Manipulating myosin to curb cardiomyopathy Hypertrophic cardiomyopathy (HCM, thickened heart muscle) is a disease typically caused by mutations in sarcomere genes. Sarcomeres are the functional and structural units in muscle that allow for contraction and relaxation. Toepfer et al. investigated how mutations in cardiac myosin-binding protein C (encoded by MYBPC3) alter cardiac muscle contraction and relaxation. Using mouse models and human muscle fibers, they showed that MYBPC3 mutations increased contractility and decreased relaxation by disrupting myosin conformations. Treating cardiomyocytes with a myosin allosteric inhibitor corrected the relaxation, contraction, and myosin conformation deficits, suggesting that the myosin inhibitor could be therapeutic for HCM.

Abstract The mechanisms by which truncating mutations in MYBPC3 (encoding cardiac myosin-binding protein C; cMyBPC) or myosin missense mutations cause hypercontractility and poor relaxation in hypertrophic cardiomyopathy (HCM) are incompletely understood. Using genetic and biochemical approaches, we explored how depletion of cMyBPC altered sarcomere function. We demonstrated that stepwise loss of cMyBPC resulted in reciprocal augmentation of myosin contractility. Direct attenuation of myosin function, via a damaging missense variant (F764L) that causes dilated cardiomyopathy (DCM), normalized the increased contractility from cMyBPC depletion. Depletion of cMyBPC also altered dynamic myosin conformations during relaxation, enhancing the myosin state that enables ATP hydrolysis and thin filament interactions while reducing the super relaxed conformation associated with energy conservation. MYK-461, a pharmacologic inhibitor of myosin ATPase, rescued relaxation deficits and restored normal contractility in mouse and human cardiomyocytes with MYBPC3 mutations. These data define dosage-dependent effects of cMyBPC on myosin that occur across the cardiac cycle as the pathophysiologic mechanisms by which MYBPC3 truncations cause HCM. Therapeutic strategies to attenuate cMyBPC activity may rescue depressed cardiac contractility in patients with DCM, whereas inhibiting myosin by MYK-461 should benefit the substantial proportion of patients with HCM with MYBPC3 mutations.

INTRODUCTION Hypertrophic cardiomyopathy (HCM) is a heritable disease of heart muscle affecting ~1 in 500 (1) individuals. Patient symptoms can be minimal or relentlessly progressive with resultant heart failure and/or sudden cardiac death (2). Adverse clinical outcomes in HCM increase with disease duration, thereby underscoring the importance of therapeutic strategies to abate disease progression (3). Dominant pathogenic variants in eight sarcomere genes cause HCM, but predominate in MYBPC3 and MYH7 [encoding β-cardiac myosin heavy chain (β-MHC)] (4). The overwhelming majority of HCM founder mutations (5–11), including one affecting 4% of South Asians (12), resides in MYBPC3. All HCM mutations in MYH7 encode missense substitutions (4), and mutant myosins are incorporated into the sarcomere. By contrast, most MYBPC3 mutations are truncating and are predicted to cause haploinsufficiency of cardiac myosin-binding protein C (cMyBPC) (13, 14). The mechanisms by which distinctive mutations in these two sarcomere proteins uniformly produce hyperdynamic contraction and poor relaxation (diastolic dysfunction) in advance of the morphologic remodeling in HCM (15–17) remain incompletely understood (18). Biophysical analyses demonstrate that HCM mutations in β-MHC, the molecular motor of the sarcomere, can increase adenosine triphosphatase (ATPase) activity, actin-sliding velocity, and power. Structural analyses predict that these interfere with the myosin IHM (interhead motif), shifting dynamic conformations of relaxed paired myosin molecules (19–21). These conformations are denoted as (i) disordered relaxation (DRX), a state where only one myosin head could be active, able to