Impact of familial hypertrophic cardiomyopathy-linked mutations in the NH₂ terminus of the RLC on β-myosin cross-bridge mechanics

Abstract

Familial hypertrophic cardiomyopathy (HCM) is associated with mutations in sarcomeric proteins, including the myosin regulatory light chain (RLC). Here we studied the impact of three HCM mutations located in the NH₂ terminus of the RLC on the molecular mechanism of β-myosin heavy chain (MHC) cross-bridge mechanics using the in vitro motility assay. To generate mutant β-myosin, native RLC was depleted from porcine cardiac MHC and reconstituted with mutant (A13T, F18L, and E22K) or wild-type (WT) human cardiac RLC. We characterized the mutant myosin force and motion generation capability in the presence of a frictional load. Compared with WT, all three mutants exhibited reductions in maximal actin filament velocity when tested under low or no frictional load. The actin-activated ATPase showed no significant difference between WT and HCM-mutant-reconstituted myosins. The decrease in velocity has been attributed to a significantly increased duty cycle, as was measured by the dependence of actin sliding velocity on myosin surface density, for all three mutant myosins. These results demonstrate a mutation-induced alteration in acto-myosin interactions that may contribute to the pathogenesis of HCM.

the interaction of actin and myosin II in striated muscle is a tightly controlled and regulated system involving many proteins on the thick and thin filaments. Mutations in any of these proteins can greatly impact acto-myosin interactions and cardiac muscle function. The most common and hence most researched group of mutations is the hypertrophic cardiomyopathy (HCM)-associated mutations. These mutations are found in the myofilament proteins and result in enlargement (hypertrophy) of the heart, most commonly the left ventricular wall, interventricular septum, and in some cases the papillary muscles (22, 23).

Myosin, the molecular motor that powers muscle contraction, is a hexameric protein consisting of two heavy chains (∼200 kDa each) that self-associate via an alpha-helical coiled-coil domain. Two light chains, one essential light chain (ELC) and one regulatory light chain (RLC), are associated with each heavy chain. The entire myosin molecule can be broadly divided into two domains: the rod domain, which functions in filament formation, and the head domain, which has been shown to be sufficient for motion and force generation (16, 46). The myosin head domain is asymmetric with a globular portion, which contains the actin binding site and ATP hydrolytic activities (the motor domain), and an elongated alpha-helical MHC-neck domain that is supported by the binding of the two light chains (the neck domain).

The neck domain of the myosin head functions to amplify conformational changes originating in the globular motor domain via a lever-like action (2, 25, 34, 38, 51). Given its role in supporting the elongated neck region of the myosin head, it is not surprising that mutations in the regulatory light chain (RLC) were found to cause HCM. To date, several RLC mutations (see Fig. 1) have been linked to HCM (3, 27, 41), with five found near the COOH-terminal end (P95A, K104E, E134A, G162R, and D166V) (3, 27, 41), two located near the cation-binding domain (N47K and R58Q) (27, 41), and four that are located at the NH₂-terminal end of the protein (A13T, F18L, M20L, and E22K) (3, 27, 41). The RLC is important due to its position on the myosin molecule (4) and its suspected role in imparting stiffness to the neck region of the myosin head (32). Consistent with this idea, a single amino acid mutation in skeletal muscle RLC, F102L, resulted in a decreased actin filament velocity in the in vitro motility assays (39). The importance of the RLC was also noted in experiments involving β-myosin containing the N47K and R58Q RLC HCM mutations that demonstrated alterations in the ability to propel actin under load (10). A decrease in force was also observed in transgenic mice containing the COOH-terminal RLC mutant (D166V) (18, 28). On the other hand, myocardium from transgenic mice expressing 10% A13T mutant RLC displayed elevated contractile force (17). These observations strongly support the notion that the RLC acts either alone or in concert with the ELC to add stiffness to the lever arm region of the myosin II molecule.

Fig. 1.

