Medical Physiology, 3rd Edition

Skeletal Muscle

Contraction of skeletal muscle is initiated by motor neurons that innervate motor units

The smallest contractile unit of skeletal muscle is a multinucleated, elongated cell called a muscle fiber or myofiber (Fig. 9-1). A bundle of linearly aligned muscle fibers forms a fascicle. In turn, bundles of fascicles form a muscle, such as the biceps. The whole muscle is contained within an external sheath extending from the tendons called the epimysium. Fascicles within the muscle are enveloped by a sheath called the perimysium. Single muscle fibers within individual fascicles are surrounded by a sheath called the endomysium. The highly organized architecture of skeletal muscle fibers and connective tissue allows skeletal muscle to generate considerable mechanical force in a vectorial manner. Beneath the endomysium surrounding each muscle fiber is the plasma membrane of the muscle cell called the sarcolemma. An individual skeletal muscle cell contains a densely arranged parallel array of cylindrical elements called myofibrils. Each myofibril is essentially an end-to-end chain of regular repeating units—or sarcomeres—that consist of smaller interdigitating filaments called myofilaments; these myofilaments contain both thin filaments and thick filaments (see pp. 25–28).

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FIGURE 9-1 Structure of skeletal muscle, from muscle to myofilament.

All skeletal muscle is under voluntary or reflex control by motor neurons of the somatic motor system. Somatic motor neurons are efferent neurons with cell bodies located in the central nervous system (CNS). A single muscle cell responds to only a single motor neuron whose cell body—except for cranial nerves—resides in the ventral horn of the spinal cord. However, the axon of a motor neuron typically branches near its termination to innervate a few or many individual muscle cells. The group of muscle fibers innervated by all of the collateral branches of a single motor neuron is referred to as a motor unit. A whole muscle can produce a wide range of forces and a graded range of shortening by varying the number of motor units excited within the muscle. The innervation ratio of a whole skeletal muscle is defined as the number of muscle fibers innervated by a single motor neuron. Muscles with a small innervation ratio control fine movements involving small forces. For example, fine, high-precision movements of the extraocular muscles that control positioning movements of the eye are achieved via an innervation ratio of as little as ~3 muscle fibers per neuron. Conversely, muscles with a large innervation ratio control coarse movement requiring development of large forces. Postural control by the soleus muscle uses an innervation ratio of ~200. The gastrocnemius muscle, which is capable of developing large forces required in athletic activities such as jumping, has innervation ratios that vary from ~100 to ~1000.

As discussed on pp. 208–210, a motor nerve axon contacts each muscle fiber near the middle of the fiber to form a synapse called the neuromuscular junction. The specialized region of sarcolemma in closest contact with the presynaptic nerve terminal is called the motor end plate. Although skeletal muscle fibers can be artificially excited by direct electrical stimulation, physiological excitation of skeletal muscle always involves chemical activation by release of acetylcholine (ACh) from the motor nerve terminal. Binding of ACh to the nicotinic receptor gives rise to a graded, depolarizing end-plate potential. An end-plate potential of sufficient magnitude raises the membrane potential to the firing threshold and activates voltage-gated Na+ channels (Navs) in the vicinity of the end plate, triggering an action potential that propagates along the surface membrane.

Action potentials propagate from the sarcolemma to the interior of muscle fibers along the transverse tubule network

As action potentials propagate along the surface membrane of skeletal and cardiac muscle fibers, they penetrate into the cell interior via radially oriented, tubular invaginations of the plasma membrane called transverse tubules or T tubules (Fig. 9-2). T tubules plunge into the muscle fiber and surround the myofibrils at two points in each sarcomere: at the junctions of the A and the I bands. A cross section through the A-I junction shows a complex branching array of T tubules penetrating to the center of the muscle cell and surrounding the individual myofibrils. Along its length the tubule associates with two terminal cisternae, which are specialized regions of the sarcoplasmic reticulum (SR). The SR of muscle cells is a specialized version of the endoplasmic reticulum (ER) of noncontractile cells and serves as a storage organelle for intracellular Ca2+. The combination of the T-tubule membrane and its two neighboring cisternae is called a triad junction, or simply a triad.imageN9-1

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FIGURE 9-2 Transverse tubules and SR in skeletal muscle. The transverse tubules (T tubules) are extensions of the plasma membrane, penetrating the muscle cell at two points in each sarcomere: the junctions of the A and I bands.

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Diversity of Mechanisms for Excitation-Contraction Coupling in Skeletal, Cardiac, and Smooth Muscle

Contributed by Ed Moczydlowski

The process by which electrical “excitation” of the surface membrane triggers an increase of [Ca2+]i in muscle is known as excitation-contraction coupling or EC coupling.

Membrane Invaginations … Bringing the Depolarization into the Cell

Skeletal Muscle

The combination of the T-tubule membrane and its two neighboring cisternae is called a triad or triad junction; this structure plays a crucial role in the coupling of excitation to contraction in skeletal muscle.

Cardiac Muscle

Cardiac myocytes have a T-tubule network similar to that of skeletal muscle myofibers except that a single terminal cisterna of the SR forms a dyad junction with the T-tubule rather than a triad junction. Furthermore, T-tubules of cardiac myocytes are located at the Z lines separating sarcomeres rather than at the A-I band junctions.

Smooth Muscle

Smooth muscle, in contrast, has more rudimentary and shallow invaginations of the plasma membrane called caveolae (see Fig. 9-15). Caveolae are considered to be a special form of membrane microdomain called lipid rafts that are enriched in glycosphingolipids and cholesterol and are involved in signal transduction. A peripheral SR compartment of smooth muscle encircles the plasma membrane in close proximity to the caveolae. A larger network of central SR runs along the long axis of the cell. The peripheral SR is involved in local Ca2+ release and interaction with plasma membrane ion channels that mediate electrical excitability, whereas the central SR has a greater role in delivering Ca2+ to intracellular myofilaments for contraction.

Source of Calcium

Although the ultimate intracellular signal that triggers and sustains contraction of skeletal, cardiac, or smooth-muscle cells is a rise in [Ca2+]i, the three types of muscle cells differ substantially in the detailed mechanism by which a depolarization of the sarcolemmal membrane results in a rise in [Ca2+]i. Ca2+ can enter the cytoplasm from the extracellular space through voltage-gated ion channels, or alternatively, Ca2+ can be released into the cytoplasm from the intracellular Ca2+ storage reservoir of the SR. Thus, both extracellular and intracellular sources may contribute to the increase in [Ca2+]i. However, the relative importance of these two sources of Ca2+ varies among the different muscle types.

Skeletal Muscle

In skeletal muscle, as noted in the text, the L-type Ca2+ channel (also known as the DHP receptor) in the T tubule directly couples to the SR Ca2+-release channel (also known as the ryanodine receptor, RYR1), which leads to Ca2+ release from the SR and thus a rise in [Ca2+]i.

Cardiac and Smooth Muscle

In contrast to skeletal muscle, in heart and smooth muscle, Ca2+ influx via the voltage-gated Ca2+ channel Cav1.2 directly activates an RYR2 isoform, leading to Ca2+ release from the SR and raising [Ca2+]i. This mechanism of EC coupling known as Ca2+-induced Ca2+ release (CICR) is quite different from the mechanical coupling mechanism of skeletal muscle. In heart and smooth muscle, colocalization of plasma membrane Cav channels with intracellular SR Ca2+-release channels allows for close coupling of Ca2+ entry from the plasma membrane and Ca2+-activation of RYR Ca2+-release channels. In the CICR coupling mechanism, the action of Ca2+ can be considered as analogous to that of a neurotransmitter or chemical messenger that diffuses across a synapse to activate an agonist-gated channel, but in this case the synapse is the intracellular diffusion gap of ~15 nm between surface Cav channels and intracellular RYR channels on the SR membrane. The CICR mechanism serves as a robust amplification system whereby local influx of Ca2+ from small clusters of L-type Cav channels in the plasma membrane trigger the coordinated release of Ca2+, the activation signal for myofilament contraction, from high-capacity internal Ca2+ stores of the SR.

