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Cardiovascular Research 1999 44(2):266-273; doi:10.1016/S0008-6363(99)00219-9
© 1999 by European Society of Cardiology
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Copyright © 1999, European Society of Cardiology

Actin—myosin interaction

Seiryo Sugiura*

Department of Cardiovascular Medicine, Graduate School of Medicine, University of Tokyo, Hongo 7-3-1, Bunkyo-ku, Tokyo 113-8655, Japan

* Tel.: +81-3-3815-5411; fax: +81-3-3814-0021 sugiura-2im{at}h.u-tokyo.ac.jp

Received 22 March 1999; accepted 25 June 1999


   Abstract
Top
 Abstract
1 Introduction
 2 Basic characterization of...
 3 Force--velocity relation in...
 4 Findings on cardiac...
 5 Clinical implications
 6 Future perspective
 References
 
Recent advances in the study of muscle physiology was made possible by the application of novel experimental techniques including in vitro motility assay, molecular biology, and X-ray crystallography. A similar approach was successfully applied in studying the properties of cardiac actin—myosin interaction. Implication in clinical cardiology is also reviewed.

KEYWORDS Cardiomyopathy; Contractile apparatus; Contractile function; Heart failure


   1 Introduction
Top
 Abstract
 1 Introduction
2 Basic characterization of...
 3 Force--velocity relation in...
 4 Findings on cardiac...
 5 Clinical implications
 6 Future perspective
 References
 
It is well established that the contraction of striated muscle results from the relative sliding of thick (composed mainly of myosin) and thin (composed mainly of actin) filaments. Since this discovery in the early 1950s [1], numerous attempts have been made to clarify the molecular mechanism driving the filament sliding. Because the fundamental function of muscle is the chemo-mechanical conversion utilizing the chemical energy derived from the splitting of the high energy bond in ATP molecules, biochemical studies which inevitably uncouple the chemical and mechanical events have not been so successful in solving this problem. Alternatively, this issue has been pursued by using intact or demembranated (skinned) muscle preparations and interpreting the obtained results in the light of a specific model. Although this type of approach has made significant progress in muscle physiology, the complex structure of the muscle cell and the large number of actin and myosin molecules involved have always been obstacles to dissect the molecular event responsible for muscle contraction.

In recent years, novel experimental systems called in vitro motility assays have been developed [2–4]. In these systems, we can observe the mechanical event taking place between a limited number (currently, even a single motor assay is possible [5,6]) of actin and myosin molecules and evaluate it qualitatively under controlled conditions. The in vitro motility assay combined with X-ray crystallography and molecular biology techniques has provided new experimental findings leading to new concept in the molecular basis of muscle contraction.

The impact was not limited to the fundamental physiology of muscle contraction. Recently, various sarcomeric proteins including myosin and actin have been reported to be responsible for the pathogenesis of various diseases affecting the cardiac muscle [7–13]. Furthermore, there is evidence that cardiac contraction and relaxation are regulated at the crossbridge level [14,15]. Accordingly, the application of these techniques in cardiology will no doubt contribute to our understanding of these clinical problems.

In this brief review, I will discuss the recent advances in muscle physiology made using these novel experimental techniques, especially emphasis will be put on the in vitro motility assays. Then application of these techniques to the study of cardiac actin—myosin interaction will be presented. Finally, clinical implication will also be discussed.


   2 Basic characterization of actin—myosin interaction
Top
 Abstract
 1 Introduction
 2 Basic characterization of...
3 Force--velocity relation in...
 4 Findings on cardiac...
 5 Clinical implications
 6 Future perspective
 References
 
2.1 In vitro studies
Since actin—myosin interaction takes place in an aqueous environment, we must utilize an optical microscope to observe this phenomenon (recently, a new development has been made in electron microscopy which enables us to observe the "living" myosin filament [16]). However, to visualize relative sliding under this setting, labeling (and fixation) of either of these molecules is necessary. The currently used experimental techniques can be classified into two categories based on the technique employed for labeling and fixation and the source of these proteins. One is the fluorescent actin filament system developed by Kron and Spudich [4] (Fig. 1A). In this system actin filaments are fluorescently labeled with rhodamine—phalloidin and introduced onto the myosin (or its subfragment) fixed on a glass coverslip. We can observe the ATP-dependent sliding movement of actin filaments under a fluorescent microscope and record it on videotapes. Because only purified proteins exist in this system, we can, not only control the experimental conditions precisely, but also study the effect of structural modifications made to the proteins either chemically or genetically.