The linear amino acid sequence of human cardiac ventricular regulatory light chain. In this figure the NH₂ and COOH termini of the molecule are shaded in gray, with the metal binding domain and the phosphorylatable serine underlined. The 11 known hypertrophic cardiomyopathy (HCM) mutations are indicated in bold and numbered. Note that 9 of the mutations reside in the gray-shaded areas with the other 2 mutations, N47K and R58Q, residing either in or near the metal binding domain. In this study we determined the mechanochemical effects of the A13T, F18L, and E22K mutant myosins, which are located near the NH₂ terminus of the molecule.

In addition to its role in stabilizing the neck region of myosin, the NH₂-terminal region and phosphorylation sites of the RLC have also been shown to be critical for the interaction of the myosin head with actin as well as impact the orientation of the myosin head in Drosophila (9). Furthermore, the phosphorylation state of the NH₂-terminal serine has been implicated in contractile function, including enhanced stretch activation response in myocardium (5, 40), increased calcium sensitivity of striated muscle contraction (31, 43), and structurally controlling myosin head orientation (8, 9). Therefore, the RLC molecule, in particular the NH₂-terminal region, may also play a role in the acto-myosin interactions.

To test the effects of the RLC-HCM mutations on the acto-myosin contractile activity, we produced myosin molecules containing the mutant light chains by exchanging bacterially expressed human cardiac RLCs onto purified porcine cardiac myosin. We used the in vitro motility assay to assess the effects of these NH₂-terminal RLC mutations on myosin motion generation. We found that the principal defect for the NH₂-terminal HCM mutant myosins is a loss in actin sliding velocity under low load. Measurements of solution ATPase activity and the dependence of actin filament velocity on myosin function indicated that the loss of velocity cannot be attributed to a decrease in the mutant myosin enzymatic activity but instead are a result of an increased duty cycle. These data demonstrate HCM mutation-induced changes in actomyosin interactions that could directly contribute to the disease phenotype.

METHODS AND MATERIALS

Solutions.

All solutions used for in vitro motility assays were made as described previously (10, 11) with some modifications. As shown in Table 1, N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES) replaced imidazole.

Table 1.

Composition of the buffers used in these experiments

Compound	Myosin Buffer	Actin Buffer	Actin + ATP	Motility Buffer
KCl	300 mM	55 mM	55 mM	55 mM
BES	25 mM	25 mM	25 mM	25 mM
MgCl2	4 mM	4 mM	4 mM	4 mM
EGTA	5 mM	5 mM	5 mM	5 mM
Glucose	0 mM	0 mM	0 mM	2 mM
ATP	0 mM	0 mM	1 mM	1 mM
Glucose oxidase	N/A	N/A	N/A	160 units
Catalase	0 mM	0 mM	0 mM	2 mM

Note that the buffer BES was used instead of imidazole due to its better buffering capacity.

Depletion of native porcine RLC and reconstitution with human ventricular WT/HCM RLC.

Endogenous porcine cardiac RLC was depleted from the myosin heavy chain using the protocol described previously (32) with modifications. Briefly, native porcine cardiac myosin was precipitated in 4°C water with 10 mM DTT for 1 h, and then centrifuged at 8,800 g for 30 min. The resulting pellet was resuspended in 500 mM KCl and 50 mM KH₂PO₄, pH 8.5, to a final concentration of 1.5–2.0 mg/ml. Triton X-100 and CDTA were added to this solution to a final concentration 1% and 5 mM, respectively. After a 15-min incubation at room temperature the reaction was terminated by diluting 10-fold in ice-cold water with 10 mM DTT followed by a 1-h incubation. The myosin was pelleted once again at 8,800 g for 30 min and the pellet resuspended in minimal reconstitution buffer (400 mM KCl, 50 mM BES, 2 mM MgCl₂, and 1 mM DTT). Myosin concentration was determined by a Bradford assay (Bio-Rad Labs, Hercules, CA). Depleted myosin was then reconstituted with bacterially expressed RLC (42) by incubation for 2 h at 4°C at a 3:1 RLC to myosin molar ratio. Reconstitution was terminated by precipitating the myosin in ice-cold 10 mM DTT. The reconstituted myosin was resuspended in minimal high-salt buffer (600 mM KCl, 50 mM BES, and 20 mM DTT) and mixed 1:1 with glycerol and stored at −20°C for up to 3 wk. The amount of exchanged RLC for each variant was determined using the ratio of RLC to ELC (in native, depleted, and reconstituted) as determined by SDS-PAGE and Coomassie stain. As shown in Fig. 2 the average amount of exchange for all four RLCs was 85.4 ± 4.2%, 81.3 ± 0.5%, 85.7 ± 5.5%, and 81.0 ± 1.9% for WT, A13T, F18L, and E22K respectively.