In smooth muscle but not in cardiac muscle, other Ca2+-activated ion channels (e.g., Ca2+-activated K+ channels, and Ca2+-activated Cl channels) also participate in repolarization and regulation of contractile tone. Activation of smooth-muscle contraction also often involves the IP3 receptor (IP3R), another Ca2+-release channel of the ER/SR membrane. In many smooth muscles, a variety of receptor agonists and chemical mediators are coupled to activation of phospholipase C (PLC). PLC activation results in cleavage of PIP2 (phosphatidylinositol 4,5-bisphosphate) and production of IP3, a chemical messenger that activates IP3R-mediated Ca2+ release (see p. 60).

Depolarization of the T-tubule membrane results in Ca2+ release from the SR at the triad

The ultimate intracellular signal that triggers and sustains contraction of skeletal muscle cells is a rise in [Ca2+]i. Ca2+ can enter the cytoplasm from the extracellular space through voltage-gated ion channels or, alternatively, Ca2+ can be released into the cytoplasm from the intracellular Ca2+ storage reservoir of the SR. Thus, both extracellular and intracellular sources may contribute to the increase in [Ca2+]i. The process by which electrical “excitation” of the surface membrane triggers an increase of [Ca2+]i in muscle is known as excitation-contraction coupling or EC coupling.

The propagation of the action potential into the T tubules of the myofiber depolarizes the triad region of the T tubules, as discussed in the previous section, thereby activating L-type Ca2+ channels (see pp. 190–193). These voltage-gated channels cluster in groups of four called tetrads (Fig. 9-3) and have a pivotal role as the voltage sensor EC coupling. Functional complexes of L-type Ca2+ channels contain the α1-subunit of the voltage-gated Ca2+ channel (i.e., Cav1.1) as well as the accessory α2-δ, β, and γ subunits (see Fig. 7-12B). The L-type Ca2+ channel is also often referred to as the DHP receptor because it is inhibited by a class of antihypertensive and antiarrhythmic drugs known as dihydropyridines or calcium channel blockers. Depolarization of the T-tubule membrane produces conformational changes in each of the four Cav1.1 channels of the tetrad, resulting in two major effects. First, the conformational changes open the Cav1.1 channel pore, which allows electrodiffusive Ca2+ entry. Second, and more importantly in skeletal muscle, the voltage-driven conformational changes in the four Cav1.1 channels mechanically activate each of the four directly coupled subunits of another channel—the Ca2+-release channel located in the portion of the terminal cisternae of the SR membrane that faces the T tubule (see Fig. 9-3).

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FIGURE 9-3 EC coupling in skeletal muscle. A tetrad of four L-type Cav channels on the T tubules faces a single RYR1 Ca2+-release channel of the SR, so that each pseudotetrameric Cav channel interacts with the foot of one of the four subunits of the RYR. Note that every other RYR interacts with Cav channels along the T-tubule in a double checkerboard pattern. imageN9-12

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Structure of the RYR1 Ryanodine Receptor

Contributed by Ed Moczydlowski

eFigure 9-4 shows the low-resolution structure of RYR1.

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EFIGURE 9-4 Image of RYR Ca2+-release channel obtained by cryo-electron microscopy at 9.6-Å resolution. A, View from the cytoplasm of the 280 × 280 Å tetrameric structure of the cytoplasmic foot domain. B, Side view showing the location of the membrane domain. TM, transmembrane region. (Reprinted with permission from Serysheva II, Ludtke SJ, Baker ML, et al: Subnanometer-resolution electron cryomicroscopy-based domain models for the cytoplasmic region of skeletal muscle RYR channel. Proc Natl Acad Sci U S A 105[28]:9610–9615, 2008, Fig 1. Copyright 2005 National Academy of Sciences, U.S.A.)

Reference

Serysheva II, Ludtke SJ, Baker ML, et al. Subnanometer-resolution electron cryomicroscopy-based domain models for the cytoplasmic region of skeletal muscle RyR channel. Proc Natl Acad Sci U S A. 2008;105:9610–9615.

The SR Ca2+-release channel (see Fig. 6-20W) has a homotetrameric structure quite different from that of the T-tubule Cav1.1 channel. This SR Ca2+-release channel is also known as the ryanodine receptor (RYR) because it is inhibited by the plant alkaloid ryanodine—an important tool in characterizing RYRs. In contrast, another plant alkaloid, caffeine, which is present in coffee, activates RYRs by increasing opening probability. imageN9-2 RYRs are the largest known channel proteins, with a molecular mass of ~550 kDa for the monomer, or ~2.1 MDa for a homotetramer. Each of the four subunits of these channels has a large extension—also known as a foot—that projects into the cytosol (see Fig. 9-3).

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Caffeine

Contributed by Ed Moczydlowski

As noted on page 230 of the text, the plant alkaloid caffeine, which is present in coffee, activates RYRs by increasing opening probability. Caffeine is often used experimentally as a research tool to open RYRs and deplete SR Ca2+ stores in muscle, but this effect is not related to the potent CNS stimulant effects of caffeine, which are the result of its action as an antagonist of CNS adenosine receptors (see Fig 13-14B).

In skeletal muscle, where the Ca2+-release channels are of the RYR1 subtype, RYR1 tetramers line up in two rows in the SR membrane. In the T-tubule membrane, half as many Cav1.1 channel tetrads are similarly aligned but are spaced such that they make intracellular contact with every other RYR1 in an alternating “double checkerboard” pattern. The monomer foot domain of each of the four RYR1 subunits is complementary to the cytoplasmic projection of one of the four Cav1.1 channels in a tetrad on the T tubule (see Fig. 9-3). The precise geometrical proximity of these two proteins as well as the ability of both DHP and ryanodine to block muscle contraction indicates that mechanical interactions between these two different Ca2+ channels underlie EC coupling in skeletal muscle. Further evidence for a direct physical interaction between Cav1.1 and RYR1 is the observation that many cycles of excitation and contraction can occur in complete absence of extracellular Ca2+. Moreover, Cav1.1 channels in the closed state physically inhibit the opening of RYR1 channels and thereby prevent the spontaneous release of SR Ca2+ in the nonactivated, resting state. Thus, EC coupling in skeletal muscle is an electromechanical process involving a voltage-induced Ca2+ release mechanism.

After depolarization of the L-type Ca2+ channel on the T-tubule membrane and mechanical activation of the Ca2+-release channel in the SR, Ca2+ stored in the SR rapidly leaves through the Ca2+-release channel. When imaged using a fluorescent Ca2+ indicator, the rapid and transient rise in local [Ca2+]i—from clusters of RYR channels—appears as a spark.imageN9-3 This increase in [Ca2+]i activates troponin C, initiating formation of cross-bridges between myofilaments, as described below. EC coupling in skeletal muscle thus includes the entire process we have just described, beginning with the depolarization of the T-tubule membrane to the initiation of the cross-bridge cycle of contraction.

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Calcium Sparks

Contributed by Ed Moczydlowski

The use of advanced fluorescent Ca2+ indicator dyes and confocal microscopy to image Ca2+ signaling in muscle cells has revealed a variety of elementary events observed as brief bursts of fluorescence corresponding to a transient and highly localized increase in intracellular Ca2+. Detailed biophysical studies of these events, termed Ca2+ sparks, has helped to refine understanding of EC coupling in skeletal, cardiac, and smooth-muscle cells.