Figure 1
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  		Fig. 1 Two types of in vitro motility assay techniques. A: Fluorescent actin filament system. An actin filament (AF) labeled with rhodamine—phalloidin slides over the myosin (M) layer fixed on a glass coverslip. B: Algal actin cable-based motility assay. A small latex bead (B) coated with myosin (M) moves along actin cables.

 
The other is an algal actin cable based motility assay developed independently by Sheetz and Spudich [2] and Shimmen and Yano [3] (Fig. 1B). In this system, well-organized actin cables (bundles of actin filaments) of algal cells are used as the substratum for the sliding movement. To visualize the movement of myosin molecules, small polystyrene beads (diameter – 1µm: visible under an ordinary light microscope) are coated with myosin and applied to the actin cables in the presence of ATP. When a bead comes in close contact with the actin cables, the myosin on its surface interacts with actin and pulls the bead. Although the structure and nature of algal actin have not been fully characterized, its straight arrangement makes the sliding movement also straight thus providing us with an opportunity to observe actin—myosin interaction under a steady-state condition.

Although these assay systems have been proven to be powerful tools for the study of actin—myosin interaction, the presented data also have created controversies some of which will be discussed below.

2.2 Step size
Determination of the step size (sliding distance of myofilament during the hydrolysis of a single ATP molecule) is a crucial issue for the study of the mechanism of muscle contraction. If the sliding movement is driven by the tilting movement of a myosin head attached to an actin filament, the step size should be at most 10 to 20 nm (the length of the myosin head). The value reported by Spudich's group using the fluorescent actin filament system was within this range [17,18]. On the other hand, Yanagida's group, using a similar experimental technique, proposed the value of 100 nm which is possible only by the multiple power strokes per one ATPase cycle [19]. The diversity may originate from the indirect estimation of the step size. The common theoretical framework used by these authors is as follows: [18,19]. The myosin step size (d) can be calculated from the velocity of actin filament sliding (Vs) and the time during which a myosin head is attached to actin and drive the filament sliding in one ATP hydrolysis cycle (ts) as:

Formula

Furthermore, ts is calculated as:

Formula
where tc is the ATPase cycle time (the time during which one ATP is hydrolyzed to ADP and Pi) and p is the ratio between ts and tc (duty ratio). The difference in the estimate of ATPase activity and the sliding velocity of actin filaments between the two groups led to this considerable diversity in results, but the causes of these differences are not clear [20].

More recently, direct measurement of d was made possible by the fluorescent actin filament system coupled with a glass micro-needle or a laser optical trap technique [5,6,21] (details will be described below). However, still considerable variation exists among the step size values reported by various research groups ranging from 4 to 17 nm. The smallest value (4 nm) has been reported by Molloy et al. [21] for skeletal muscle subfragment 1 (S1). However, this value was obtained as the shift in the mean value of Boltzman distribution, thus being not the result of direct measurement. For heavy meromyosin (HMM), larger values (11 nm by Finer et al. [5], 7 to 30 nm by Miyata et al. [22]) have been reported. We also note that in the same paper Molloy et al. reported the value of 7.9 nm for HMM. Furthermore, Ishijima et al. identified a unitary step size of 17 nm [6] for physiologically oriented myosin molecules (myosin-rod cofilaments). All these data can be taken to indicate that, although the essential part of the motor function resides in the head (S1) portion, the step size may increase as more complete forms of myosin molecules are used for the assay.

Very recently, Kitamura et al. [23], using a novel assay system, observed the displacements caused by single S1 molecules. The mean value of all the observed events was 13 nm, but each consisted of regular steps of 5.3 nm, suggesting multiple steps during one ATP hydrolysis cycle (chemo-mechanical loose coupling). If this loose coupling mechanism applies to a myosin power stroke under low load, it could be another source of variation in the step size.

2.3 Unitary force
Like the step size, measurement of the force generated by a single myosin molecule (unitary force) is of interest. Kishino and Yanagida used a compliant glass micro-needle to measure the force generated by actin and myosin [24]. They attached a single fluorescent actin filament to the tip of a glass micro-needle and brought it close to the myosin layer fixed on a glass coverslip. As the actin filament slid over the myosin layer, it pulled and bent the needle, the stiffness of which was pre-determined. From the density of the myosin head on a glass coverslip and the length of the filament, they estimated the force per myosin head averaged over one ATPase cycle (average force) at about 1 pN. This value was consistent with that estimated from the force generated by a muscle fiber and the density of the myosin head in it [25]. Later, they measured the unitary force using a similar experimental setting but over a myosin layer with much lower density (expecting that only one myosin head interacted with an actin filament) and reported a peak value (Fig. 3) of 5–6 pN [6].