hydrolyze adenosine triphosphate (ATP) and potentiate force and (ii) super relaxation (SRX), a state of dual inactivation of myosins with both ATPases inhibited. The IHM is an evolutionarily conserved motif that is found in all muscle myosins and in primitive animals with nonmuscle myosin II, indicating the importance of inhibiting myosin during relaxation (22). cMyBPC has structural and functional roles in sarcomere biology (23). cMyBPC is generally thought to serve as a brake that limits cross-bridge interactions (23) through its biophysical interactions of its amino and carboxyl termini with both myosin (22) and actin (23). Phosphorylation of the amino terminus of cMyBPC reduces myosin interactions and increases ATPase activity and actin interactions to promote cross-bridge formation (24), events that are reversed by calcium concentrations that maximally activate thin filaments (25, 26). Hence, the phosphorylation state of cMyBPC is hypothesized to regulate the number of myosin heads available for force production (24). Interpreting these interactions in the context of human HCM mutations that reduce cMyBPC expression is complex for several reasons. Cardiac histopathology and in vivo function of Mybpc3t/+ mice, which genetically recapitulate human HCM mutations, are indistinguishable from wild type (WT) mice. Mybpc3t/t mice have a developmental defect in the normal pathways for cytokinesis that results in increased numbers of cardiomyocytes that are mononuclear (27, 28), contributing to ventricular dilatation and decreased contractile force (25, 29). Recent studies also demonstrate that loss of cMyBPC alters proportions of relaxed myosin in DRX and SRX conformations (30, 31), but whether this relates to contractility is unknown. To better understand how MYBPC3 mutations cause HCM, we assessed sarcomere function in the setting of cMyBPC deficiency and genetically altered myosin or pharmacologically attenuated myosin ATPase activity. In combination, these assays shed light on unifying mechanisms that drive HCM pathophysiology and demonstrate that a single pharmacologic manipulation of myosin corrects sarcomere dysfunction caused by MYBPC3 mutations, thereby providing a promising avenue for treating this prevalent cause of human HCM.

RESULTS We studied three mouse models with altered cMyBPC expression (fig. S1). Mybpc3t/+ and Mybpc3t/t mice carry endogenous heterozygous or homozygous truncating mutations (32, 33) and express graded reductions of cMyBPC protein. MyBPC-RNAi (RNA interference) (27) are WT mice transfected (P10) with a cardiotropic adenoassociated virus serotype 9 (AAV9) carrying green fluorescent protein (GFP) and RNAi targeting Mybpc3 transcripts. Injection of 5 × 1011 viral genomes (vg)/kg reduced Mybpc3 transcripts to less than 10% of quantities in WT mice and abolished protein expression (fig. S1 to S4). This strategy bypassed the developmental defect in Mybpc3t/t mice and also excluded potential functional effects of a small amount of truncated residual cMyBPC protein in homozygous mice (fig. S1, C and D). We also studied heterozygous (Myh6F764L/+) and homozygous (Myh6F764L/F764L) mice, which carry a human MYH7 mutation that causes dilated cardiomyopathy (DCM). This mutation reduces both actin-activated myosin ATPase activity and actin filament velocity (34, 35). In vivo cardiac phenotypes in Mypbc3 mutant mice Previous studies demonstrate that Mybpc3t/+ mice have normal cardiac contractility and minimally increased left ventricular (LV) wall thickness in comparison to WT. cMyBPC phosphorylation was comparable to that in WT mice (fig. S1, E to G). Mybpc3t/t mice have increased LV volumes and mass but depressed contractile function due in part to increased numbers of cardiomyocytes from additional perinatal divisions before permanent exit from the cell cycle (27). MyBPC-RNAi mice exhibited minimal LV hypertrophy (fig. S2, A to D) (27), without changed ventricular volumes