Cellular Physiology of Skeletal, Cardiac, and Smooth 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 ).

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 Ca ²⁺ . The combination of the T-tubule membrane and its two neighboring cisternae is called a triad junction, or simply a triad. N9-1

Depolarization of the T-tubule membrane results in Ca ²⁺ release from the SR at the triad

The ultimate intracellular signal that triggers and sustains contraction of skeletal muscle cells is a rise in [Ca ²⁺ ] _i . Ca ²⁺ can enter the cytoplasm from the extracellular space through voltage-gated ion channels or, alternatively, Ca ²⁺ can be released into the cytoplasm from the intracellular Ca ²⁺ storage reservoir of the SR. Thus, both extracellular and intracellular sources may contribute to the increase in [Ca ²⁺ ] _i . The process by which electrical “excitation” of the surface membrane triggers an increase of [Ca ²⁺ ] _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 Ca ²⁺ 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 in EC coupling. Functional complexes of L-type Ca ²⁺ channels contain the α ₁ -subunit of the voltage-gated Ca ²⁺ channel (i.e., Cav1.1) as well as the accessory α ₂ -δ, β, and γ subunits (see Fig. 7-12 B ). The L-type Ca ²⁺ 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 Ca ²⁺ 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 Ca ²⁺ -release channel located in the portion of the terminal cisternae of the SR membrane that faces the T tubule (see Fig. 9-3 ).

N9-12

Structure of the RYR1 Ryanodine Receptor

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

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: pp. 9610-9615.

The SR Ca ²⁺ -release channel (see Fig. 6-20 W ) has a homotetrameric structure quite different from that of the T-tubule Cav1.1 channel. This SR Ca ²⁺ -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. N9-2 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 ).

In skeletal muscle, where the Ca ²⁺ -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 Ca ²⁺ 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 Ca ²⁺ . Moreover, Cav1.1 channels in the closed state physically inhibit the opening of RYR1 channels and thereby prevent the spontaneous release of SR Ca ²⁺ in the nonactivated, resting state. Thus, EC coupling in skeletal muscle is an electromechanical process involving a voltage-induced Ca ²⁺ release mechanism.

After depolarization of the L-type Ca ²⁺ channel on the T-tubule membrane and mechanical activation of the Ca ²⁺ -release channel in the SR, Ca ²⁺ stored in the SR rapidly leaves through the Ca ²⁺ -release channel. When imaged using a fluorescent Ca ²⁺ indicator, the rapid and transient rise in local [Ca ²⁺ ] _i —from clusters of RYR channels—appears as a spark. N9-3 This increase in [Ca ²⁺ ] _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.

N9-3

Calcium Sparks

The use of advanced fluorescent Ca ²⁺ indicator dyes and con­focal microscopy to image Ca ²⁺ 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 Ca ²⁺ . Detailed biophysical studies of these events, termed Ca ²⁺ sparks, has helped to refine understanding of EC coupling in skeletal, cardiac, and smooth-muscle cells.

Ca ²⁺ 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 Ca ²⁺ dye indicator such as fluo-3 ( eFig. 9-1 A ). The spark is a brief increase in fluorescence intensity corresponding to Ca ²⁺ binding to the dye resulting from a local and rapid increase in Ca ²⁺ concentration that rises to a peak within ~10 ms and decays within ~50 ms (see eFig. 9-1 B ). Such spontaneous sparks in cardiac myocytes are due to the small opening probability of SR RYR Ca ²⁺ -release channels that depends upon [Ca ²⁺ ] in the cytoplasm and SR lumen. Biophysical analysis indicates that a single spark event corresponds to the simultaneous opening of a cluster of Ca ²⁺ -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 Ca ²⁺ sparks generally requires low activation conditions of Ca ²⁺ release, they can be observed in single cardiac myocytes activated by a depolarizing voltage pulse at the leading edge of a transient rise of [Ca ²⁺ ] _i (see eFig. 9-1 A ).

eFigure 9-1 C shows a series of Ca ²⁺ 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 Ca ²⁺ 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 [Ca ²⁺ ] in muscle cells is the result of the stochastic summation of many individual spark events corresponding to localized bursts of intracellular Ca ²⁺ release. Studies of Ca ²⁺ sparks have also confirmed that voltage-activated mechanical coupling underlies EC mechanisms in skeletal muscle, whereas Ca ²⁺ -induced Ca ²⁺ release underlies these mechanisms in cardiac and smooth muscle.

Due to the tight voltage control of Ca ²⁺ release and termination by brief ~2-ms Na ⁺ action potentials in mammalian skeletal muscle, classical Ca ²⁺ 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 Ca ²⁺ release, which presumably facilitates precise control of many body movements.

Although we have stressed that EC coupling in skeletal muscle primarily involves direct mechanical coupling between the L-type Ca ²⁺ channel in the T-tubule membrane and the Ca ²⁺ -release channel of the SR, N9-1 other mechanisms modulate the activity of RYR1. For example, RYR1 is subject to regulation by cytoplasmic Ca ²⁺ , Mg ²⁺ , ATP, and calmodulin (CaM) as well as protein kinases such as protein kinase A (PKA; see p. 57 ) and Ca ²⁺ -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 Ca ²⁺ , and thus stronger skeletal muscle contraction ( Box 9-1 ).

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. 9-4 A, 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.

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. 9-4 B ), 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. 9-4 B ). 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. 9-4 C ). 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. 9-5 A ). N9-4 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.

N9-4

F-Actin

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. 9-5 A ).

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. 9-5 A ). 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 t ropomyosin

• 2

  Troponin C (TnC or TNNC), which binds Ca ²⁺ . Troponin C is closely related to another Ca ²⁺ -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 [Ca ²⁺ ] _i .

Thick Filaments

Like actin thin filaments, thick filaments are also an intertwined complex of proteins (see Fig. 9-5 B ). 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), N9-5 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
Ca 2+ release mechanisms	RYR1, Ca 2+ -release channel or ryanodine receptor (RYR1)	RYR1 (RYR1)	RYR1 (RYR1)	RYR2 (RYR2)	IP 3 R1, IP 3 R2, IP 3 R3 (ITPR1, ITPR2, ITPR3) RYR3 (RYR3)
Ca 2+ sensor	Troponin C 1 (TNNC1)	Troponin C 2 (TNNC2)	Troponin C 2 (TNNC2)	Troponin C 1 (TNNC1)	CaM (multiple isoforms)

* Gene names in parentheses.

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

N9-5

Muscle Myosin

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 ).

A myosin heavy chain molecule has ~2000 amino acids in three regions: N9-5 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 Ca ²⁺ triggers the interaction between a thin filament and a myosin head group from a thick filament.

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. 9-4 B ). 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 (P _i ) 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. N9-6 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

  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. 9-2 A ). 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. 9-2 B ). 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. 9-2 C ). 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 ).

  

• 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 P _i from the myosin. Dissociation of P _i 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).

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 [Ca ²⁺ ] _i —as we will see in the next section.