Figure 3
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  		Fig. 3 (A) Unitary force of rat cardiac myosin recorded by an optical trap system. (B) Schematic diagram of the crossbridge cycling. A single crossbridge repeats cyclic interaction with an actin filament while splitting an ATP molecule.

 
Unitary force can be measured by the laser optical trap technique in which an infrared laser beam introduced through the objective lens of an inverted microscope creates a field of force at the focal point so that we can trap and manipulate a small particle. Finer et al. [5] attached two microspheres at both ends of an actin filament and trapped them by two independent optical traps. They brought this filament to myosin molecules sparsely fixed on a glass coverslip to observe a single event. By projecting the bright field image of the microsphere onto a quadrant photo-diode, they could monitor the actin—myosin sliding at nm resolution. Furthermore, by feeding back the position signal to a driver circuit which steered the laser beam, they could also measure the force under isometric conditions. With this system they clearly demonstrated that a single molecular event between actin and myosin took place in a pulse like fashion. The unitary force thus measured was about 3–4 pN. More recently, Molloy et al., using a similar system, reported the unitary force generated by heavy meromyosin to be 1.7 pN [21]. These values are a little smaller than that reported by Yanagida's group using a glass micro-needle (5.7 pN) [6]. They contended that the more physiological actin—myosin interaction in their experiment (actin filament vs. myosin filament) made the unitary force larger. In contrast to the step size, however, the unitary forces reported so far by various authors are not so divergent.


   3 Force—velocity relation in vitro and crossbridge kinetics
Top
 Abstract
 1 Introduction
 2 Basic characterization of...
 3 Force--velocity relation in...
4 Findings on cardiac...
 5 Clinical implications
 6 Future perspective
 References
 
Oiwa et al. studied the force—velocity relation in vitro using a centrifuge microscope [26]. This microscope, developed by Kamitsubo et al. [27], is equipped with a rotating stage and a stroboscopic light (Fig. 2A). After mounting an algal cell preparation in which myosin coated beads are sliding on straight actin cables at a constant velocity, they could apply a centrifugal force as a load to the moving bead by rotating the stage. As the load (rotation rate) was increased, the sliding velocity of the bead decreased. Finally, the bead stopped sliding under a load which was equal to the maximal force generated by myosin molecules on the bead. From the maximal force value, they estimated that the number of myosin molecules participating in the sliding would be only 1 to 20. The obtained force—velocity relation was hyperbolic in shape but its curvature switched in the middle range of the load (Fig. 2B). We can find similarly complex force—velocity relations in studies on single skeletal muscle fibers [28] and cardiac trabeculae [29], but the deviation from the single hyperbola was more pronounced in the in vitro motility assay.


Figure 2
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  		Fig. 2 Force—velocity relation in vitro. A: Schematic presentation of a centrifuge microscope system. The algal cell preparation is mounted on a rotating stage and a centrifugal force is applied to the moving bead as a load. VC: video camera; VTR: video-tape recorder; PC: pulse counter; PS: position sensor; PG: pulse generator; Xe: xenon lamp. B: Force—velocity relation in vitro obtained using a centrifuge microscope for rat cardiac myosin by Sugiura et al. [30].

 
To account for the strange shape of the force—velocity relation in vitro, Sugiura et al. proposed a model [30], the essence of which was the small number of myosin molecules involved and the small duty ratio (p) of each molecule of myosin. They assumed that the p value was very small under zero load and increased in proportion to the external load. This assumption was based on the Huxley's contraction model [31] and was supported by the stiffness measurement during muscle shortening [32]. Furthermore, small duty ratio values under unloaded sliding were reported in the in vitro motility studies [5,33].

Contradictory results have been reported by Yanagida's group [34]. In the in vitro motility assay using a glass micro-needle, they showed that the force fluctuation decreased as the actin filament slid faster on the myosin layer. This result implies that the duty ratio increases as the load is reduced, reaching 90% under the unloaded condition, which is thus in disagreement with Huxley's model. To explain this discrepancy, they contended that multiple working strokes during one ATP hydrolysis cycle can take place under low load (loose coupling hypothesis). This hypothesis was further supported by the following experiments [23,35].

