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1  INTRODUCTION

Cardiac contraction is driven by the cyclic interaction of the motor protein type II myosin with the actin filament, consuming ATP as the energy source to produce tension or shortening. Characterization of the functions of myosin and its subunits on the molecular level provides important steps in the understanding of the physiological and pathophysiological states of the heart.

1.1 Cardiac Excitation-Contraction-Coupling

The release of Ca2+from the sarcoplasmic reticulum (SR) following membrane depolarization and subsequent Ca2+-induced myofilament activation is referred to as excitation–contraction coupling (ECC) (Fig. 1). Myocardial contraction is activated by a transient rise in cytosolic free calcium concentration [Ca2+] to about 1 μM from a resting diastolic [Ca2+]of about 0.1 μM. Despite a rather modest rise in the average free [Ca2+], activation of the contractile proteins typically requires the binding of between 40 and 60 μM total Ca2+. The bulk of this Ca2+required for activation of contraction originates from the sarcoplasmic reticulum (SR), with Ca2+influx through L-type Ca2+channels and Na+–Ca2+exchangers (NCX) making more minor contributions, although the relative contribution varies between species (Bers 2002).

A central feature of ECC is the gating of SR Ca2+release channels (called ryanodine or RyRs receptors), where gating refers to the opening (or activation) and the closing of individual RyRs channels located within the terminal cisternae of the SR. Gating of RyRs channels is controlled primarily by elevations of [Ca2+]in the subsarcolemmal space between the T-tubular membrane of the sarcolemma and the terminal cisternae of the SR, which occurs following the opening of sarcolemmal L-type Ca2+ channels in response to membrane depolarization (Lopez-Lopez et al. 1994; Lopez-Lopez et al. 1995). The amplification process whereby elevated subsarcolemmal Ca2+ ‘triggers’ further Ca2+release from the SR is called Ca2+-induced Ca2+release (CICR).

Relaxation of myocyte contraction involves the extrusion of Ca2+from the cytosol. At steady state, the majority of relaxation involves Ca2+reuptake into the SR by the Ca2+-[page 11↓]ATPase (SERCA2a), with the remaining Ca2+(equivalent to the amount entering via L-type Ca2+channels) being extruded from the myocyte via the NCX (Bers 2002).

Figure 1 . Scheme of cardiac excitation-contraction-coupling events in a ventricular myocyte.

Electrical excitation at the sarcolemmal membrane activates voltage-gated Ca2+channels, and the resulting Ca2+entry activates Ca2+ release from the sarcoplasmic reticulum (SR) via ryanodine receptors (RyRs), resulting in contractile element activation. NCX, Na+/Ca2+ exchange; ATP, ATPase; PLB, phospholamban; SR, sarcoplasmic reticulum. Inset shows the time course of an action potential, Ca2+ transient and contraction. (Bers 2002).


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1.2  Cardiac myofilaments

Cardiomyocytes contains large number of myofilaments that are organized in a regular array of cross-striations. The cross striations of the myocardium reflect the organization of the contractile proteins into thick and thin filaments. The thick filaments, which are composed largely of myosin, extend the length of the A-band to which they contribute both its darkly staining characteristics and its high birefringence. The thin filaments are composed of actin and the regulatory proteins tropomyosin and troponin complex, they extend the length of the I-bands to which they contribute both its lightly staining striations and less birefringent. A broad dense M-band is found in the center of each A-band, while the I-bands are bisected by Z-lines. The latter delimit the fundamental morphologic unit of striated muscle, the sarcomere, which is defined as the region between two Z-lines. Each sarcomere thus consists of a central A-band plus two adjacent half I-bands. (Fig. 2) (Katz 2001).

During systole the thin filaments are drawn toward the center of the sarcomere by movements of the myosin cross-bridges, which project from the thick filaments to establish bonds with the thin filaments. The thin filaments extend from the Z-lines at either end of the sarcomere into the A-bands, where the extent of overlap between the thick and thin filaments depends on sarcomere length (Katz 2001).


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Figure 2 . Ultra-structure of the working myocardial cells.

The A-band, the I-band, the Z-lines, the sarcoplasmic reticulum, the transevers tubular system (t-tubule) and the mitochondria are shown in the central sarcomere (see text for explanation) (Katz 2001).