Ca2+ sparks were first characterized in cardiac myocytes and later also described in smooth muscle and skeletal muscle. Such spark events can be observed in resting cardiac myocytes loaded with a fluorescent Ca2+ dye indicator such as fluo-3 (eFig. 9-1A). The spark is a brief increase in fluorescence intensity corresponding to Ca2+ binding to the dye resulting from a local and rapid increase in Ca2+concentration that rises to a peak within ~10 ms and decays within ~50 ms (see eFig. 9-1B). Such spontaneous sparks in cardiac myocytes are due to the small opening probability of SR RYR Ca2+-release channels that depends upon [Ca2+] in the cytoplasm and SR lumen. Biophysical analysis indicates that a single spark event corresponds to the simultaneous opening of a cluster of Ca2+-release channels, termed calcium release units (CRUs), that may represent the opening of ~10 to 100 RYR channels, depending on the recording conditions and preparation. Although visual resolution of Ca2+ sparks generally requires low activation conditions of Ca2+ release, they can be observed in single cardiac myocytes activated by a depolarizing voltage pulse at the leading edge of a transient rise of [Ca2+]i (see eFig. 9-1A).

eFigure 9-1C shows a series of Ca2+ sparks from a cardiac myocyte imaged by a line scan of a confocal microscope oriented along the long axis of the cell. The recording shows that synchronized voltage-activated Ca2+ sparks appear at the locations of T tubules at a spacing of ~1.8 µm apart. Such experiments have shown that the macroscopic or global increase in cytoplasmic [Ca2+] in muscle cells is the result of the stochastic summation of many individual spark events corresponding to localized bursts of intracellular Ca2+ release. Studies of Ca2+ sparks have also confirmed that voltage-activated mechanical coupling underlies EC mechanisms in skeletal muscle, whereas Ca2+-induced Ca2+ release underlies these mechanisms in cardiac and smooth muscle.

Due to the tight voltage control of Ca2+ release and termination by brief ~2-ms Na+ action potentials in mammalian skeletal muscle, classical Ca2+ sparks in these striated muscle cells can be resolved only after strenuous exercise of the muscle and under certain nonphysiological and pathological conditions. This implies that spontaneous opening of RYRs in skeletal muscle is suppressed by mechanical linkage to Cav channels in the resting state and that mechanical EC coupling of mammalian skeletal muscle involves fine temporal and voltage control of Ca2+ release, which presumably facilitates precise control of many body movements.

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EFIGURE 9-1 Calcium spark imaging in single cardiac ventricular myocytes. A, A recording was made from a myocyte under whole-cell voltage clamp with fluo-3 Ca2+-imaging dye in the pipette (top right diagram). A line scan along the long dimension of the cell was imaged by confocal microscopy. Upon mild voltage depolarization from −50 to −40 mV for 400 ms, randomly appearing spark events (red-orange spots) are observed in the top left time course of the line scan. The lower trace shows the time course of total integrated intracellular Ca2+, which increases in consecutive jumps corresponding to the appearance of Ca2+ sparks seen in the upper image. B, A myocyte loaded with 4 mM EGTA (ethylene glycol-bis(2-aminothylether)-N,N,N′,N′-tetraacetic acid, or egtazic acid) and 1 mM Oregon Green 488 BAPTA-5N Ca2+-imaging dye was imaged by confocal line scan as in A. Here the line scan was placed along a row of T tubules. Depolarization of the cell to 0 mV evoked a series of simultaneous Ca2+ spark events at the T-tubule spacing of ~1.8 mm. The lower trace shows the average time course of Ca2+ sparks corresponding to SR Ca2+-release events. C, Surface plot of line-scan image from another myocyte showing both spontaneous Ca2+ sparks at a resting Vm of −70 mV and a row of Ca2+ sparks evoked by depolarization to −30 mV. (Reprinted with permission from Cheng H, Lederer J: Calcium sparks. Physiol Rev 88:1491–1545, 2008, Figs 2 and 3.)

Although we have stressed that EC coupling in skeletal muscle primarily involves direct mechanical coupling between the L-type Ca2+ channel in the T-tubule membrane and the Ca2+-release channel of the SR, imageN9-1 other mechanisms modulate the activity of RYR1. For example, RYR1 is subject to regulation by cytoplasmic Ca2+, Mg2+, ATP, and calmodulin (CaM) as well as protein kinases such as protein kinase A (PKA; see p. 57) and Ca2+-calmodulin–dependent kinase II (CaMKII; see p. 60). In the fight-or-flight response (see p. 347), the sympathetic autonomic nervous system activates β-adrenergic receptors, causing PKA-mediated phosphorylation of RYR1 and other muscle proteins; this results in faster and larger increases in cytoplasmic Ca2+, and thus stronger skeletal muscle contraction (Box 9-1).

Box 9-1

Defective EC Coupling in Muscle Due to Cav Channel Mutations

Ca2+ channels have been linked to a large variety of genetic defects of skeletal muscle. In mice, an interesting mutation results in muscular dysgenesis, or failure of normal skeletal muscle to develop. These mice lack a functional Ca2+ channel α1 subunit in their skeletal muscle. They die shortly after birth, but their cultured muscle cells provide an assay system to investigate the mechanism of EC coupling. Contraction of such defective muscle cells can be rescued by expression of cloned genes for either the skeletal Cav1.1 (CACNA1S gene) or the cardiac Cav1.2 (CACNA1C gene) L-type Ca2+ channels. A key distinguishing feature of EC coupling in normal skeletal muscle versus cardiac muscle is the requirement for extracellular Ca2+ in cardiac muscle (see pp. 242–243) but not in skeletal muscle (see pp. 242–243). imageN9-1 Indeed, when the rescue is accomplished with skeletal Cav1.1, contraction does not require extracellular Ca2+. On the other hand, when the rescue is accomplished with cardiac Cav1.2, contraction does require extracellular Ca2+. Such studies provide strong support for the concept that EC coupling (1) in skeletal muscle involves direct mechanical coupling of Cav1.1 to the RYR1 but (2) in cardiac muscle involves Ca2+ entry through Cav1.2 channels, which causes Ca2+-induced Ca2+ release (see pp. 242–243). Experiments with chimeric cardiac and skeletal Cav channel isoforms have shown that the intracellular linker region between domains II and III (see Fig. 7-12B) determines whether EC coupling is of the skeletal or cardiac type.

Hypokalemic periodic paralysis (not to be confused with hyperkalemic periodic paralysis, discussed in Box 7-1) is an autosomal dominant muscle disease of humans. Affected family members have a point mutation in the CACNA1S gene encoding the skeletal Cav1.1, located in transmembrane segment S4 of domain II. This finding explains the basis for a human disorder involving defective EC coupling of skeletal muscle.

Striations of skeletal muscle fibers correspond to ordered arrays of thick and thin filaments within myofibrils

Myofilaments are of two types: thick filaments composed primarily of a protein called myosin and thin filaments largely composed of a protein called actin (see pp. 25–28). The sarcomere is defined as the repeating unit between adjacent Z disks or Z lines (Fig. 9-4A, B). A myofibril is thus a linear array of sarcomeres stacked end to end. The highly organized sarcomeres within skeletal and cardiac muscle are responsible for the striped or striated appearance of muscle fibers of these tissues as visualized by various microscopic imaging techniques. Thus, both skeletal muscle and cardiac muscle are referred to as striated muscle. In contrast, smooth muscle lacks striations because actin and myosin have a less regular pattern of organization in these myocytes.

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FIGURE 9-4 Structure of the sarcomere.