3.1 Molecular mechanism
Although the in vitro motility assay has been a powerful tool in identifying and characterizing the single molecular event between actin and myosin, we have to combine other experimental techniques to understand the molecular mechanism underlying it. Rayment et al. [36] identified the 3-D structure of the head portion (subfragment 1) of chicken skeletal myosin with 2.8 Å resolution using X-ray crystallography. In this structure, the C-terminus region of subfragment 1 is composed of an {alpha}-helix about 9 nm long (lever) and a pair of light chains (essential and regulatory ones) are attached to it. Based on this finding, they proposed that a small structural change induced by hydrolysis of ATP in the ATP-binding site is amplified and transmitted to the lever portion of the molecule (lever arm model). This model was reinforced by the following work which clearly showed the conformational change among the S1 crystals bound with various nucleotide analogues (MgADP•BeFx, MgADP•AlF4, MgADP•vanadate, MgPPi, each of which corresponds to a different stage in ATP hydrolysis) [37]. Motion of the {alpha}-helix region was demonstrated by the measurement of a fluorescent signal emitted from the probes attached to the light chain (LC2) [38]. In vitro motility assay using recombinant proteins also gave a support to this idea. Uyeda et al. expressed mutant Dictyostelium myosins in which the length of the {alpha}-helix was made to half, twice, and three times that of the wild type and measured the sliding velocity of the actin filament on them [39]. Interestingly, the sliding velocity correlated with the length of these modified {alpha}-helices, thus suggesting the lever function of this segment.

Estimation of the step size can be approached by X-ray crystallography. Whittaker et al. [40] compared the structure of S1 (truncated form) of smooth muscle myosin with and without MgADP and concluded that the lever arm tilted by 23°. Considering the length of the "lever" ({alpha}-helix; 9nm), a rotation of 23° results in a step of 3.5 nm being consistent with the result obtained by Molloy et al. [21]. On the other hand, a recent study by Dominguez et al. [41], in which they analyzed the more complete form of smooth muscle S1, suggested a rotation of 70° which could easily accommodate the step size in the order of 10nm.

Currently, the lever arm model seems to dominate the research in this field. However, as it is known from the diversity in step size values, we have not obtained conclusive evidence on the molecular mechanism driving the relative sliding of actin and myosin. Further studies are necessary to solve this problem.


   4 Findings on cardiac myosin
Top
 Abstract
 1 Introduction
 2 Basic characterization of...
 3 Force--velocity relation in...
 4 Findings on cardiac...
5 Clinical implications
 6 Future perspective
 References
 
The in vitro motility assay has also proved to be a useful tool in the study of the mechanical property of cardiac myosin. In mammalian cardiac muscle, three myosin isoforms (V1, V2 and V3) have been recognized, depending on the mobility on gel electrophoresis [42]. Biochemical studies have revealed that V1 has the highest ATPase activity, whereas V3 has the lowest one. Structurally, V1 and V3 are homodimers of {alpha}- and β- heavy chains, respectively, and the V2 is a heterodimer of {alpha} and β. Since the expression of these isoforms is regulated by development and aging [43], hormonal stimulation [44], and mechanical overload [45–47], its physiological significance has been a focus of study. Although those studies using muscle fibers have shown that the unloaded shortening velocity correlated with the myosin isoform content [48–50], the complex membrane structure and regulatory proteins conserved in these preparations have made the interpretation of results difficult. To circumvent this problem, Yamashita et al. measured the sliding velocity of beads coated with myosin obtained from either control (predominantly V3) and hyperthyroid (predominantly V1) rabbit ventricular muscle [51]. The results clearly showed a positive correlation between V1 isoform content (and the ATPase activity) of the myosin preparation and the sliding velocity of beads, thus indicating a direct relation between myosin structure and its mechanical property.

Probably the more important mechanical characteristics about the actin—myosin interaction is its behavior under load. Sugiura et al. studied the force—velocity relation of beads coated with cardiac V1 or V3 isoform using a centrifuge microscope [30]. The obtained force—velocity relation was complex in shape as in the previous studies with skeletal muscle myosin [26]. Although the maximum velocity under zero load was about three times faster for the V1 isoform, the maximum load under which the bead stopped moving was not different between the two isoforms, suggesting an equal force generating ability. On the other hand, the shape of the force—velocity relation differed between V1 and V3, suggesting a distinct kinetic property of these isoforms.