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1.2.1  Sarcomeric proteins associated with cardiac myofilaments

Although these proteins do not participate directly in the contractile process, they maintain sarcomere structure and provide mechanical linkages that convey the tension developed by the contractile proteins to adjacent sarcomeres and ultimately to the ends of the muscle (Schiaffino et al. 1996)

At least five proteins are present in the thick filaments contain amino acid sequences that resemble both fibronectin and immunoglobulins (Helmes et al. 1996; Linke et al. 1997; Small et al. 1992). Titin which is also called connectin, is one of the largest known proteins with a molecular weight of about 3,000,000, extends into the thick filaments from the Z-lines (Fig. 3). Titin includes both an elastic region, which is located in the I-band, and a rigid region that supports the thick filaments within the A-bands.

Myosin binding protein C, also called C protein (MW-140,000), which links the thick filaments and titin in the central third of the A-band, plays a role in sarcomere formation (Fig. 3) and contains a number of phosphorylation sites that regulate contractility (Winegrad 1999). Myosin binding protein H (MW-74,000) is present in the purkinje fibers but not in working cardiac myocytes (Alyonycheva et al. 1997). M-protein and myomesin (MW-185,000) contribute to a set of transverse striations, called M-band, which link the thick filaments in the center of the A-band (Fig. 3). These proteins participate in myofibrillogenesis and stabilize interaction between titin and the thick filaments. Nebulette (MW-107,000) is oriented along the axis of the thin filament and projects from the Z-lines into the I-bands (Moncman et al. 1995). Tropomodulin (MW-43,000) is a capping protein, and is found at the ends of the of the thin filaments where, by covering the end of the F-actin polymer (Fig. 3), it helps to determine thin filament length (Sussman et al. 1994). α –Actinin (MW-140,000) and Cap Z (β-actinin) (MW-38,000), they weave the ends of thin filaments into the Z-lines at the ends of each sarcomere (Fig. 3).


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Figure 3 . Major components of the cardiac sarcomere:

Actin (green), myosin (red). Rod-like tropomyosin molecules (black lines). Thin filaments in muscle sarcomeres are anchored at the Z-disk by the cross-linking protein α -actinin (gold) and are capped by CapZ (pink squares). The thin-filament pointed ends terminate within the A band, are capped by tropomodulin (bright red). Myosin-binding-protein C (MyBP-C; yellow transverse lines), (Gregorio et al. 2000).


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1.2.2  Thin Filament

The main site for Ca+2regulation is the thin filament. Figure 4A is a diagram of the thin filament in striated muscle showing the three components: actin, tropomyosin (Tm), and troponin (Tn) with the three Tn subunits (Gordon et al. 2000).

Monomeric globular actin (G-actin) polymerizes spontaneously to form the backbone of the thin filament, F-actin. The double helical structure of F-acin as two-stranded long-pitch helical structure is evident form electron microscopy of isolated filaments and X-ray diffraction patterns form actin bundles and intact muscle. There are grooves between the two so-called long pitch strands; tropomyosin molecules lie along these grooves such that each tropomyosin molecule interacts with seven actin monomers, There is one troponin complex associated with each tropomyosin molecule (Fig. 4A) (Gordon et al. 2000).

G-actin comprises a single polypeptide chain (mol. Wt 42 000), which binds Ca+2 and ATP reversibly. The structure of the actin monomer (Fig. 4B) was determined at atomic resolution (Holmes et al. 1990; Kabsch et al. 1990) into the actin filament helix by modeling against fiber X-ray diffraction patterns from aligned gels of actin filaments (Popp et al. 1987; Popp et al. 1991). The resulting structure is shown in Figure 4B. Each actin monomer comprises four subdomains. Structure positions of subdomains 3 and 4 close to the helix axis, where they interact with subdomains 3 and 4 of other actin monomers. Subdomains 1 and 2 are on the outside of the helix. Subdomain 1 contains the amino and carboxyl termini of the molecule (Holmes et al. 1990).