Fig. 2.

SDS-PAGE of native (lane 1), depleted (lane 2), and exchanged regulatory light chain (RLC) onto porcine cardiac beta myosin. As indicated by the gel the depletion reaction is able to remove ∼80–85% of the native RLC as indicated by densitometry traces, with the subsequent exchanges of WT, A13T, F18L, or E22K (lanes 3–6, respectively) yielding an average exchange of 85.4% ± 4.2%, 81.3% ± 0.5%, 85.7% ± 5.5%, and 81.0% ± 1.9%, respectively. ELC, essential light chain.

In vitro motility assay.

Experiments were performed as described previously (32) with some modifications. BSA concentration was increased from 0.5 to 1 mg/ml. Briefly, ∼20 μl flow cells were constructed by attaching nitrocellulose-coated coverslips and a standard glass slide with 100-μm-thick 3M double stick tape. Myosin and α-actinin (50 μg/ml) were mixed in myosin buffer (Table 1) and incubated in the flow cell for 1 min. This was followed by 25 μl of 1 mg/ml BSA, for 1 min, in myosin buffer to remove any unbound myosin and block the remaining surface sites to avoid nonspecific attachment of actin filaments. After blocking, 50 μl of actin buffer was flowed through the chamber, followed by the addition of unlabeled actin. This actin was allowed to incubate for 2 min before being washed out with 50 μl of actin + ATP buffer and then 100 μl of actin buffer to remove the ATP. TRITC labeled actin filaments in low-salt buffer was then added to the flow cell and allowed to incubate for 30 seconds. Unbound actin was washed out and motility initiated by the addition of low-salt buffer with 1 mM ATP and oxygen scavengers.

Myosin duty cycle.

To examine if the HCM mutants caused changes in myosin duty ratio, we measured the actin sliding velocity as a function of increasing myosin concentrations. In this assay, actin filaments are translocated at maximum velocity (V_max) if at least one head is interacting with the actin filament at all times. The probability of this occurring is related to the number of heads available to interact with actin (loading concentration, ρ) and the fraction of ATPase cycle time that myosin molecule remains bound, r (i.e., duty cycle). Assuming that the motors act asynchronously, the probability that at least one head is interacting with actin at all times is given by 1 − (1 − r)^ρ. Accordingly, at low motor surface densities, velocity will decrease as a function of r and ρ, according to the equation:

$$V_{\text{obs}}=V_{\text{max}}[1-{\left(1-r\right)}^{\rho }]$$ (1)

where V_obs is the observed velocity of the actin filament, and V_max is the maximal sliding velocity at saturating myosin concentrations (26, 48). Therefore, a myosin with a higher duty cycle will be able to move an actin filament at maximal velocity at a lower concentration, whereas a myosin with a lower duty cycle will require a greater concentration on the motility assay surface in order to ensure that at least one myosin head is attached to actin at any given time.

Data acquisition and analysis.