Force measurement can be seriously affected by the number of force generating units, which is not readily determined in the algal actin cable based motility assay. In case of the sliding filament assay, however, the number of myosin heads interacting with an actin filament can be estimated by the length of the actin filament (visible by fluorescent labeling) and the density of myosin heads fixed on the glass coverslip (estimated from the ATPase activity on the coverslip). Using this assay system, Sugiura et al. [52] compared the force generating ability of two cardiac myosin isoforms to find no significant difference between them. This result was in contrast to that obtained by Van Buren et al. [53], showing a greater force generating ability of the V3 isoform obtained in a similar experiment. The reason for this discrepancy is not clear, but similarly diverse results have been reported by experiments comparing the force generated by ventricular papillary muscle or trabeculae whose myosin composition was predominantly V1 or V3 [54–56].

Recently, Sugiura et al. [57] directly measured and compared the unitary step and force generated by rat cardiac myosin isoforms using an experimental technique similar to those employed by Finer et al. [5], Miyata et al. [22], and Molloy et al. [21] (Fig. 3A). The step size they obtained was about 15 nm and did not differ between the two cardiac myosin isoforms (V1: 15.3 nm vs. V3: 14.9 nm), but V1 had a shorter duration of events (V1: 204.7 ms vs. V3: 282.7 ms, ATP concentration; 0.5 µM). The value they obtained (15 nm) was in the larger range of the step size values reported so far, probably because they used intact myosin molecules.

As for the unitary force, V1 and V3 also did not differ in amplitude (V1: 1.2 pN vs. V3: 1.6 pN), but duration was longer for V3 (V1: 332.7 ms vs. V3: 488.1 ms). This result implies that the crossbridge force—time integral (the product of the unitary force and its duration, Fig. 3B) is significantly smaller for the V1 isoform. In their series of studies on heat measurement, Alpert's group [58–60] suggested that the small force—time integral was the basis for the reduced economy of energy utilization by the V1 isoform. In this respect, the results of Sugiura et al. were consistent with Alpert's view and might be the molecular basis for the isoform redistribution as an adaptation process.

The reduced economy can also be described by considering the relation between the peak force (unitary force) and the average force. As is shown in Fig. 3A,B, each myosin head (cross-bridge) repeats the attached (force generating) state and the detached (non-force generating) state while splitting an ATP molecule and the average force (F) is calculated as the product of the unitary force (f) and the duty ratio (p). As for the cardiac myosin isoforms (V1 and V3), with similar unitary forces [57], the average force would be mainly influenced by the duty ratio. Because the V1 isoform, with a shorter duration of force-generating state (ts) is known to have a higher ATPase rate, i.e. shorter ATPase time (tc), the duty ratio (=ts/tc) and the average force may not be significantly different from those of the V3. However, to maintain a similar level of average force, the V1 isoform must consume more ATP per unit time.


   5 Clinical implications
Top
 Abstract
 1 Introduction
 2 Basic characterization of...
 3 Force--velocity relation in...
 4 Findings on cardiac...
 5 Clinical implications
6 Future perspective
 References
 
The importance of the cardiac myosin isoform redistribution discussed above is recognized only in small animals. However, actin myosin interaction has been shown to play a significant role in the pathogenesis of human cardiac disease.

Recently, mutations in the genes coding various sarcomeric proteins have been identified as the cause of familial hypertrophic cardiomyopathy (FHC) [7–12]. Among these, mutations in the β-myosin heavy chain (β-MHC) account for a significant part. Characteristically, all the mutation sites reported so far are clustered in the head portion, which is believed to be functionally important. To clarify the impact of mutant myosin on its contractile function, several groups of investigators have studied the property of mutant myosin from various sources.