Tropomyosin (Tm) forms an α helical coiled-coil dimmer. Each tropomyosin molecule extends across seven actin monomers in the thin filament and overlaps with neighbouring tropomyosin by about eight residues to form a continuous, rather flexible structure. Image reconstruction studies indicate that tropomyosin lies in the groove of the actin bound to the inner (large) domain of actin (Fig. 4A). One troponin is bound to each tropomyosin dimer, the bulk of the troponin lying about 20nm from the tropomyosin C-terminus (Fig. 4A) (Milligan et al. 1990).


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Figure 4 . Thin filaments:

(A) A model of the molecular arrangement of troponin (Tn), tropomyosin (Tm), and actin in the cardiac muscle thin filament. The various troponin subunits are indicated [TnC (red), TnT (yellow), and TnI (green)] as they lie along the two-stranded tropomyosin shown (brown) and (orange) that in turn lies along an actin (gray) (Cohen 1975). (B) Ribbon diagram (i.e.polypeptide backbone only) of the actin monomer structure subdomains: 1 (red), 2 (green), 3 (blue), and 4 (yellow) (Kabsch et al. 1990).


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Troponin comprises three subunits, designated, TnI, TnC, TnT, on the basis of their function. Structural studies showed that Tn belongs to the calcium binding protein family (Kretsinger 1980).

TnC is the myofibrillar protein that by binding Ca2+, transmits the signal to the thin filament. The structure of TnC has been solved to atomic resolution by X-ray crystallography; it consists of two globular domains corresponding to the amino and carboxyl termini, and these are linked by a central, nine-turn α-helix. Each globular domain contains two divalent metal binding sites formed from the helix-loop-helix EF hand motif. The two metal binding sites in the amino-terminal domain are Ca2+specific and are believed to be the regulatory sites (i.e., Ca2+ binding to these sites is believed to trigger activation). TnC interacts both with TnI and TnT. TnC interacts more strongly with TnI when Ca2+is not bound to the TnC regulatory sites; modulation of this interaction is believed to be the regulatory mechanism within the troponin complex (Herzberg et al. 1985).

Troponin and tropomyosin exert their regulatory effect in the thin filament by causing inhibition when TnC has no Ca2+bound to the regulatory sites. The actin filament without its regulatory proteins is intrinsically active; that is, it activates myosin ATPase and supports force production and movement. TnI has a major role in inhibiting this activity. In fact, on its own, TnI has an effect on actin filament activity similar to the whole troponin complex at low [Ca2+]. Besides its interaction with TnC, TnI appears

to interact with TnT and actin. The interaction with actin is particularly important: it is thought that through this link TnI exerts its regulatory effect and this interaction is modulated by Ca2+ binding to TnC (Squire et al. 1998).

TnT is involved in the attachment of the troponin complex to tropomyosin and thus to actin. It is a rodlike molecule with amino and carboxyl termini at opposite ends. The amino-terminal region binds to the head–tail junction of tropomyosin; the carboxyl-terminal region is closely associated with TnC and TnI (Squire et al. 1998).


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1.2.2.1  Thin filament regulation in myocardial contraction

Contraction occurs when myosin S1 heads of the thick filament attach to and exert force on actin molecules in the thin filament (Geeves et al. 1999; Xu et al. 1999). This force causes the thin filament to slide over the thick filament and the sarcomere to shorten and develop force against a load. Ca2+binding to troponin (Tn) on the thin filament initiates the force-generating interaction of myosin and actin, and ATP hydrolysis provides the energy for the molecular changes that drive force generation and muscle shortening. Regulation by Ca2+is mediated through changes in the thin filament, although modulation can occur through myosin (Gordon et al. 2001).

Figure 5A shows the salient structural features of thin filament regulation. Helically arranged actins form the backbone of the thin filament, with the regulatory proteins tropomyosin (Tm) and troponin Tn attached to actin in a 7:1:1 (A:Tm:Tn) ratio. Tm, a long, flexible molecule, binds to seven actin monomers in the thin filament helix and overlaps the adjacent Tm. Tn attaches to two actins in the absence of Ca2+ through its TnI subunit and to Tm through the TnT subunit.