or fractional shortening (FS), an in vivo measure of contractility. The graded loss of cMyBPC in these mice produced less hypertrophy than occurs in other HCM mouse models (3). Contractility and relaxation in cardiomyocytes from Mypbc3 mutant mice Because in vivo contractility and relaxation reflect sarcomere performance, as well as myocardial geometry, histopathology, and hemodynamic load, we studied ex vivo cardiomyocytes to assess biophysical functions of sarcomeres with altered cMyBPC expression (Fig. 1). Isolated cardiomyocytes from at least four mice of each genotype were studied. Cardiomyocytes from Mybpc3t/t mice were more fragile and more heterogeneous than in other models. Fig. 1 Contractile characterization of cMyBPC mouse models. (A) A schematic depiction of the WT sarcomere with normal cMyBPC integration (left side) and the consequences of mutations that deplete cMyBPC quantities in the sarcomere (right side). (B) Representative contractile waveforms from isolated cardiomyocytes from mice paced at 1 Hz. SLs of isolated cardiomyocytes were tracked to define the percentage shortening per cell and duration of relaxation. Each trace is the averaged waveform across all cells (numbered in parenthesis) analyzed for each genotype: MyBPC-RNAi (34), Mybpc3t/t (13), Mybpc3t/+ (61), and WT (30). (C) Comparisons of cellular shortening of isolated cardiomyocytes (numbered in parenthesis) from four mice with different genotypes: MyBPC-RNAi (36), Mybpc3t/t (23), Mybpc3t/+ (118), and WT (53). Data are plotted as mean ± SEM. (D) Measures of duration from peak contraction to relaxation in seconds plot as mean ± SEM (cells analyzed: MyBPC-RNAi (34), Mybpc3t/t (13), Mybpc3t/+ (61), and WT (30). Resting sarcomere lengths (SLs) (fig. S3) were comparable in cardiomyocytes from WT and Mybpc3t/+ mice, modestly decreased in cells from MyBPC-RNAi mice, and variable in cardiomyocytes from Mybpc3t/t mice. We assessed SLs throughout the contractile cycle to define percent shortening, a surrogate for systolic function (Fig. 1, B and C). Cardiomyocytes isolated from naïve or sham-RNAi mice had comparable cell shortening, thereby excluding an effect of AAV9 on contractility (fig. S2D). Cardiomyocytes with altered cMyBPC expression had dosage-dependent increases in maximal cellular shortening. In comparison to WT, Mybpc3t/+ cardiomyocytes had 50% increased shortening (7.2 ± 0.25 %; P < 0.0001), whereas cellular shortening was increased by 100% in cardiomyocytes lacking cMyBPC (MyBPC-RNAi: 10 ± 0.6%; Mybpc3t/t: 9.5 ± 0.9%; P < 0.0001). Augmentation of cellular shortening of isolated cardiomyocytes with cMyBPC deficiency did not result in increased contractility in vivo (27, 32), an observation that implies that additional (biochemical, transcriptional, and morphologic) processes can modulate ensemble systolic performance of cardiomyocytes. Human HCM is characterized by impaired diastolic performance, a parameter that is difficult to assess in mice. Instead, we tracked SLs in isolated cardiomyocytes across the contractile cycle to assess the duration of relaxation as a quantitative proxy for diastolic function (Fig. 1D). Relaxation was prolonged in MyBPC-RNAi and Mybpc3t/t cardiomyocytes [0.31 ± 0.02 (P < 0.0001) and 0.38 ± 0.06 (P < 0.0056), respectively] compared to WT cells (0.21 ± 0.02), whereas the duration of relaxation in Mybpc3t/+ cardiomyocytes was indistinguishable from WT (0.19 ± 0.02). Genetic repression of myosin function corrects cMyBPC deficiency Rare myosin missense mutations cause DCM, a disorder characterized by ventricular enlargement and diminished cardiac contractility. Mice engineered to carry the human DCM mutation (Myh6F764L/+ and Myh6F764L/F764L) recapitulate these phenotypes (34). Analyses of isolated cardiomyocytes from these models showed a genotype-dependent depression of cellular shortening (Fig. 2A); contractility in Myh6F764L/+ and Myh6F764L/F764L cardiomyocytes was 75 and 