In striated muscle, thin filaments—composed of actin—are 5 to 8 nm in diameter and 1 µm in length. The plus end of the thin filaments attach to opposite faces of a dense disk known as the Z disk (see Fig. 9-4B), which is perpendicular to the axis of the myofibril and has the diameter of the myofibril. Cross-linking the antiparallel thin filaments at the Z disk are α-actinin proteins. Each α-actinin is a rod-shaped antiparallel homodimer, 35 nm long and belonging to the spectrin family of actin-binding proteins. Two large proteins, titin and nebulin, are also tethered at the Z disks, as are other diverse proteins thought to be involved in stretch sensing and signal communication to the nucleus. Not only do Z disks tether the thin filaments of a single myofibril together, but connections between the Z disks also tether each myofibril to its neighbors and align the Z disks and thus the sarcomeres. In summary, Z disks have an important protein-organizing and tension-bearing role in the sarcomere structure.

The thick filaments—composed of myosin—are 10 to 15 nm in diameter and, in striated muscle, 1.6 µm in length (see Fig. 9-4B). They lie between and partially interdigitate with the thin filaments. This partial interdigitation results in alternating light and dark bands along the axis of the myofibril. The light bands, which represent regions of the thin filament that do not overlap with thick filaments, are known as I bands because they are isotropic to polarized light as demonstrated by polarization microscopy. The Z disk is visible as a dark perpendicular line at the center of the I band. The dark bands, which represent the myosin filaments, are known as A bands because they are anisotropic to polarized light. When the A band is viewed in cross section where the thick and thin filaments overlap, six thin filaments (actin) are seen to surround each thick filament (myosin) in a tightly packed hexagonal array (see Fig. 9-4C). During contraction, the I bands (nonoverlapping region of actin) shorten, while the A bands (myosin) do not change in length. This observation led to the idea that an energy-requiring ratcheting mechanism causes the thick and thin filaments to slide past each other—the sliding filament model of muscle contraction.

Thin and thick filaments are supramolecular assemblies of protein subunits

Thin Filaments

The backbone of the thin filament is a right-handed, two-stranded helix of noncovalently polymerized actin molecules, forming filamentous or F-actin (Fig. 9-5A). imageN9-4 The fundamental unit is a supramolecular helix with a total of 13 molecules in the two strands and a length of ~36 nm. The muscle thin filament is an association of F-actin with two important regulatory actin-binding proteins: tropomyosin and the troponin complex.

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FIGURE 9-5 Structure of thin and thick filaments. (A, Courtesy of Roberto Dominguez, University of Pennsylvania.)

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F-Actin

Contributed by Ed Moczydlowski

Actin is perhaps the most abundant and highly conserved protein in eukaryotic cells. It is engaged in numerous protein-protein cytoskeletal interactions in the cytoplasm. The 43-kDa, 375-residue, soluble monomer form of actin is called G-actin. Aside from other cellular forms of cytoskeletal actin (see p. 25), there are three human isoforms of α-actin involved in muscle contraction that correspond to separate actin genes expressed in skeletal muscle (ACTA1), smooth muscle (ACTA2), and cardiac muscle (ACTC1). As noted beginning on pages 25–28, binding and hydrolysis of ATP controls polymerization of G-actin into the filamentous form of actin (F-actin) by sequential addition of actin monomers at the plus end of the molecule. Each actin molecule in F-actin interacts with four other actin molecules. The fundamental unit is a supramolecular helix (a double strand) with a total of 13 molecules in the two strands, and a length of ~36 nm (see Fig. 9-5A).

The tropomyosin monomer of striated muscle is an α-helical protein of 284 amino acids, consisting of seven pseudo-repeats of ~40 residues along the length of the molecule. The pseudo-repeats of the monomer determine its linearly coiled shape and define the binding to seven actin monomers along the thin filament. Two tropomyosin monomers form a dimer aligned in parallel and wound about each other in a coiled-coil structure. Two such tropomyosin dimers flank each supramolecular helix of actin (see Fig. 9-5A). Overlapping head-to-tail contacts between two tropomyosin dimers produce two nearly continuous double-helical filaments that shadow the actin double helix. As we describe below, tropomyosin acts as a gatekeeper in regulating the binding of myosin head groups to actin.

Troponin or the troponin complex is a heterotrimer consisting of the following:

1. Troponin T (TnT or TNNT), which binds to a single molecule of tropomyosin

2. Troponin C (TnC or TNNC), which binds Ca2+. Troponin C is closely related to another Ca2+-binding protein, calmodulin (see p. 60).

3. Troponin I (TnI or TNNI), which binds to actin and inhibits contraction.

Thus, each troponin heterotrimer interacts with a single tropomyosin molecule, which in turn interacts with seven actin monomers. The troponin complex also interacts directly with the actin filaments. The coordinated interactions of troponin, tropomyosin, and actin allow the binding of actin and myosin to be regulated by changes in [Ca2+]i.

Thick Filaments

Like actin thin filaments, thick filaments are also an intertwined complex of proteins (see Fig. 9-5B). In fast skeletal muscle, the thick filament is a bipolar superassembly of several hundred myosin II molecules, which are part of a larger family of myosins (see p. 25). Myosin II is responsible for ATP-dependent force generation in all types of myocytes. The myosin II molecule is a pair of identical heterotrimers, each composed of a myosin heavy chain (MHC), and two myosin light chains (MLCs). One MLC is an essential light chain (ELC or MLC-1), imageN9-5 and the other is a regulatory light chain (RLC or MLC-2). Both the MHCs and MLCs vary among muscle types (Table 9-1).

TABLE 9-1

Isoform Expression of Contractile and Regulatory Proteins*

 

SKELETAL SLOW (TYPE I)

SKELETAL FAST OXIDATIVE (TYPE IIa)

SKELETAL FAST FATIGABLE (TYPE IIx/IIb)

CARDIAC

SMOOTH

Myosin heavy chain

MHC-I (MYH1) and βMHC (MYH7)

MHC-IIa (MYH2)

MHC-IIb (MYH4), MHC-IIx (MYH1)

αMHC (MYH6) and βMHC (MYH7)

MHC-SM1, MHC-SM2 (MYH11)

Myosin light chain (essential)

MLC-1aS, MLC-1bS (MYL3)

MLC-1f, MLC-3f (MYL1)

MLC-1f, MLC-3f (MYL1)

MLC-1v, MLC-1a (MYL3)

MLC-17a, MLC-17b (MYL6)

Myosin light chain (regulatory)

MLC-2 (MYL2)

MLC-2fast (MYLPF)

MLC-2fast (MYLPF)

MLC-2v (MYL2), MLC-2a (MYL7)

MLC-2c (MYL9)

SR Ca-ATPase

SERCA2a (ATP2A2)

SERCA1 (ATP2A1)

SERCA1 (ATP2A1)

SERCA2a (ATP2A2)

SERCA2a, SERCA2b (b > > > a) (ATP2A2)

Phospholamban

Present

Absent

Absent

Present

Present

Calsequestrin

CSQ1, CSQ2

CSQ1

CSQ1

CSQ2

CSQ2, CSQ1

Ca2+ release mechanisms

RYR1, Ca2+-release channel or ryanodine receptor (RYR1)

RYR1 (RYR1)

RYR1 (RYR1)

RYR2 (RYR2)

IP3R1, IP3R2, IP3R3 (ITPR1, ITPR2, ITPR3)
RYR3 (RYR3)

Ca2+ sensor

Troponin C1 (TNNC1)

Troponin C2 (TNNC2)

Troponin C2 (TNNC2)

Troponin C1 (TNNC1)

CaM (multiple isoforms)

*Gene names in parentheses.

In normal adult ventricular muscle, αMHC is the dominant form.

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Muscle Myosin

Contributed by Ed Moczydlowski

For historical reasons, the parts of the myosin II molecule in muscle often have more than one name.

• Myosin heavy chains (MHCs) consist of the following:

• The N-terminal head

• A neck or lever (or lever arm) or linker or hinge

• The C-terminal rod or tail

• Essential myosin light chains (ELCs or MLC-1) are also called alkali chains.