Movies of actin filament motion on the cover glass surface were captured at three frames per second on an intensified charge-coupled device (ICCD) camera (IC300; PTI, Birmingham, NJ) using software (XCAP) and PCIe card (PIXCI SV7) purchased from EPIX (Buffalo Grove, IL). All movies were analyzed using an ImageJ plug-in, wrMTRCK [written by J. S. Pedersen (jsp@phage.dk)]. This program is a centroid-tracking program that automatically locates and tracks objects within a movie and reports the velocity of the object. All filaments from a single movie that have been determined to be moving are then averaged to give a single value for that movie. The average velocity for each movie is considered an N of 1 for each movie collected. For both the myosin dose response, and α-actinin curves 10 movies were collected for each data point for each exchanged myosin with 4–5 exchanges run for all RLC variants tested here.

Actin-activated ATPase assays.

WT-RLC and the three HCM mutants were diluted 1:10 in 4°C water, 10 mM DTT for 1 h. Precipitated (filamentous) myosin was centrifuged for 30 min at 8,800 g. The resulting pellet was resuspended in a minimal amount of 10X salt buffer (800 mM KCl, 50 mM BES, and 10 mM DTT) and the protein concentration was determined using a Bradford assay (Bio-Rad Labs). Actin-activated ATPase activity was measured using the ATPase Kinetic ELIPA Assay Kit (Cytoskeleton, Denver, CO) that is based on an absorbance shift from 330 to 360 nm when 2-amino-6-mercapto-7-methylpurine ribonucleoside (MESG) is converted to 2-amino-6-mercapto-7-methyl purine in the presence of inorganic phosphate. The reaction was run according to the manufacturer's instructions with a slight modification: the reaction volume was doubled from 300 to 600 μl to accommodate the 1-ml cuvettes used to measure the absorbance at 360 nm. Prior to measuring the ATPase activity of the exchanged myosins, the three components of the reaction were mixed in the amounts recommended by the manufacturer and allowed to mix briefly for 5 min before pipetting 540 μl of the reaction volume into 1.5-ml microcentrifuge tubes. G-actin was added to the reaction tubes to a final concentration of 1.9 μM along with 55 μl of myosin with one of four RLC exchanged myosins (wild-type, A13T, F18L, or E22K) and incubated for 30 min at room temperature. For myosin-free controls 55 μl of high-salt buffer was added thereby mimicking the condition for myosin-containing samples. The ATPase reaction was initiated by the addition of ATP (final concentration 10 mM). MESG, and thus phosphate production, was monitored by absorbance at 360 nm at room temperature. Measurements were made every 20 s for 20 min and the resulting measurements, two or three runs for each mutant myosin, were pooled to obtain the average ATPase for each myosin.

Statistics.

Statistical analysis referred to in this paper was performed using Prism 6 (GraphPad 7825, La Jolla, CA). One-way ANOVA was used to compare the actin sliding velocities between WT and HCM mutants at all amounts of applied α-actinin. Significance between the curves was determined using the compare function within the analyze window using the extra sum-of squares F-test. Significance was determined if the returned P value was <0.05.

RESULTS

Impact of RLC HCM mutations on sliding velocity.

To examine the direct impact of the NH₂-terminal HCM mutations on the motility and kinetics of β-myosin, we performed experiments on isolated porcine β-myosin, which is 97% identical to human β-myosin, in which the native cardiac RLC was exchanged with the recombinant human ventricular WT and mutant RLC. As described previously, we found that RLC depletion and reconstitution had no statistically significant effects on steady-state ATPase activity and restored actin filament velocity to near untreated myosin values (32).

We used the in vitro motility assay to assess the mechanical properties of the myosin mutants used in this study. As shown in Fig. 3, all three NH₂-terminal RLC mutations have significantly reduced unloaded actin sliding velocities when compared with WT-exchanged myosin. Porcine RLC is 97% identical to human RLC and control experiments show no significant differences in actin filament velocity between porcine myosin reconstituted with human or porcine RLC (not shown). To further scrutinize the impact of these mutations on actin sliding velocity, we introduced frictional loads with increasing amounts of the actin binding protein α-actinin. Under little to no frictional load (0–1 μg/ml α-actinin), all three mutants had significantly reduced velocities, indicated by the closed star. Increasing the frictional load reduced the velocities of all four strains with significant differences between WT and two of the mutants (A13T and F18L; open star), Table 2 gives the significance values compared with WT for the sliding velocities. At the higher loads (>3 μg/ml α-actinin) the velocities of the different RLC containing myosin constructs converged to a similar value as observed for WT, indicating similar force-generating capability.