Ca2+binding to the TnC subunit strengthens the TnC-TnI interaction and detaches TnI from its contacts with actin. Recent structural studies (Vibert et al. 1997) show that this Ca2+-mediated detachment of TnI from actin allows Tm to move over the surface of the thin filament. On the actin surface, there are sites for weak (mainly electrostatic) and strong myosin binding (Fig. 5A). Tm either rolls around its axis or slides from a position near the outer edge of the thin filament [where it covers many of the myosin binding sites on actin (Fig. 5B)] to a position allowing increased weak and some strong myosin head binding (Fig. 5C). Tm is a flexible molecule, and its positioning should be considered dynamic; Tm does not occupy a single fixed position in the presence of elevated [Ca2+] but “rocks and rolls” or “slips and slides” back and forth over the actin surface. The Tm positions (Fig. 5, B–D) should therefore be considered average positions. When myosin cross-bridge is strongly bound to actin, Tm is locally stabilized in a position that makes both weak and strong myosin binding sites available on nearby actin monomers (Fig. 5D).


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Thus structural data suggests that thin filament activation is achieved by the movement of Tm over the actin surface, which is controlled both by Ca2+binding to TnC and initial cross-bridge binding to actin to allow additional strong cross-bridge binding. This Tm motion permits force generation and shortening (Gordon et al. 2001).


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Figure 5 . Regulation of thin filament in contraction

Actin residues are light gray, except for residues (1–4, 24–25, 95–100; shown in light blue) making weak electrostatic interaction with myosin and residues (144–148, 340–346, 332–334; shown in red) forming stronger attachments with myosin. In B–D, a surface rendering of a cardiac tropomyosin segment (residues 61–112 of each tropomyosin strand) in dark gray. Arg90 of tropomyosinm in each strand is shown in yellow to illustrate the putative rolling motion of tropomyosin. (Gordon et al. 2001).


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1.2.3  Myosin filament

The motor protein, myosin II, is composed of two heavy chains with molecular masses of 200 kDa each and four light chains, two each of so called essential and regulatory light chains (MLC 1 and MLC 2,respectively), of molecular mass 20 kDa. The heavy chains form a parallel two-chain coiled-coil structure over most of their length except for the large, globular NH2-terminal regions, termed heads or S1 (subfragment 1). One pair of light chains bind to each S1. The coiled-coil region of the myosin, termed the myosin rod (tail). The S1 head structure projecting from the backbone of the thick filament interacts with actin to generate force and filament sliding (Fig. 6) (Sellers et al. 1995).

Figure 6 . Schematic representation for myosin molecule structure:

The myosin heavy chains (MHC) have a globular "head" with the ATP and actin binding sites at the amino terminal and a long α-helical "tail" at the carboxy terminal. The essential myosin light chain (MLC-1) and the regulatory myosin light chain (MLC-2) at the neck region.


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Limited proteolysis of S1 reveals three major segments, a 25-KDa N-terminal, a central 50-KDa and a 20-KDa C-terminal domain (Mornet et al. 1979). The three dimensional structure of the S1 domain of the myosin molecule has been elucidated by X-ray crystallographic analysis (Fig. 7) (Rayment et al. 1993). It consists of a heavy chain (MHC) which folds at the N-terminus into an asymmetric globular head domain. This head is 16.5 nM long, 6.5 nM wide, and 4 nM thick and comprises a seven-stranded β-sheet connected by flanking α-helices and /or loops which constiute the catalytic domain.

The 50-KDa domain is split by a long narrow cleft with actin-binding sites located on both sides of the cleft. Also the ATP-binding (active) site forms an open cleft and is located opposite to the actin-binding site at the 25/50 KDa junction. The apex of the long cleft through the 50 KDa domain is very close to the nucleotide binding cleft, thus mediating signal transduction between the actin and nucleotide binding clefts.

Amino acids 771-843 at the C-terminus form an 8.5 nM α- helical structure, the neck region that binds two types of light chains. The essential myosin light chain binds between amino acid 783 and 806 and the regulatory light chains further downstream between amino acids 808 and 842. A so-called converter domain joins the catalytic and light chain binding domains (around amino acids 711-771).