• Regulatory myosin light chains (RLCs or MLC-2).

myosin heavy chain molecule has ~2000 amino acids in three regions: imageN9-5 an N-terminal head region, a neck, and a C-terminal rod. The α-helical rod portions of two MHCs wrap around each other to form a dimer; these dimers self-assemble into thick filaments. At the neck regions, the two MHCs of the dimer flare apart, leading to the two globular heads. Each MHC head has, at its tip, several loops that bind actin and, at its middle, a nucleotide site for binding and hydrolyzing ATP.

The essential light chain and regulatory light chain—both structurally related to the CaM superfamily—bind to and mechanically stabilize the α-helical neck region. The phosphorylation of RLC by myosin light-chain kinases (MLCKs)—members of the CaMK family—enhances myosin cross-bridge interactions. Phosphatases have the opposite effect. In skeletal muscle, this phosphorylation is an important mechanism for force potentiation. Figure 9-6 illustrates how Ca2+ triggers the interaction between a thin filament and a myosin head group from a thick filament.

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FIGURE 9-6 Role of Ca2+ in triggering the contraction of skeletal and cardiac muscle.

Running alongside the thick filaments of skeletal muscle is a protein named titin—the largest known protein, with ~25,000 amino acids (~3 MDa). The linear titin molecule spans one half the length of a sarcomere, with its N terminus tethered in the Z disk and its C terminus in the M line (see Fig. 9-4B). Within the M line are other proteins that cross-link the antiparallel myosin molecules at the middle of the thick filament. Titin—the elastic filament of sarcomeres—includes ~300 immunoglobulin-like domains that appear to unfold reversibly upon stretch.

Nebulin is another large protein (600 to 900 kDa) of muscle that runs from the Z disk along the actin thin filaments. Nebulin interacts with actin and controls the length of the thin filament; it also appears to function in sarcomere assembly by contributing to the structural integrity of myofibrils.

During the cross-bridge cycle, contractile proteins convert the energy of ATP hydrolysis into mechanical energy

The fundamental process of skeletal muscle contraction involves a biochemical cycle, called the cross-bridge cycle, that occurs in six steps (Fig. 9-7). We start the cycle in the absence of both ATP and ADP, with the myosin head rigidly attached to an actin filament. In a corpse soon after death, the lack of ATP prevents the cycle from proceeding further; this leads to an extreme example of muscle rigidity—called rigor mortis—that is limited only by protein decomposition.

Step 1: ATP binding. ATP binding to the head of the MHC reduces the affinity of myosin for actin, which causes the myosin head to release from the actin filament. If all cross-bridges in a muscle were in this state, the muscle would be fully relaxed.

Step 2: ATP hydrolysis. The breakdown of ATP to ADP and inorganic phosphate (Pi) occurs on the myosin head; the products of hydrolysis are retained within the myosin active site. As a result of hydrolysis, the myosin head/neck pivots into a “cocked” position in which the head/neck are more colinear with the rod. This pivot causes the tip of the myosin head to move ~11 nm along the actin filament so that it now lines up with a new actin monomer two monomers farther along the actin filament. imageN9-6 If all cross-bridges in a muscle were in this state, the muscle would be fully relaxed.

N9-6

Measuring the Force of a Single Cross-Bridge Cycle

Contributed by Ed Moczydlowski

The force of a single cross-bridge cycle has been measured directly. Finer, Simmons, and Spudich used optical tweezers to manipulate a single actin filament and to place it in proximity to a myosin molecule immobilized on a bead (eFig. 9-2A). With the use of video-enhanced microscopy these investigators were able to detect movements of the actin filament as small as 1 nm. The optical tweezers could also exert an adjustable force opposing movement of the actin filament. When the tweezers applied only a small opposing force and the experiment was conducted in the presence of ATP, the researchers observed that the actin moved over the myosin bead in step-like displacements of 11 nm. This observation, made under “microscopically isotonic” conditions, suggests that the quantal displacement of a single cross-bridge cycle is ~11 nm (see eFig. 9-2B). When the tweezers applied a force sufficiently large to immobilize the actin filament, the investigators observed step-like impulses of force that averaged ~5 pN (see eFig. 9-2C). This observation, made under “microscopically isometric” conditions, suggests that the quantal force developed during a single cross-bridge cycle is ~5 pN. Interestingly, these isometric force impulses lasted longer when the ATP concentration was lower. This last finding is consistent with the notion that ATP binding to myosin must occur to allow detachment of the cross-bridges (step 1 in the cycle in Fig. 9-7).

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EFIGURE 9-2 Microscopic measurements of cross-bridge force and displacement. A, An actin filament is attached at each end to a polystyrene bead. The optical tweezers, a finely focused beam of laser light, can trap the bead at its focal point and physically move it. By adjusting the laser intensity, the experimenter can alter the strength of the trap (i.e., the force with which the bead is held). In this experiment, two optical tweezers were used to suspend the actin filament above a coverglass. Attached to this coverglass is a silica bead, and myosin molecules are bound to the bead. B, In an isotonic experiment, the force between the actin filament and the fixed myosin/silica bead is kept constant by use of a stable laser intensity. The experimenter measures, as a function of time, the displacement of the polystyrene bead away from the center of the trap. Thus, in one cross-bridge cycle, the myosin-actin interaction pulls the polystyrene bead ~11 nm away from the center of the trap. C, In an isometric experiment, the experimenter measures, as a function of time, the extra force that needs to be applied (i.e., increase in laser intensity) to keep the polystyrene bead at a fixed position near the center of the trap. Thus, in one cross-bridge cycle, the myosin-actin interaction exerts a force of ~5 pN. (Data from Finer JT, Mehta AD, Spudich JA: Characterization of single actin-myosin interactions. Biophys J 68:291s–296s, 1995.)

Step 3: Weak cross-bridge formation. The cocked myosin head now binds loosely to a new position on the actin filament, scanning for a suitable binding site. Recall that six actin filaments surround each thick filament.

Step 4: Release of Pi from the myosin. Dissociation of Pi from the myosin head triggers an increased affinity of the myosin-ADP complex for actin—the strong cross-bridge state. The transition from weak to strong binding is the rate-limiting step in the cross-bridge cycle.

Step 5: Power stroke. A conformational change causes the myosin neck to rotate around the myosin head, which remains firmly fixed to the actin. This bending pulls the rod of the myosin, drawing the actin and myosin filaments past one another by a distance of ~11 nm. The myosin head/neck is now angled with respect to the rod. At the macroscopic level, this activity pulls the Z lines closer together and shortens the sarcomere, with concurrent force generation.

Step 6: ADP release. Dissociation of ADP from myosin completes the cycle, and the actomyosin complex is left in a rigid, “attached state.” The relative positions of the actin versus the myosin head, neck, and rod remain the same until another ATP molecule binds and initiates another cycle (step 1).

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FIGURE 9-7 Cross-bridge cycle in skeletal and cardiac muscle. Each cycle advances the myosin head by two actin monomers, or ~11 nm.

The ADP–free myosin complex (“Attached State” in Fig. 9-7) would quickly bind ATP at the concentrations of ATP normally found within cells. Each round of the cross-bridge cycle consumes one molecule of ATP; we discuss the regeneration of ATP in muscle beginning on pages 1208–1209. If unrestrained, this cross-bridge cycling would continue until the cytoplasm is depleted of ATP—rigor mortis.

The biochemical steps of the cross-bridge cycle reveal that [ATP]i does not regulate the cross-bridge cycle of actin-myosin interaction. In skeletal and cardiac muscle, temporal control of the cycle of contraction occurs at the third step by prevention of cross-bridge formation until the tropomyosin moves out of the way in response to an increase in [Ca2+]i—as we will see in the next section.