Fig. 3.

Impact of NH₂-terminal HCM RLC mutations on actin sliding velocity with varying amounts of frictional load. Note that the unloaded actin filament velocities driven by the mutant myosins are reduced compared with WT (P < 0.001). Similarly, actin filament velocities were significantly slower than wild type under low frictional load (up to 2 μg/ml α-actinin; solid stars; open stars indicate significance for A13T and F18L only). Yet, when the frictional load is increased beyond 3 μg/ml the actin sliding velocity for the mutants is not significantly different from the wild-type. Except for F18L at 6 μg/ml α-actinin (N = 9), all points represent the average of more than 25 movies; error bars represent SE.

Table 2.

P values of the frictional loaded assay illustrating the level of significance in the difference in sliding velocities relative to the wild-type sliding velocity

	Strain
α-Actinin, mg/ml	A13T	F18L	E22K
0	***	***	***
0.5	***	***	***
1	***	***	***
2	*	***	NS
3	***	***	NS
4	NS	NS	NS
5	NS	NS	NS
6	NS	NS	NS

^***P < 0.001,

^*P < 0.05. NS, not significant.

Actin-activated ATPase.

In the in vitro motility assay the actin filament velocity can be affected by changes in ATPase activity and/or duty ratio; therefore we measured these parameters to determine the underlying mechanism for the HCM-mutation-induced changes in velocity (12). To determine if the reduction in unloaded velocity observed with the HCM mutations resulted from a reduced ATPase activity, we examined the actin-activated ATPase activity of all four strains of RLC. As shown in Fig. 4, the ATPase activity of β-myosin containing exogenous human RLC demonstrated no significant difference in ATPase activity between WT and the three RLC mutants.

Fig. 4.

Actin-activated ATPase activity of β-myosin bearing WT and HCM-exchanged RLC. The ATPase activity of all four RLC exchanged β-myosins were not significantly different (average from 3 exchanges); error bars represent SE.

Myosin duty cycle.

As shown in Fig. 5A, actin filament velocity was independent of myosin concentration over a broad range (50–200 μg/ml) for all of the myosins studied. Consistent with the data presented in Fig. 3, all three mutants (V_max = 0.414 ± 0.008, 0.404 ± 0.004, and 0.404 ± 0.006 for A13T, F18L, and E22K, respectively) were significantly slower than WT (V_max = 0.496 ± 0.009). As the concentration of myosin loaded onto the motility surface was lowered below 50 μg/ml, the normalized velocity for all strains decreased but the velocity at low myosin concentrations was higher for the mutants compared with WT (Fig. 5B). The relationship between actin filament velocity and surface density was used to estimate duty cycle, r (Eq. 1). The calculated r values are presented (see Fig. 5 legend). The HCM-RLC mutant myosins showed a significantly higher duty cycle when compared with the wild-type control myosin (Fig. 5B). This result indicates that, consistent with the inverse relationship between duty cycle and velocity (30), the mutation-induced increase in duty cycle contributes to the reduced V_max.

Fig. 5.

Effect of HCM mutations on the duty cycle of β-myosin. As shown in A, consistent with the results from Fig. 1, the calculated V_max for the 3 HCM mutants were significantly (P < 0.0001) slower than WT-RLC exchange myosin (0.496 ± 0.009, 0.414 ± 0.008, 0.404 ± 0.004, and 0.404 ± 0.006 mm/s for WT, A13T, F18L, and E22K, respectively), as indicated by the black star. B: normalizing the sliding velocities to the calculated V_max illustrates that all 3 HCM mutants had a significantly (P < 0.01) larger duty cycle (6.3± 0.6%, 6.2± 0.3%, and 6.0± 0.4% for A13T, F18L and E22K, as shown in inset of B) than the wild-type exchanged myosin (4.4± 0.3%) (as indicated by the gray star). Data from 0-50 μg/l are plotted in the inset of B. For WT, A13T, and E22K there were 15–25 measurements per myosin concentration while for F18L the N was 25–35 movies for each point; error bars represent SE.