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Figure 7 . Three-dimensional ribbon structure of Myosin S1

(Rayment et al. 1993). The MHC consists of a 25-K N-terminal (green), a 50-K (red), and a C-terminal joining 20-K (blue) domain. The pear-shaped head is called the catalytic domain. Essential MLC-1 (yellow) and regulatory myosin light chain MLC-2 (magenta) are associated with the α-helical C-terminal part of the 20-K domain (light chain binding domain)


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1.2.3.1  Myosin cross-bridge

The swinging-cross-bridge hypothesis of muscle contraction (featured in most text books) was recently modified into swinging-lever arm hypothesis in which the bulk of the cross-bridge is envisaged to bind to actin without rolling on the surface during the power stroke (as had been initially suggested); large movements are envisaged as coming from the distal (C-terminal) part of the myosin cross-bridge moving as a lever arm (Geeves et al. 1999).

The structural changes in the cross-bridges are associated with the strain of an elastic component present in the cross-bridges, which operates over a range of 8–10 nm (Huxley et al. 1971). The α-helical neck domain is believed to function as a lever arm which swings relative to the catalytic domain: picometer changes in the active site of S1 are magnified into nanometers of motion by rotation of the converter domain (Eisenberg et al. 1985; Holmes 1997). In fact, truncation and elongation of the light chain binding domains show that the sliding velocity of actin filaments is proportional to the length of the lever arm (Uyeda et al. 1996). Furthermore, the light chain binding domain, rather than the catalytic domain, reveals tilting motions during length perturbations of an isometrically contracting muscle (Hopkins et al. 1998; Irving et al. 1995; Lombardi et al. 1995). The elastic element of the cross-bridge may therefore reside in the lever arm (Howard et al. 1996).

1.2.3.2 Cross-bridge cycle kinetics

In the intact contractile structure the ATP- or ADP-Pi- loaded MHC binds (as the cross-bridge) to the N-terminus of actin. Myosin undergoes changes in actin affinity and structure, being strongly attached to actin (having high affinity) or weakly attached (having low affinity) (Brenner 1988; Eisenberg et al. 1985). Force is generated upon the transition from the weakly to the strongly attached state. This transition is considered to be coupled to the Pi release step (Dantzig et al. 1992). Figure 8 shows the structural and biochemical events in a cross bridge cycle (Gordon et al. 2001):


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Step 1: At physiological ATP concentrations (3–5 mM), ATP binding to myosin is very rapid and irreversible.

Step 2: The subsequent detachment of actin from the actin-myosin·ATP (A~M·ATP) complex is similarly rapid and is caused by an opening between myosin’s upper and lower 50-kDa regions like the opening of jaws.

Step 3: A “flexing” or bending of the myosin neck region accompanies the hydrolytic cleavage of ATP, whose equilibrium constant (K3, defined as k+3/k–3) is only ~10.

Step 4: Following ATP cleavage, myosin again binds weakly to actin at a high rate, but in the absence of Ca2+Tm sterically blocks access of the myosin head to strong binding sites on actin (Fig. 8B). However, when Ca2+is bound to TnC, TnI detaches from actin, allowing the Tm/Tn complex to roll or slide over the thin filament surface. This exposes weak binding sites on actin and transiently exposes strong binding sites on actin (Fig. 8B) for binding to the complementary regions in myosin’s 50-kDa domain.

Step 5:The greater the [Ca2+], the greater the fraction of time the Tm/Tn complex allows myosin access to strong binding sites on actin. Consequently, the rate of strong cross-bridge attachment, the flux through is dependent on [Ca2+] and Tm position (i.e., in the simplest case, the value of k+5 is proportional to the fraction of Tn having bound calcium). Strong binding of myosin to actin (Fig. 8B) is associated with movement of the upper and lower 50-kDa subdomains toward each other (or closing the jaws). This movement may allow the neck region of myosin to extend, opening a pathway for inorganic phosphate release from the ATP binding pocket in myosin. Alternatively, closing the jaws might promote inorganic phosphate release from the binding pocket, which then allows the extension of myosin’s neck region.

Step 6: In any event, myosin neck extension is the power stroke that, in isometric muscle, stretches an elastic element by some 10 nm and produces force. In nonisometric conditions, shortening of the neck extension causes the thick and thin filaments to slide past each other.


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Step 7: Is an irreversible isomerization and is strain sensitive; i.e., when the force on the cross-bridge is large as in isometric contractions, k+7 is slow (3–10 s–1) and is the rate-limiting step for the cross-bridge cycle. Finally, ADP is released from A·Mf·ADP (where f is a cross-bridge exerting force).