An increase in [Ca2+]i triggers contraction by removing the inhibition of cross-bridge cycling

In skeletal, cardiac, and smooth muscle, an increase in [Ca2+]i initiates and allows cross-bridge cycling to continue. During this excitatory increase, [Ca2+]i may rise from its resting level of <10−7 M to >10−5 M. The subsequent decrease in [Ca2+]i—discussed in the next section—is the signal to cease cross-bridge cycling and relax.

Regardless of the muscle type, Ca2+ exerts its effect by binding to regulatory proteins rather than directly interacting with contractile proteins. In the absence of Ca2+, these regulatory proteins act in concert to inhibit actin-myosin interactions, thus inhibiting the contractile process. When Ca2+ binds to one or more of these proteins, a conformational change takes place in the regulatory complex that releases the inhibition of contraction. In both skeletal and cardiac muscle, the regulatory proteins form the troponin complex, which is composed of troponin C, troponin I, and troponin T. The troponin T (TnT) binds to tropomyosin, establishing the linkage between the troponin complex and tropomyosin.

In skeletal muscle, the TNNC2 subtype of troponin C (TnC) has two pairs of Ca2+-binding sites. Two high-affinity sites—located on the C-lobe of TNNC2—are always occupied by Ca2+ or Mg2+ under physiological conditions. These sites on TNNC2 bind to troponin I (TnI). On the other hand, two low-affinity sites—located on the N-lobe of TNNC2—bind and release Ca2+ as [Ca2+]i rises and falls. At low [Ca2+]i, the N-lobe of TnC does not bind to TnI (see Fig. 9-6), which allows the TnI to bind to a particular spot on F-actin and thereby prevent the binding of myosin. At high [Ca2+]i, the N-lobe of TnC can now interact with TnI in such a way as to cause tropomyosin to translocate by 25 degrees on the F-actin surface, which allows the cocked myosin head group to interact weakly with actin (see Fig. 9-7, step 3 of the cross-bridge cycle). As long as [Ca2+]i remains high and the tropomyosin is out of the way, cross-bridge cycling will continue indefinitely. imageN9-7

N9-7

Tropomyosin-Troponin Interactions—the “Functional Group” of the Thin Filament

Contributed by Ed Moczydlowski

As discussed in the text, troponin (which consists of troponin T, C, and I) interacts with one tropomyosin molecule, which in turn interacts with seven actin monomers (see Fig. 9-6). The region along a thin filament that falls under the control of a single troponin molecule is a functional group. However, overlap of troponin T onto the junction between two tropomyosin dimers (recall that the tropomyosin molecules stack end to end, with overlap, to create a continuous filament) may allow a single troponin complex to control—via the two tropomyosin dimers—a functional group of 14 or more actin molecules.

In the absence of Ca2+, tropomyosin is bound in a position along the actin filament that blocks its interaction with myosin. When Ca2+ binds to the troponin complex, the tropomyosin shifts from its original position along the actin (eFig. 9-3, red area)—the blocked off-state—to a new position caused by an azimuthal rotation of ~25 degrees on the F-actin surface (see eFig. 9-3, yellow area)—the Ca2+-activated state. This displacement enables myosin to bind to actin, which results in another ~10-degree azimuthal movement of tropomyosin (see eFig. 9-3, green area); this shift allows myosin to engage in mechanical activity—the fully activated state.

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EFIGURE 9-3 Model of the interaction of tropomyosin with actin illustrating the shift in position of tropomyosin from the off, myosin-blocked state (red), to the Ca2+-activated state (yellow), and to the fully activated state (green), which allows full interaction of the myosin head group. (Reprinted with permission from Brown JH, Zhou Z, Reshetnikova L, et al: Structure of the mid-region of tropomyosin: Bending and binding sites for actin. Proc Natl Acad Sci U S A 102[52]:18878–18883, 2005, Fig 4a. Copyright 2005 National Academy of Sciences, U.S.A.)

Termination of contraction requires re-uptake of Ca2+ into the SR

After the action potential in the skeletal muscle has subsided, Ca2+ must be removed from the sarcoplasm for contraction actually to cease and for relaxation to occur. Removal of Ca2+ from the sarcoplasm occurs by two mechanisms. Ca2+ may be extruded across the cell plasma membrane or sequestered within intracellular compartments (Fig. 9-8).

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FIGURE 9-8 Mechanisms of Ca2+ removal from the cytoplasm.

The cell may extrude Ca2+ by use of either an NCX (Na-Ca exchanger, SLC8 family; see p. 123) or PMCA (plasma membrane Ca-ATPase or pump; see p. 118). Extrusion across the cell membrane, however, would eventually totally deplete the cell of Ca2+ and is therefore a minor mechanism for Ca2+ removal from the cytoplasm. Instead, Ca2+ re-uptake into the SR is the most important mechanism by which the cell returns [Ca2+]i to resting levels. Ca2+ re-uptake by the SR is mediated by a SERCA (sarcoplasmic and endoplasmic reticulum Ca-ATPase or pump; see p. 118). imageN9-8

N9-8

SERCA Isoforms

Contributed by Emile Boulpaep, Walter Boron

SERCA is an acronym for sarcoplasmic and endoplasmic reticulum Ca-ATPase. The energy for Ca pumping comes from the hydrolysis of ATP. As discussed on page 126 in the text (also see imageN5-10), the Ca2+-transporting protein is an E1-E2 (or P-type) ATPase that has a molecular weight of 110 kDa. Three different SERCA isoforms are known.

All of the SERCA isoforms (SERCA1, SERCA2, and SERCA3) are Ca-H exchange pumps. The SERCA2 isoform may be expressed as two alternatively spliced variants. The SERCA1 isoform is expressed in fast-twitch skeletal muscle, which is a subtype of skeletal muscle fibers that contract rapidly (see Table 9-1). The SERCA2a isoform is found in slow-twitch skeletal muscle as well as cardiac and smooth muscle. The SERCA2b isoform is found in smooth-muscle cells; it is also heavily expressed in the ER of nonmuscle cells. Table 9-1 summarizes the distribution of the SERCA isoforms among muscle types.

Note that all of the SR Ca pumps (i.e., SERCAs) are distinct from the Ca pumps in the plasma membrane, which are known as PMCAs (see p. 126).

High [Ca2+] within the SR lumen inhibits the activity of SERCA, an effect attenuated by Ca2+-binding proteins within the SR lumen. These Ca2+-binding proteins buffer the [Ca2+] increase in the SR during Ca2+re-uptake and thus markedly increase the Ca2+ storage capacity of the SR. The principal Ca2+-binding protein in skeletal muscle, calsequestrin (CSQ), is also present in cardiac and some smooth muscle. Calreticulin is a ubiquitous Ca2+-binding protein that is found in particularly high concentrations within the SR of smooth muscle. These proteins have a tremendous capacity to bind Ca2+, with up to 50 binding sites per protein molecule.

CSQ forms oligomers in the SR lumen and is highly localized to the region of the SR immediately beneath the triad junction, where it forms a complex with the Ca2+-release channel and with two other RYR-anchoring proteins—junctin and triadin—and senses free [Ca2+] inside the SR. During high action potential frequency, SR Ca2+ content falls, and dissociation of Ca2+ from CSQ leads to deactivation of RYR, thereby preserving SR Ca2+ (Box 9-2).