50%, respectively, of normal (P < 0.0001 for each). DCM cardiomyocytes also had small but significantly reduced durations of relaxation (Fig. 2B) [Myh6F764L/+, 0.17 ± 0.01 (P = 0.0002); Myh6F764L/F764L, 0.14 ± 0.005 (P < 0.0001)] compared to WT cardiomyocytes (P = 0.21 ± 0.02). Fig. 2 Genetic and pharmacological manipulation of cardiomyocytes depleted for cMyBPC. (A) Sarcomere contractility (cellular shortening) in isolated cardiomyocytes (numbered in parenthesis) from WT (63), Myh6764/+ (55), Myh6F764L/F764L (71) (abbreviated as Myh6764/+ and Myh6764/764), Myh6764/764 + MyBPC-RNAi (43), and MyBPC-RNAi (36) mice. (B) Sarcomere relaxation of isolated cardiomyocytes (numbered in parenthesis) from WT (30), Myh6764/+ (45), Myh6764/764 (29), Myh6764/764 treated with MyBPC-RNAi (31), and MyBPC-RNAi (45) mice. Individual data points are plotted with mean ± SEM indicated. All significant P values are indicated on the graph. (C) Sarcomere contractility of cardiomyocytes treated with MYK-461 (0.03 to 0.3 μM). More than 20 cardiomyocytes were analyzed for each drug concentration and treatment group. (D) Sarcomere relaxation of cardiomyocytes treated with MYK-461 (0.03 to 0.3 μM). All data are displayed as mean ± SEM. *P < 0.01 and **P < 0.0001 denote comparisons with WT without MYK-461. To determine whether manipulation of myosin properties would alter contractile phenotypes in Mybpc3 mutant cardiomyocytes, we injected AAV9 carrying GFP and MyBPC-RNAi into P10 Myh6F764L/F764L mice. Forty days after injection, the cellular shortening and duration of relaxation (Fig. 2, A and B) were significantly improved in comparison to MyBPC-RNAi cardiomyocytes (P < 0.0001); both contractile and relaxation parameters were indistinguishable from WT cardiomyocytes. Inhibition of myosin ATPase corrects cMyBPC defects in cardiomyocytes and cardiac tissues Because genetic deficits in myosin normalized the performance of Mybc3-deficient cardiomyocytes, we hypothesized that MYK-461, a cardiac-selective, pharmacologic allosteric myosin ATPase inhibitor, would also be effective. To initially determine whether MYK-461 (2.5 mg/kg per day via drinking water) elicited deleterious effects in Mybpc3t/+ and MyBPC-RNAi mice, we used in vivo echocardiography to assess LV posterior wall dimensions and FS 5 and 20 weeks after dosing. There were significant changes in cardiac morphology when comparing treated and untreated mice within genotypes for FS at 20 weeks in WT, Mypbc3t/+, and MyBPC-RNAi cohorts (P < 0.05) (table S1). On the basis of our findings that isolated cardiomyocytes from Mypbc3t/+ and Mypbc3t/t mice exhibited contractile differences that were not apparent from in vivo mouse imaging, we then acutely treated WT and mutant cardiomyocytes with MYK-461. The hypercontractility in Mypbc3t/+ and Mypbc3t/t cardiomyocytes was reduced in a dose-dependent manner (Fig. 2C). Two-way analysis of variance (ANOVA) within treatment groups also showed a reduction in cellular shortening after MYK-461 treatment. Contractile function was normalized in Mypbc3t/+ cardiomyocytes at 0.15 μM MYK-461 and at 0.3 μM in Mypbc3t/t cardiomyocytes. This higher dose of MYK-461 reduced sarcomere contractility by ~50% in both mutant genotypes and depressed contractility by ~30% in WT cardiomyocytes. Concurrently, 0.3 μM MYK-461 normalized relaxation times in Mypbc3t/t cardiomyocytes (Fig. 2D) but did not alter the duration of relaxation in Mypbc3t/+ or WT cardiomyocytes. Increased ratios of myosin heads in DRX:SRX caused by cMyBPC deficiency are normalized by modulation of myosin ATPase The proportions of myosin heads in DRX and SRX conformations correlate with the rate of ATP cycling in relaxed muscle, which can be measured by the decay of a fluorescent, nonhydrolyzable ATP (Mant-ATP) from skinned muscle (36) or cell fibers (Fig. 3 and fig. S5). Myosin heads in the DRX configuration have ~5× more ATPase activity than myosin heads in the SRX configuration (36). Hence, the fraction of myosin heads in the SRX configuration and DRX configuration can be estimated from the fraction of Mant-ATP that is released rapidly (DRX) or slowly (SRX). Fig. 3 Mant-ATP assessment and correction of SRX and DRX ratios in mouse and human myocardium. (A) Average Mant-ATP fluorescence decay curves plot from fluorescence decay due to dark ATP wash with acquisition duration of 5 min. Data points are the mean of ~12 separate experiments from three separate individuals in each genotype/treatment group. Data are fit by a double exponential decay to assess ratios of DRX and SRX heads. (B) Plot of the initial rapid decay amplitude corresponding to DRX heads. (C) Plot of the second exponents slow decay amplitude corresponding to SRX heads. (D) Average Mant-ATP fluorescence decay curves of unrelated human hearts: three without HCM (WT) and three HCM with MYBPC3t/+ mutations. Each curve is the average of 12 experiments from three separate samples in each treatment group. Data are fit by double exponential decay to assess ratios of DRX and SRX heads in the myocardium. (E) Plot of the initial rapid decay amplitude corresponding to DRX heads. (F) Plot of the second exponents slow decay amplitude corresponding to SRX heads. All data are presented as mean ± SEM with significances indicated with P values. Assays of skinned cardiac fibers from WT, Mypbc3t/+, and Mypbc3t/t mice showed different proportions of myosins in DRX and SRX confirmations (Fig. 3, A to C). Compared to WT cardiac tissues, Mypbc3t/+ and Mypbc3t/t had a 50% increase (P = 0.02) and 94% increase (P = 0.001), respectively, of myosins in DRX (Fig. 3B), changes that paralleled the dose-dependent increase in cardiomyocyte contractility (Fig. 1C). The proportions of SRX and DRX found in Myh6F764L/F764L (fig. S6) were comparable to those of WT mice (P = 0.053). This finding, combined with previous biophysical analyses of Myh6F764L/F764L molecules (34, 35), indicated that the depressed contractility associated with this DCM genotype largely reflects intrinsic deficits in the mutant myosin rather than a major shift in ratio of SRX and DRX conformations. Consistent with this model, fibers from MyBPC-RNAi–treated Myh6F764L/F764L mice had improved proportions of myosins in SRX and DRX compared to fibers from MyBPCt/t mice, but levels were not corrected to the physiologic proportions found in WT fibers (P = 0.004; fig. S6). We then asked whether an allosteric inhibitor of myosin ATPase, MYK-461, influenced the dynamic ratios of myosin heads in the SRX and DRX conformations. Skinned cardiac fibers from WT and cMyBPC-deficient mice treated with MYK-461 (0.3 μM) increased the proportion of myosins in SRX and reduced myosins in DRX by 60% in WT (P = 0.007), 65% in Mypbc3t/+ (P = 0.0002), and 70% in Mypbc3t/t fibers (P = 0.0001) in comparison to untreated corresponding genotypes (Fig. 3, A to C). This MYK-461 dose normalized cardiomyocyte contractility (Fig. 2, C and D), indicating a direct relationship between the proportion of myosins in DRX and cellular hypercontractility and relaxation rates. Analyses of Mant-ATP release from skinned human HCM heart fibers with heterozygous MYBPC3 truncations showed abnormalities comparable to those in mutant mouse hearts. The proportion of myosin in DRX was increased (~50%, P = 0.006) compared to normal human heart fibers (Fig. 3, D and F). Moreover, treatment of MYBPC3-mutant fibers with 0.3 μM MYK-461 normalized the ratio of DRX:SRX by reciprocally reducing the proportion in DRX and increasing the proportion in SRX by 40% (P = 0.003, versus untreated). Together, these observations indicate that MYBPC3 mutations in humans and mice disrupted normal myosin conformations, resulting in the increased contractility, diminished relaxation, and excessive ATP consumption—prototypic findings in HCM. These abnormalities can be pharmacologically corrected with the myosin ATPase inhibitor, MYK-461.