Step 8: Is reversible to form the rigor state, A·Mf. During isometric contractions, the slowness of k+7 causes the population of cross-bridges in the initial force-bearing (A·Mf*·ADP) state to rise and with it force.

Cross-bridges attach and exert force constantly during steps 7, 8, and 1 during isometric contraction, and force drops to zero when the cross-bridges detach in step 2. During shortening contractions the filaments slide past each other, the strain on the cross-bridge is reduced, and step7 occurs more rapidly. This accounts for the Fenn effect (an increased rate of energy liberation above the isometric rate as shortening velocity increases). The chemomechanical mechanism shown in figure 8 implies that during an isometric contraction, a cross-bridge remains strongly attached to actin for a relatively long time (>100 ms/cycle). Strongly bound cross-bridges prevent Tm/Tn from returning to its blocked or closed position, maintaining the thin filament in a “switched on” position (Fig. 8B). In the absence of Ca2+, cross-bridge detachments at the end of the cycle allows Tm/Tn to cover the strong myosin binding sites on actin and deactivate the thin filament (Fig. 8A)


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Figure 8 . Cross-Bridge cycle kinetics.

(A) shows the cross-bridge cycle in terms of biochemical changesand the corresponding structural changes are shown in (B) (Gordon et al. 2001)


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1.2.3.3  Myosin heavy chains (MHC)

In the human heart two sarcomeric myosin II genes are expressed generating two isoenzymes of the myosin heavy chain (designated α and β-MHC). These are located in tandem on the long arm of human chromosome 14q11.2-q13 (Saez et al. 1987). In the normal heart both MHC isoenzymes are expressed in a tissue-specific manner: the α -MHC is preferentially expressed in the atrium, and the β -MHC is almost exclusively expressed in the ventricle (Bouvagnet et al. 1984).

The human α –MHC contains 1939 amino acids residues (Matsuoka et al. 1991) while the β –MHC contains 1935 amino acids (Jaenicke et al. 1990). A total of 131 residues differ between both heavy chains and most of these differences are confined to regions of biological significance in the S1 subfragment such as the N-terminus, the ATP binding pocket, the actin binding cleft, the light chain binding domain and in the two hinge regions further down in the rod (tail) domain (Schaub et al. 1998).

In the hypertrophied atrium considerable amounts of β-MHC are expressed (Gorza et al. 1984; Mercadier et al. 1983), which is associated with a decrease in maximal shortening velocity (Arndt et al. 1989). α-MHC, however, is either found to be a minor component of human ventricular myosin (Gorza et al. 1984; Hirzel et al. 1985; Mercadier et al. 1983) or could not be detected at all (Ritter et al. 1999). Furthermore, there are no changes in α-MHC expression during hypertrophy of the human ventricle (Gorza et al. 1984; Hirzel et al. 1985; Mercadier et al. 1983; Ritter et al. 1999).

1.2.3.4 Myosin light chains (MLC)

Two types of myosin light chains (MLC), essential and regulatory, are associated with the neck region of the MHC. The essential MLC is designated as the MLC-1 or alkali MLC. The regulatory MLC is designated as MLC-2 or phosphorylatable MLC or 5,5-dithio-bis-(2-nitrobenzoate)-MLC (Ritter et al. 1999). Together with calmodulin and troponin C, both types belong to the superfamily of EF hand Ca2+binding proteins (Moncrief et al. 1990).


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In the human heart two different genes encode the essential myosin light chain isoforms, the ventricular-specific (VLC-1) and an atrial specific (ALC-1). VLC-1 is located on chromosome 3p21 (Fodor et al. 1989), coding for a 194 amino acid protein, which is the same isoform as the MLC-1 present in the adult slow skeletal muscle. ALC-1 is located on chromosome 17q21 (Seharaseyon et al. 1990) coding a 196 amino acid protein. Several MLC-2 isoforms exist in the human, preferentially expressed in the atrium (ALC-2) or in the ventricle (VLC-2). VLC-2 has been mapped to chromosome12q23-q24 (Macera et al. 1992) in human encoding a 166 amino acid protein. ALC-2 is composed of 175 amino acids (Hailstones et al. 1992), but no data are available yet about the chromosomal localization. Electrophoretic analysis suggests the existence of two different VLC-2 isoforms in the human heart having the same molecular weight but different isoelectric points (VLC-2a for the more acidic and VLC-2b for the more basic isoform) (Price et al. 1980).