Box 9-2

Malignant Hyperthermia and Central Core Disease—RYR Channelopathies

Malignant hyperthermia (MH) affects between 1 in 15,000 children and 1 in 50,000 adults undergoing anesthesia. It is a genetic disorder that may affect as many as 1 in 2000 to 3000 individuals in the general population. Affected individuals are at risk of a potentially life-threatening syndrome on exposure to particular inhalation anesthetic agents, especially halothane, sevoflurane, and desflurane. Administration of the muscle relaxant succinylcholine (see pp. 225–226) can also trigger or exaggerate MH. Onset of MH syndrome in the setting of the operating room is typified by the development of tachypnea (rapid breathing), low plasma [O2], high plasma [CO2], tachycardia (rapid heart rate), and hyperthermia (rising body temperature) as well as by rigidity, sweating, and dramatic swings in blood pressure. The patient's temperature may rise as rapidly as 1°C every 5 minutes. The onset of MH is usually during anesthesia, but it can occur up to several hours later. If the condition is untreated, the patient will develop respiratory and lactic acidosis, muscle rigidity, and a breakdown of muscle tissue that leads to the release of K+ and thus profound hyperkalemia. These episodes reflect a progressively severe hypermetabolic state in the muscle tissues. Fortunately, our evolving understanding of the pathophysiology of MH has led to the development of a therapeutic regimen that has greatly improved the once-dismal prognosis.

The major features of the syndrome—hyperthermia, muscle rigidity, and an increased metabolic rate—led early investigators to suggest that MH is a disease of abnormal regulation of muscle contraction. We now understand that uncontrolled Ca2+ release—somehow triggered by the administration of halothane and succinylcholine—causes excessive contraction and ATP hydrolysis. As muscle tries to replenish its ATP stores, mitochondrial oxidative metabolism increases. Hyperthermia develops because of the heat liberated by these metabolic processes.

The incidence of MH is particularly high in swine, in which episodes are triggered by a variety of physical and environmental stresses (porcine stress syndrome). MH in animals has significant economic importance in view of the potential loss from fatal episodes and the devaluation of meat as a result of muscle destruction during nonfatal episodes.

When exposed to halothane, muscle biopsy samples from susceptible individuals develop more tension than fibers from normal individuals. In muscle fibers from both humans and a strain of swine susceptible to MH, Ca2+-induced Ca2+ release from the SR is enhanced compared with that in fibers from unaffected subjects. Furthermore, caffeine, which causes the Ca2+-release channels to open, induced greater contractions in fibers from susceptible subjects. Taken together, these observations suggested that MH results from an abnormality in the Ca2+-release channel in the SR membrane.

In both humans and animals, inheritance of MH follows a mendelian autosomal dominant pattern. Approximately 200 mutations in the human RYR1 have been linked to MH and central core disease (discussed below). The mutations tend to cluster in three “hot spots” of the channel protein: the N-terminal, central, and C-terminal regions. Some of these mutations increase the sensitivity of the RYR1 to activation by halothane and caffeine. They may also act by a “gain-of-function” effect that promotes the constitutive leakage of Ca2+ from the SR. In swine, MH results from a single amino-acid substitution in RYR1 (cysteine for arginine at position 615), the skeletal muscle isoform. The analogous R614C substitution is present in some human kindreds as well. This substitution increases the open probability of the Ca2+-release channel.

Therapy for MH involves intravenous administration of the drug dantrolene, cessation of anesthesia, and aggressive efforts aimed at cooling the body. Dantrolene is an effective therapeutic agent because it blocks RYR1, thus interrupting the otherwise uncontrolled release of SR Ca2+ and progression of muscle contractions. The drug can be given acutely in an effort to abort an ongoing attack or, in a person known to be at risk, it can be given before the initiation of anesthesia to prevent onset of the syndrome. Dantrolene treatment has decreased mortality due to MH from 80% to <10% today. Therapy also includes intravenous hydration and the judicious use of diuretics to keep the urine flowing; this lessens damage to the kidneys from the release of breakdown products, such as myoglobin from the damaged muscles. Sodium bicarbonate is given to counter the lactic acidosis, and patients may be mechanically hyperventilated to blow off the excess CO2. For patients known to be at risk of MH, alternative forms of pain suppression that can be safely used for necessary surgical procedures include nitrous oxide, propofol, opiates, and benzodiazepines. Local anesthetics such as lidocaine are also safe.

Despite the intensive protocol just outlined, MH is still regarded as an emergency in the operating room. The relatives of a patient with a documented history of one episode of MH should be carefully screened to see whether they, too, carry the inherited trait; many of the affected relatives may demonstrate baseline elevations in muscle enzyme levels in their blood (e.g., an increase in creatine kinase levels).

As noted above, mutations in RYR1 have also been linked to central core disease (CCD), also called central core myopathy. CCD is a congenital autosomal dominant genetic condition defined by the histopathological appearance of so-called core regions within skeletal muscle cells that are devoid of mitochondria and are inactive with respect to oxidative metabolism. Symptoms of CCD include muscle weakness in young patients and skeletal abnormalities. Some CCD patients are also susceptible to MH.

Muscle contractions produce force under isometric conditions and force with shortening under isotonic conditions

The total force generated by a muscle is the sum of the forces generated by many independently cycling actin-myosin cross-bridges. The number of simultaneously cycling cross-bridges depends substantially on the initial length of the muscle fiber and on the pattern or frequency of muscle cell stimulation. When muscle is stimulated to contract, it exerts a force tending to pull the attachment points at either end toward each other. This force is referred to as the tension developed by the muscle.

Two mechanical—and artificial—arrangements can be used to study force and length relationships in muscle contraction. In one, the attachment points are immobile, so that the muscle length is fixed. Here, stimulation causes an increase in tension, but no shortening. Because these contractions occur at constant length, they are referred to as isometric contractions (Fig. 9-9A). In the second arrangement, one of the two attachment points is mobile and tethered to a variable load, which tends to pull this mobile point away from the fixed one. Here, stimulation causes shortening, provided the tension developed by the muscle is greater than the opposing load. Because these shortenings occur at constant load, they are referred to as isotonic contractions (see Fig. 9-9B). imageN9-6 Both isometric and isotonic contractions can be examined at different initial muscle lengths. Moreover, they can be measured during individual muscle twitches evoked by single muscle action potentials as well as during other patterns of stimulation. Physical activity involves various combinations of isometric and isotonic contractions.

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FIGURE 9-9 Isometric and isotonic contraction. A, Experimental preparation for study of muscle contraction under isometric conditions. B, Experimental preparation for study of muscle contraction under isotonic conditions. C, The “Passive” curve represents the tension that is measured at various muscle lengths before muscle contraction. The “Total” curve represents the tension that is measured at various muscle lengths during muscle contraction. Muscle length is expressed as the percent of “optimal” length; that is, the length at which active isometric tension is maximal. D, The active tension is the difference between the total and the passive tensions in C. E, Points A, A1, A2, and A3 on the blue curve show that the velocity of muscle shortening is greater if the muscle lifts a lighter weight—it is easier to lift a feather (left side of each curve/low load) than to lift a barbell (right side of each curve/high load). The blue, green, and gold curves also show that for any given velocity of shortening, a longer muscle can develop a greater tension than can a shorter muscle.

Muscle length influences tension development by determining the degree of overlap between actin and myosin filaments

The isometric force of contractions depends on the initial length of the muscle fiber. Unstimulated muscle may be elongated somewhat by applying tension and stretching it. The tension measured before muscle contraction is referred to as passive tension (see Fig. 9-9C). Because muscle gets stiffer as it is distended, it takes increasing amounts of passive tension to progressively elongate the muscle cell. If at any fixed length (i.e., isometric conditions) the muscle is stimulated to contract, an additional active tension develops because of cross-bridge cycling. The total measured tension is thus the sum of the passive and active tension. This incremental or active tension—the difference between total tension and passive tension—is quite small when the muscle is less than ~70% of its normal resting length (see Fig. 9-9D). As muscle length increases toward its normal length, active tension increases. Active tension is maximal at an optimal length (blue point in Fig. 9-9D)—usually called L0—that is near the normal muscle length. Active tension decreases with further lengthening; thus, active tension is again small when the muscle is stretched beyond 150% of its normal resting length. Although the relationship between muscle length and tension has been best characterized for skeletal muscle, the tension of cardiac and smooth muscle also appears to depend on length in a similar manner.