DISCUSSION

HCM is a disease of the myocardium manifested by hypertrophy, myofibrillar disarray, and fibrosis and was shown to result from mutations in sarcomeric proteins. In this report, we examined the impact of three of the four known NH₂-terminal RLC-HCM mutations on the kinetics and mechanical properties of isolated myosin using the in vitro motility assay. To determine the effect of the mutations on acto-myosin contractile activity, reconstituted myosin preparations with the endogenous regulatory light chain extracted and replaced with bacterially expressed mutant RLCs were generated. The actin filament velocity, actin-activated ATPase, and duty cycle of the mutant myosins were examined to elucidate the mechanical and biochemical properties altered by the mutations. The results provide crucial information about the initial effects of these mutations on acto-myosin interactions, which ultimately may lead to the disease phenotype.

The RLC is found ubiquitously expressed in all muscle myosins and is proposed to play an important role in stiffening of the myosin neck region (lever arm) (19, 20, 47). Since the neck region of the myosin head is thought to undergo large changes in orientation during myosin motion generation (13, 21, 33, 36, 37), we hypothesized that the RLC-HCM mutations may disrupt actin filament velocity by either altering its mechanical properties or its biochemistry.

This present study demonstrates that the A13T and F18L mutations impose dramatic reductions in unloaded shortening velocity at saturating myosin surface densities (Fig. 3). On the other hand, the myosin bearing the E22K mutation was shown to produce a much more subtle effect (Fig. 3), strongly indicating that the amino acids around the phosphorylatable serine, along with the phosphorylatable serine, play an important role in stabilizing the lever arm region of the myosin molecule. No significant differences in myosin ATPase activity were observed between WT and the three RLC mutants (Fig. 4). Interestingly, differences in velocity observed for the NH₂-terminal HCM mutants were minimized by the addition of an α-actinin-induced frictional load to the motility assay. These reported results are consistent with those published earlier for transgenic A13T mice (17), but differ when compared with results reported for transgenic E22K mice (7, 45). These differences are likely due to the myosin heavy chain isoform (porcine β vs. mouse α) bearing the mutant RLC. Therefore, in contrast to what has been observed for the more malignant mutations in the RLC, R58Q, (10) and D166V (29), the NH₂-terminal RLC-HCM mutations do not appear to disrupt transfer of load to the active site.

There is precedence for mutations in the RLC resulting in a decrease in actin sliding velocity. Respective mutations in Dictyostelium myosin, E12T, and G18K also showed an unloaded contractile defect, evident as a ∼50% reduction in actin filament velocity with no change in ATPase activity (4). Similarly, the Dictyostelium RLC mutant, N94A, exhibited defects in cytokinesis as well as a dramatic reduction in ATPase activity and actin filament velocity. Furthermore, while not an HCM mutation, a single amino acid substitution in skeletal muscle myosin, F102L, resulted in decreased actin filament velocity (39).

Despite reductions in unloaded shortening velocity at saturating myosin surface densities, the RLC-HCM mutations studied here reached maximal sliding velocity at lower myosin concentrations compared with WT-RLC, indicating a higher unloaded duty cycle. This, combined with the fact that no change was seen for the actin-activated ATPase activity for the mutants compared with WT (Fig. 4), indicates that the reduced unloaded velocity observed for myosin containing the RLC-HCM mutations results, at least in part, from an increase in duty cycle. A decreased velocity and an increase in duty cycle are both consistent with a reduction in the acto-myosin detachment rate (14).

