standfield PR. Studies of the unitary properties of adenosine-5′-triphospate-regulated potassium channels of frog skeletal muscle. J Physiol. 1987;382:213–236.

123. Standen NB, Stanfield PR, Ward TA. Properties of single potassium channels in vesicles formed from the sarcolemma of frog skeletal muscle. J Physiol. 1985;364:339–358.

124. Stanfield PR. A calcium-dependent inward current in frog skeletal muscle fibers. Pflügers Arch. 1977;368:267–270.

125. Stephenson EW. Activation of fast skeletal muscle: contributions of studies on skinned fibers. Am J Physiol. 1981;240:C1–19.

126. Steinmeyer K, Ortland C, Jentsch TJ. Primary structure and functional expression of a developmentally regulated skeletal muscle chloride channel. Nature. 1991;354:301–304.

127. Stoller D, Kakkar R, Smelley M, et al. Mice lacking sulfonylurea receptor 2 (SUR2) ATP-sensitive potassium channels are resistant to acute cardiovascular stress. J Mol Cell Cardiol. 2007;43:445–454.

128. Tasker P, Simon SE, Johnstons BM, Shankly KH, Shaw FH. The dimensions of the extracellular space in sartorius muscle. J Gen Physiol. 1959;43:39–53.

129. Thabet M, Miki T, Seino S, Renaud JM. Treadmill running causes significant fiber damage in skeletal muscle of KATP channel-deficient mice. Physiol Genomics. 2005;22:204–212.

130. Vinogradova NA. Distribution of nonpenetrating sugars in the frog’s sartorius muscle under hypo- and hypertonic conditions. Tsitologiya. 1968;10:831–838.

131. Vivaudou MB, Arnoult C, Villaz M. Skeletal muscle ATP-sensitive K⁺ channels recorded from sarcolemmal blebs of split fibers: ATP inhibition is reduced by magnesium and ADP. J Membr Biol. 1991;122:165–175.

132. Vogel S, Harder D, Sperelakis N. Ca⁺⁺ dependent electrical and mechanical activities in skeletal muscle. Fed Proc. 1978;37:517.

133. Wallace N, Sommer JR. Fusion of sarcoplasmic reticulum with ruthenium red. Proc Electron Microsc Soc A. 1975;33:500–501.

134. Weiss DS, Magleby KL. Voltage-dependent gating mechanism for single fast chloride channels from rat skeletal muscle. J Physiol (London). 1992;453:279–306.

135. Werman R, Grundfest H. Graded and all-or-none electrogenesis in arthropod muscle II The effects of alkali-earth and strontium ions on lobster muscle fibers. J Gen Physiol. 1961;44:997–1027.

136. Werman R, McCann FV, Grundfest H. Graded and all-or-none electrogenesis in arthropod muscle I The effects of alkali-earth cations on the neuromuscular system of Romalea microptera. J Gen Physiol. 1961;44:979–995.

137. Winegrad S. Intracellular calcium movements of frog skeletal muscle during recovery from tetanus. J Gen Physiol. 1968;51:65–83.

138. Woll KH, Lonnendonker U, Neumcke B. ATP-sensitive potassium channels in adult mouse skeletal muscle: different modes of blockage by internal cations, ATP, and tolbutamide. Pflügers Arch. 1989;414:622–628.

139. Yokoshiki H, Sunagawa M, Seki T, Sperelakis N. ATP-sensitive K⁺ channels in pancreatic, cardiac, and vascular smooth muscle cells Invited Review. Am J Physiol. 1998;274:C25–C37.

140. Zifarelli G, Pusch M. CLC chloride channels and transporters: A biophysical and physiological perspective. Rev Physiol Biochem Pharmacol. 2007;158:23–76.

¹ To produce glycerol osmotic shock, about 300 mOsm glycerol is added to Ringer’s solution (Eisenberg and Gage, 1969). The glycerol rapidly permeates into the fiber interior (so the fiber shrinks transiently) and equilibrates. But when the glycerol is washed out, there is a great hypotonic shock produced that disrupts the T-tubules.

² In hypertonic solutions, skeletal muscle fibers shrink (fiber diameter decreases) like a perfect osmometer (but with an osmotically-inactive volume of about 32%), but their T-tubules swell.

Chapter 42

Skeletal Muscle Excitability

Chapter Outline

I Summary