DISCUSSION Truncating germline mutations in one or both Mybpc3 alleles and RNAi silencing of Mybpc3 transcripts caused comparable abnormalities in cardiomyocyte contraction and relaxation, thereby supporting the conclusion that MYBPC3 mutations cause HCM by haploinsufficiency. We show that the severity of cardiomyocyte phenotypes is dependent on cMyBPC quantities; truncation of one allele that reduced protein expression without altering cMyBPC phosphorylation produced milder abnormalities in systolic and diastolic performance than biallelic mutations. The phenotypes of MyBPC-RNAi cardiomyocytes confirmed this dose-dependent relationship; extinguishing postnatal protein expression substantially amplified hypercontractility and impaired relaxation, evidencing a direct role of cMyBPC across the cardiac cycle. Our studies uncovered a dichotomous reduction of systolic contraction in Mypbc3t/t mice despite prominent hypercontractility in isolated cardiomyocytes from Mypbc3t/t or MyBPC-RNAi mice. Several factors may account for these observations. The hypercontractility because of profoundly depressed cMyBPC quantities in Mypbc3t/t mice may evoke life-long compensatory mechanisms such as reduced phosphorylation of regulatory light chains that could normalize sarcomere performance (21, 37–40). Compensatory effects in other model systems with germline mutations have similarly mitigated the expected phenotype (41). Second, the Mypbc3t/t cardiomyocytes with impaired post-natal cytokinesis that increases numbers of cardiomyocytes, particularly mononuclear cardiomyocytes (27), may be dysfunctional. Increased numbers of fragile mononuclear cardiomyocytes, and hypercontractility of binuclear cells, and altered myocardial geometry may each increase energy demands, which when unmet, could compromise contraction, promote cardiomyocyte death and fibrosis, and thereby diminish the performance of Mypbc3t/t hearts. By eliminating these factors, our analyses of isolated Mypbc3t/t and post-natal MyBPC-RNAi cardiomyocytes provided a more proximal readout, revealing that reduced cMyBPC increased sarcomere contractility. Through genetic and pharmacological approaches, we demonstrated that myosin dysregulation provides a unifying mechanism by which thick filament gene mutations in MYH7 and MYPBC3 cause HCM (Fig. 4). Genetic repression of myosin’s motor function, as occurs in Myh6F764L/F764L cardiomyocytes (34), improved the hypercontractile phenotype of cMyBPC deficiency, a finding that centrally positions myosin dysregulation in the pathogenicity of cMyBPC mutations. Conversely, depletion of cMyBPC protein activated sarcomere performance, similar to the effects associated with phosphorylation or the regulatory light chain (21, 37–40). These observations indicate that strategies to reduce cMyBPC can, at least transiently, increase myosin contractility. Fig. 4 Schematic of the mechanism by which haploinsufficiency of cMyBPC causes HCM. (A) Schematic of a WT sarcomere (left side) with normal cMyBPC quantities and physiologic contractility and relaxation due to appropriate proportions of myosins in state of SRX with low-energy consumption or DRX with high-energy consumption. An HCM sarcomere with reduced cMyBPC quantities that dysregulates the proportions of myosins in DRX (increased) and SRX (reduced) is shown on the right. The increased proportion of DRX myosins causes inappropriate sarcomere hypercontractility. Yellow denotes the approximate interaction site of MYK-461 on myosin, which abates the hypercontractile phenotype and shifts the myosin DRX:SRX equilibrium back toward normal. (B) Contractile waveform of an individual cardiomyocyte isolated from a healthy human showing normal sarcomere shortening and normal relaxation duration. (C) Contractile waveform from a cardiomyocyte isolated from a patient with HCM with cMyBPC haploinsufficiency showing hypercontractility with increased sarcomere shortening and slowed relaxation. When exposed to MYK-461, the HCM phenotypes of hypercontractility are normalized by restoring physiologic balance of myosin DRX:SRX. Depletion of cMyBPC also slowed cardiac relaxation, as has been previously observed (42–45), an abnormality that precedes the development of hypertrophy in patients with HCM with heterozygous MYBPC3 mutations (46–49). Although we could not demonstrate significant relaxation deficits in Mypbc3t/+ cardiomyocytes, diastolic abnormalities were prominent in Mypbc3t/t and MyBPC-RNAi cardiomyocytes. Whereas Myh6764/764 reduced the increased duration of relaxation associated with cMyBPC deficiency, we demonstrated that pharmacologic treatment with MYK-461 more effectively improved relaxation. Mant-ATP experiments provided a mechanism for these observations. Our experiments and those by others (30, 31) show altered proportions of myosins in DRX and SRX in mouse and human myocardium with truncating cMyBPC mutations. Loss of cMyBPC increased the proportions of myosin in the more