Because of the absence of the NH₂-terminal region of the RLC from the atomic structure of myosin S1 (35), little is known about the interaction of the NH₂-terminal region of the RLC with the rest of the myosin molecule. Mechanically, alterations in the NH₂-terminal portion of the RLC in Drosophila indirect flight muscle (IFM), although larger than vertebrate RLC, reduce work and power output in these muscles (6, 15, 24) potentially by altering the proximity of the myosin head to the thin filament (20). Furthermore, substituting the phosphorylatable serines with alanines was also shown to significantly alter myosin head orientation (9). These results indicate that alteration of the RLC NH₂ terminus, especially the regions around the phosphorylatable serines, greatly impacts the ability of myosin to interact with actin.

Consistent with this notion, structural studies of myosin thick filaments obtained from human biopsy samples from patients carrying the E22K mutation revealed myosin head disorder when compared with thick filaments from control human fibers (20). RLC phosphorylation has also been observed to affect myosin head orientation in a similar way as the E22K mutation (20), suggesting an important role for the NH₂-terminal region of the RLC in head mobility and the ability to bind the thin filament. HCM mutation-induced alterations in myosin head orientation would allow for increased attachment to actin, which is consistent with the increased duty cycle observed here. Thus, alteration in the ability of the myosin heads to bind the thin filament could also be a contributing factor for the NH₂-terminal RLC-HCM mutant contractile deficits (17) rather than from a loss in power stroke (39) or load transfer (10, 49).

In transgenic mice, disease-causing mutations of the RLC have shown alterations in contractility and faithfully recapitulated the disease phenotype at the heart level with myocyte disarray and fibrotic lesions (1, 17, 28, 30). In contrast to the results presented here, skinned muscle fibers from transgenic mice expressing E22K (17) and A13T (7, 45) mutant RLC show decreases and increases in force, respectively. Further complicating interpretation of these data, fibers extracted form transgenic mice expressing 10% mutant A13T RLC showed a 25% decrease in ATPase. This dominant effect of A13T RLC protein on myocardial function indicates that higher-order effects of the mutation at the fiber level (myocyte disarray, fibrosis, post-translational modification of contractile proteins) contribute to the observed phenotype. Indeed, endogenous mutant proteins can be posttranslationally modified to help restore normal function (1, 17, 18, 28, 30, 44). The uniqueness of our approach in using recombinant mutant proteins reconstituted with β-myosin heavy chain allows for the study and definition of the initial causative mutation-induced disruptions without the secondary effects that often confound the interpretation. Further, it should be noted that the results found here are particular to the β-myosin isoform and may not reflect the impact these mutations may have on α-myosin, the primary isoform expressed in adult mice. In this study we have determined that the principal contractile defect in the NH₂-terminal HCM mutant myosin is a loss in actin sliding velocity due to an increase duty cycle. Unlike the cation-binding mutants or mutants near the COOH terminus, the NH₂-terminal mutants do not manifest a severe disruption in strain dependent kinetics.

Given the proximity between the mutations and the RLC phosphorylation site it is conceivable that the mutations may affect the degree of phosphorylation. Consistent with this notion, several disease-linked RLC mutations have been associated with altered RLC phosphorylation(18, 28, 30, 50). Further, phosphorylation of the RLC has been shown to reverse some of biochemical and structural alterations of the isolated mutant RLCs (42). Future studies are needed to address the effectiveness of mutant RLC as a substrate for myosin light chain kinase as well as phosphorylation as a potential compensatory mechanism for mutation-induced alterations in myosin function (50).

GRANTS

This work was supported by National Heart, Lung, and Blood Institute Grants HL-071778 and HL-108343 (D. Szczesna-Cordary) and HL-077280 (J. R. Moore).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: G.P.F., P.M., K.K., D.S.-C., and J.R.M. conception and design of research; G.P.F. performed experiments; G.P.F. analyzed data; G.P.F. and J.R.M. interpreted results of experiments; G.P.F. prepared figures; G.P.F. and J.R.M. drafted manuscript; G.P.F., P.M., K.K., D.S.-C., and J.R.M. edited and revised manuscript; G.P.F., D.S.-C., and J.R.M. approved final version of manuscript.