In the normal human heart mainly the VLC-2b isoform is expressed (LC-2b to LC-2a ratio of 2.3) (Morano et al. 1997; Morano et al. 1996). In most patients with limited cardiac functions, this ratio remains at its normal level. However, recently it was found that in patients with HOCM VLC-2a expression declines in favor of the VLC-2b form (Ritter et al. 1999).

Expression of ALC-1 is tissue specific and developmentally regulated. Human embryos express large amounts of ALC-1 both in the whole heart and in skeletal muscle (Barton et al. 1985). ALC-1 protein levels decrease in the ventricle to undetectable levels during early postnatal development but persisted in the atrium throughout the whole life (Fallot 1963).

The situation differs in patients with congenital heart disease such as Tetralogy of Fallot. Tetralogy of Fallot is a complex congenital heart disease characterized by four components: right ventricular infundibular stenosis, ventricular septal defect, dextroposed aorta overriding the interventricular septal defect, and right ventricular hypertrophy (Auckland et al. 1986). The hypertrophied right ventricle of children with tetralogy of Fallot express large amounts of VLC-1 in the atrium (Shi et al. 1991) and ALC-1 in the ventricle, up to adulthood (Auckland et al. 1986). Similarly, the hypertrophied left ventricle of patients with ischemic, dilative, and hypertrophic [page 31↓]cardiomyopathy express ALC-1 (Morano et al. 1997; Ritter et al. 1999; Schaub et al. 1998). Surgical intervention and subsequent normalization of the hemodynamic state decrease ALC-1 expression in these patients (Sutsch et al. 1992).

MLC-1 binds not only to the neck domain of the MHC (Rayment et al. 1993) but also with its N-terminus to the C-terminus domain of actin (Hayashibara et al. 1994; Morano et al. 1997; Sutsch et al. 1992; Trayer et al. 1987). Thus MLC-1 tethers the MHC to the actin filament. It has been demonstrated that MHC and MLC-1 bind to different actin monomers (Timson et al. 1998). Binding of MLC-1 to actin cannot be predicted from the crystal structure of myosin S1 (Rayment et al. 1993). This is due to the limited resolution of the three-dimensional structure of the N-terminus of MLC-1 (Rayment et al. 1993). In fact, around 40 N-terminal amino acid residues of MLC-1 are not seen in the crystal structure (Rayment et al. 1993). This missing part of MLC-1 contains ten Pro and ten Ala residues, which could form an antenna-like structure long enough to bridge the gap to the actin filament (Morano 1999).

Experimental evidence for the functional importance of the MHC/actin tether has been obtained by weakening the tether on the MLC-1/actin interface and/or MLC-1/MHC interface and simultaneous registration of cross-bridge function. Inhibition of the MLC-1/actin interaction by peptide competition using synthetic N-terminal MLC-1 peptides increases force production and shortening velocity of both demembrated (skinned) and of intact electrically driven human ventricular fibers (Morano et al. 1995) as well as myofibrillar ATPase activity (Rarick et al. 1996).

Assuming that tethering MHC to the actin filament via MLC-1 imposes a load on the myosin cross-bridge, thus relieving or weakening the MHC/actin tether decreases this load and accelerates cross-bridge cycling kinetics and at the same time enhances tension output per cross-bridge, thus increasing contractility (Morano et al. 1995).

The ALC-1 and VLC-1 differ in the primary structure of the N-terminus (Fodor et al. 1989). Indeed, the affinity for actin of the N-terminal peptide 5–14 derived from ALC-1 is significantly lower than the actin affinity of the corresponding N-terminal peptide of VLC-1 (Morano et al. 1997). These results support the idea that binding of ALC-1 to actin is weaker than the binding of VLC-1 to actin, representing a weaker MHC/actin tether [page 32↓](Morano et al. 1996): because of its low actin affinity ALC-1 is a weaker MHC/actin tether than VLC-1 and has increased cross-bridge cycling kinetics and force generation.