This length-tension relationship is a direct result of the anatomy of the thick and thin filaments within individual sarcomeres (see Fig. 9-9D). As muscle length increases, the ends of the actin filaments arising from neighboring Z disks are pulled away from each other. When length is increased beyond 150% of the resting sarcomere length, the ends of the actin filaments are pulled beyond the ends of the myosin filaments. Under this condition, no interaction occurs between actin and myosin filaments and hence no active tension develops. As muscle length shortens from this point, actin and myosin filaments begin to overlap and tension can develop; the amount of tension developed corresponds to the degree of overlap between the actin and the myosin filaments. As the muscle shortens further, opposing actin filaments slide over one another and the ends of the myosin filaments and—with extreme degrees of shortening—eventually butt up against the opposing Z disks. Under these conditions, the spatial relationship between actin and myosin is distorted and active tension falls. The maximal degree of overlap between actin and myosin filaments, and hence maximal active tension, corresponds to a sarcomere length that is near its normal resting length.

At higher loads, the velocity of shortening is lower because more cross-bridges are simultaneously active

Under isotonic conditions, the velocity of shortening decreases as the applied load opposing contraction of the muscle fiber increases. This point is obvious; anyone can lift a single French fry much faster than a sack of potatoes. As shown for any of the three downward-sloping curves in Figure 9-9E—each of which represents a different initial length of muscle—there is an inverse relationship between velocity and load. Note that for isotonic contractions, the applied load is the same as the tension in the muscle.

The load (or tension)–velocity relationship is perhaps best understood by first considering the condition at points A, B, and C in Figure 9-9E, where—for each muscle length—we choose a different load that is just adequate to prevent shortening or lengthening (i.e., isometric conditions). Point A represents the optimal length for supporting the largest isometric tension (represented by the blue point in Fig. 9-9D). Here, all available cross-bridges are engaged in resisting the opposing force, and none is left over to make the muscle shorten. If we could somehow decrease the number of engaged cross-bridges, the muscle would lengthen. On the other hand, starting at point A, if we decrease the load at the same initial muscle length, the number of cross-bridges already engaged is now more than needed to resist the opposing load. The extra cross-bridges are available to ratchet the thick myosin filaments over the thin actin filaments, but at a very low initial velocity (point A1). Returning to the isometric condition at point A, if we were to reduce the lower load even more, then even more cross-bridges would be available for ratcheting the myosin over the actin, and the initial velocity would be even higher (point A2). Again returning to the isometric condition at point A, if we make the load vanishingly small, the initial velocity is very high because the speed with which the thick and thin filaments slide over each other is limited only by the speed at which the cross-bridges can cycle. Viewed from a different perspective, as we increase velocity, the probability of actin-myosin interactions decreases, fewer cross-bridges are simultaneously active, and less tension develops.

Recall that the blue curve in Figure 9-9E applies to a single initial length of the muscle, that is, the optimal length. We already saw in Figure 9-9D that, starting at the optimal length, reducing initial muscle length causes isometric tension to fall. The points A, B, and C in Figure 9-9E restate this relationship: the shorter the initial length, the smaller the maximal load under zero-velocity conditions. As we decrease the load, starting from point B (green curve) or point C (brown curve), the initial velocity of shortening rises, as we saw for the blue curve. Note that all three curves extrapolate to the same initial maximal velocity at zero load. The explanation for this effect, as already noted, is that maximal velocity (at no load) depends on the maximal rate of cross-bridge turnover, not on the initial overlap of the thin and thick filaments.

The velocity-tension curve reveals an interesting relationship between muscle power and applied load. Muscle does measurable mechanical work only when it displaces a load. This mechanical work (W) is the product of load (F) and displacement (Δx). Power (P) is the rate at which work is performed, or work per unit time (Δt):

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(9-1)

Because velocity (v) is Δxt, it follows that

image

(9-2)

For a given load (F), we can calculate the power by reading the velocity (v) from the uppermost of the three blue load-velocity relationships in Figure 9-9E. Power is maximal at intermediate loads (where both F and v are moderate) and falls to zero at maximum load (where v = 0) and at zero load (where F = 0).

In a single skeletal muscle fiber, the force developed may be increased by summing multiple twitches in time

At sufficiently low stimulation frequencies, the tension developed falls to the resting level between individual twitches (Fig. 9-10A). Single skeletal muscle twitches last between 25 and 200 ms, depending on the type of muscle. Although each twitch is elicited by a single muscle action potential, the duration of contraction is long compared with the duration of the exciting action potential, which lasts only several milliseconds. Because the muscle twitch far exceeds the duration of the action potential, it is possible to initiate a second action potential before a first contraction has fully subsided. When this situation occurs, the second action potential stimulates a twitch that is superimposed on the residual tension of the first twitch and thereby achieves greater isometric tension than the first (compare Fig. 9-10A and 9-10B). This effect is known as summation.

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FIGURE 9-10 Frequency summation of skeletal muscle twitches.

If multiple action potentials occur close enough in time, the multiple twitches can summate and thus greatly increase the tension developed. Summation is more effective at increasing tension when the action potentials are grouped more closely in time, as in Figure 9-10C. In other words, tension is higher when action potentials are evoked at higher frequency. Because this type of tension enhancement depends on the frequency of muscle stimulation, it is referred to as frequency summation.

When the stimulation frequency is increased sufficiently, the individual twitches occur so close together in time that they fuse (see Fig. 9-10D) and cause the muscle tension to remain at a steady plateau. The state in which the individual twitches are no longer distinguishable from each other is referred to as tetanus. Tetanus arises when the time between successive action potentials is insufficient to return enough Ca2+ to the SR to lower [Ca2+]i below a level that initiates relaxation. In fact, a sustained increase in [Ca2+]i persists until the tetanic stimulus ceases. At stimulation frequencies above the fusion frequency that causes tetanus, muscle fiber tension increases very little.

In a whole skeletal muscle, the force developed may be increased by summing the contractions of multiple fibers

In addition to determining the frequency with which it stimulates a single muscle fiber, the CNS can control muscle force by determining the number of individual muscle fibers that it stimulates at a given time. As each additional motor-neuron cell body within the spinal cord is excited, those muscle fibers that are part of the motor unit of that motor neuron are added to the contracting pool of fibers (Fig. 9-11). This effect is known as multiple-fiber summation. In general, smaller motor neurons serve motor units consisting of fewer individual muscle fibers. Because a given excitatory stimulus will generate a larger excitatory postsynaptic potential (see p. 210) in motor neurons with smaller cell bodies, the small motor units are recruited even with minimal neuronal stimulation. As neuronal stimulation intensifies, larger motor neurons innervating larger motor units are also recruited. The progressive recruitment of first small and then larger and larger motor units is referred to as the size principle (see pp. 1204–1205). The group of all motor neurons innervating a single muscle is called a motor-neuron pool.

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FIGURE 9-11 The motor unit and the motor-neuron pool.

Multiple-fiber summation, sometimes referred to as spatial summation, is an important mechanism that allows the force developed by a whole muscle to be relatively constant in time. It is true that the CNS could direct the force to be relatively constant over time merely by driving a fixed number of motor units within the muscle to tetanus, where the force fluctuations are very small (see Fig. 9-10D). However, adding tetanic motor units would increase total muscle force by rather large individual increments. Instead, the CNS can activate individual motor units asynchronously so that some units are developing tension while others are relaxing. Thus, whole-muscle force can be relatively constant with time, even when individual fibers are not stimulated to tetanus. Smooth, nontetanic contraction is essential for fine motor control.