Cuda et al. purified a mutant myosin (Arg403Gln) from the soleus muscle (slow twitch fiber expressing β-MHC) of FHC patients. They found a decrease in sliding velocity suggesting an abnormality in actin—myosin interaction [61]. Similar results have been obtained for Thr124Ile, Tyr162Cys, Gly256Glu, Val606Met, Arg870His, and Leu908Val mutations [62]. In vitro expression systems have also been used for the study. Sweeney et al. [63] expressed a mutant rat {alpha}-cardiac MHC using a baculo virus/insect cell expression system and reported a depressed sliding velocity. Human β-MHC has also been expressed by Sata and Ikebe [64] using a similar system. Again, a decrease in sliding velocity was observed for β-MHC mutants Arg403Gln, Arg249Gln, Arg453Cys, and Val606Met. All these studied strongly suggested that an abnormality in actin—myosin interaction was the cause of familial hypertrophic cardiomyopathy. Furthermore, Arg403Gln mutation which is associated with poor prognosis showed the lower actin sliding velocity than others with better prognosis. However, interpretation of filament sliding velocity in view of clinically available indices of cardiac performance may not be straightforward.

Fujita et al. [65] measured the force generated by mutant myosins expressed in Dictyostelium discoideum (slime mold) cell. Dictyostelium myosin, despite its low sequence identity (40%), has a crystal structure very similar to that of chicken skeletal muscle myosin, thus it can be a good model for the study of the β-MHC mutant [37]. The mutations Fujita et al. studied could be classified into three groups based on the functional properties. Three mutations, Ala699Arg (corresponding to human FHC Gly716Arg), Lys703Glu (Arg719Glu), and Lys703Try (Arg719Try), are on a short {alpha}-helix in the COOH terminal subdomain. The COOH-terminal subdomain has been shown to be part of the "lever" which swings against the remaining part of the motor domain of myosin. These mutated loci are also close to the essential light chain. Notably, this group of mutations are characterized by the lowest level of force generating ability. Clinically, patients with these mutations are known to have the worst prognosis among the mutations examined in this study. In the second group is the mutation Phe506Cys (Phe513Cys), located away from the actin-binding and ATPase sites, and is characterized by a nearly normal force generating ability. Patients with this mutation can expect good prognosis. The third group consists of two mutations Arg397Glu (Arg403Glu) and Gly575Arg (Gly584Arg). Arg403Glu is known to be part of the actin-binding site. Gly584Arg is located closely to the 50-K cleft and at a turn connecting two β-strands. This group of mutation shows moderately depressed force generating ability and its prognosis is between the first and the second. The authors speculated that the force generating ability could be a determinant of the prognosis. A depressed force generating ability can be compensated by an increase in ventricular wall thickness resulting in a pathologic hypertrophy.

In human heart, two types of {alpha}-actin isoforms are expressed, the amino acid sequences of which have been shown to differ at residues 2 and 3 by a Glu to Asp substitution and by substitutions of Met to Leu and Ser to Thr at positions 299 and 358 [66]. According to Sutoh et al. [67], the highly charged N-terminal region of actin (residues 1–4) is in contact with myosin and the alteration of these amino acids caused reduced sliding velocity of the actin filaments polymerized from these mutants on myosin in vitro. Hewett et al. [68] found a significant correlation between the {alpha}-skeletal actin content in the cardiac tissue and the contractility of the heart evaluated by the maximum rate of contraction (dP/dt) in the BALB/c mice model. These results may suggest the role of actin in the pathogenesis of cardiac disease. In fact, the relative content of skeletal and cardiac {alpha}-actin also changes in the human heart during the course of aging but no significant difference was detected between control and failing adult hearts [69]. Recently, mutations in cardiac actin gene (Arg312His and Glu361Gly) have been identified as the cause of hereditary dilated cardiomyopathy [13]. Since, however, these amino acids are located in the domain that attaches to the Z band, contractile dysfunction is probably introduced by an abnormality in transmission of force rather than the actin—myosin interaction itself.


   6 Future perspective
Top
 Abstract
 1 Introduction
 2 Basic characterization of...
 3 Force--velocity relation in...
 4 Findings on cardiac...
 5 Clinical implications
 6 Future perspective
References
 
Currently, more and more sophisticated experimental techniques are being developed so that we can dissect the crossbridge mechanics in more detail [23]. This line of information is useful not only for the study of the molecular mechanism of cardiac contraction and relaxation, but also for designing a new strategy for the treatment of heart disease. Especially, gene therapies targeting the contractile proteins can be an alternative for the pharmacological intervention. Further studies are eagerly needed.

Time for primary review 28 days.


   References
Top
 Abstract
 1 Introduction
 2 Basic characterization of...
 3 Force--velocity relation in...
 4 Findings on cardiac...
 5 Clinical implications
 6 Future perspective
 References
 

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