1.2.3.5 Regulation of ALC-1 expression in the heart

Regulation of ALC-1 expression in the human heart is still not well understood. In the mouse two E boxes, which interact with muscle-specific basic helix-loop-helix (bHLH) regulatory proteins of the MyoD family, and a diverged CArG box, which binds to the serum response factor, exist within the first 630 bp of the ALC-1 promoter region (Catala et al. 1995).

E boxes have been shown to be sufficient for ALC-1 transcription regulation during skeletal muscle differentiation (Catala et al. 1995). bHLH regulatory factors of the MyoD family regulate skeletal muscle differentiation by forming heterodimers with E12 bHLH factors that bind to E-box elements, thus increasing the transcription rate of target genes (Olson 1992). Recently two cardiac-specific bHLH proteins homologous to the MyoD family have been detected in the heart, designated as the E and D forms of heart–autonomic nervous system–neural crest derivative (HAND) (Cserjesi et al. 1995; Srivastava et al. 1995). They are important for early development and looping of the embryonic heart (Srivastava et al. 1997). Since both HAND transcription factors bind weakly to E-box as E12 heterodimers, the ALC-1 may be a target gene. Recently it has been reported that in the human hypertrophied ventricle there is upregulation of HAND gene expression and a positive correlation between HAND and ALC-1 mRNA (Ritter et al. 1999). Furthermore, ALC-1 expression may be regulated by endogenous antisense ALC-1 mRNA present in the human ventricle (Ritter et al. 1999). Moreover Ca2+-calmodulin-dependent processes, which are involved in the development of human heart hypertrophy, increased the activity of the human ALC-1 promoter. Both activation of calcineurin (CaN) and Calmodulin dependant kinase IV (CaMKIV) revealed a potent role in human ALC-1 gene regulation (Woischwill et al. 2004).


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1.2.3.6  Functions of ALC-1 in the heart

Skinned fiber studies showed that the shortening velocity, rate of tension development, isometric force generation, and Ca2+sensitivity of isometric force generation increase upon partial replacement of VLC-1 by ALC-1 in the human ventricles (Morano et al. 1997; Morano et al. 1996). Furthermore, there was a significant positive correlation between ALC-1 expression and dP/dtmax of patients with hypertrophic obstructive cardiomyopathy (HOCM) in vivo (Ritter et al. 1999). Thus crossbridge cycling kinetics and tension generation per cross-bridge are modulated by differential expression of MLC-1 genes. These results demonstrated for the first time that there is a molecular mechanism which allows the ventricular cardiomyocyte to adjust to enhanced work load through modification of the structure of the molecular motor – the partial substitution of VLC-1 by ALC-1 which increases power output of the sarcomeric motor macromolecules and improves cardiac contractility (Morano 1999). These results have been supported in a transgenic mouse model overexpressing the mouse ALC-1. By the examination of the chemically skinned fibre function, the ventricular fibres from the transgenic mice had a higher Vmax of shortening when compared to non-transgenic mice. Furthermore this light chain Isoform switch in the transgenic mouse model lead to increased contractile functions at the whole heart level (Fewell et al. 1998).


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1.3  Aim of the study

The functional properties of the human atrial essential myosin light chain (hALC-1) has never been evaluated in an intact heart preparation on the whole organ level. Functional examination of the chemically skinned ventricular fibers from a transgenic mouse model overexpressing the mouse ALC-1 showed a higher maximal shortening velocity (Vmax) when compared to non-transgenic mice: In the mouse heart, 95% replacement of mouse VLC-1 by the mouse ALC-1 was accompanied by a 1.78-fold increase in Vmax of shortening. However chemically skinned ventricular fibers of patients expressing the hALC-1 with 20% replacement of human VLC-1 by human ALC-1 showed 1.88-fold increase in maximal shortening velocity (Vmax). Although the data are consistent, it appears that the effect of mouse ALC-1 is attenuated as compared with hALC-1. These data suggest that there is a significant difference between the human and mouse ALC-1 isoforms at the functional level. To verify the functional efficiency of hALC-1 compared with mouse ALC-1 and to analyze hALC-1 functions in intact whole heart preparations, a transgenic rat-model overexpressing the hALC-1 was produced and characterized at the protein and functional levels.


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