active DRX conformation. MYK-461 shifted DRX:SRX proportions in mice and human tissues at concentrations that alleviated enhanced cardiomyocyte contractility, data that strongly suggest that increases in the proportion of myosins in DRX contribute to the hypercontractile phenotype of HCM. MYK-461 was acutely administered to permeabilized myocardium in our study; therefore, its binding to myosin (and not secondary signaling events) is most likely the driver of these beneficial biophysical changes. MYK-461 represses filamentous function both by directly decreasing myosin contractile function and increasing relaxation properties, evidence supporting dual mechanisms for MYK-461 modulation of muscle function. We propose that shifting the DRX:SRX ratio to favor the normal SRX abundance reduces the pool of myosin heads available for strong cross-bridge formation and diminishes the abundance of active heads that must detach from actin to allow relaxation, thereby shortening the time for cardiomyocytes to restore resting SL. We demonstrated the therapeutic potential of targeting myosin in patients with MYBPC3-truncating mutations. As previously shown in HCM mice with myosin mutations (3), MYK-461 treatment of mouse or human heart tissues with cMyBPC mutations resulted in dose-dependent attenuation of hypercontractility. Because MyBPCt/+ mice do not exhibit in vivo morphologic or hemodynamic parameters of human HCM, these models can only confirm that MYK-461 is well tolerated. However, in combination with correction of cellular abnormalities in isolated mouse, MyBPCt/+ cardiomyocytes, and normalized rates of ATP cycling in human HCM hearts with MYBPC3 mutations, we expect that MYK-461 will also be effective in patients. We recognize several limitations in this study. Mypbc3t/+ mice do not recapitulate the extent of hypertrophy nor the degree of relaxation deficits found in patients with heterozygous MYBPC3 mutations, factors that diminish the value of longitudinal treatment trials in mice. Contractility measurements and relaxation assays were performed in an unloaded cardiomyocytes. Despite these limitations, we suggest that these preclinical data support the need for detailed studies in human patients with HCM. In summary, cMyBPC truncation causes HCM by a mechanism of haploinsufficiency, wherein myosin SRX conformations are destabilized, leading to deleterious ratios of DRX:SRX conformations that drive hypercontractility, impair relaxation, and increase energy consumption. This triad of abnormalities explains the clinical phenotypes of hyperdynamic contraction, diastolic dysfunction, and energy deprivation observed in HCM hearts. In addition, these observations support the conclusion that myosin dysregulation is a central mechanism of HCM pathophysiology in cMyBPC haploinsufficiency and substantially contributes to diastolic dysfunction. By targeting myosin functions genetically or pharmacologically, these phenotypes are normalized in cardiomyocytes and will likely reduce disease pathogenesis in vivo. Demonstrating that myosin is a central player in the pathophysiology of cMyBPC truncation may extend the therapeutic utility of MYK-461 to the proportionally largest subset of patients with HCM, those with MYBPC3 mutations.

MATERIALS AND METHODS Study design We hypothesized that cardiac contractile abnormalities observed in patients with MYBPC3 mutations that truncate the cMyBPC protein could be modeled at the cellular level. To test this, we studied mouse cardiomyocytes (from n > 3 randomized animals per genotype and treatment group), interrogating cells with heterozygous mutations, and cells depleted of cMyBPC. No data were excluded, and all analyses were performed under blinded conditions. Cardiomyocyte contractility was sampled until 80% of initial contractility was observed, at which point analysis was halted. Cellular replicates were performed (n > 10 per experiment), and experiments were performed at least thrice for all samples. After establishing a contractile phenotype in cardiomyocytes with decreased cMyBPC protein, we hypothesized that myosin activities were involved. We tested this model by studying cardiomyocytes with a damaging mutation in myosin and with a myosin allosteric inhibitor MYK-461. Pharmacologic perturbations were performed to test the dose dependence of MYK-461 on contractile function. On the basis of experimental observations, we hypothesized that depletion of cMyBPC caused contractile phenotypes by disturbing the balance of relaxed conformational states of myosin DRX:SRX and that MYK-461 corrected this imbalance. We extended this model by comparing the proportions of each confirmation in human heart tissue from patients with heterozygous mutations that truncate cMyBPC (n = 3) and donor tissues (n = 2) without MYBPC3 mutations at baseline and after treatment with MYK-461. cMyBPC truncation All animal protocols were compliant with the approved protocols of the Association for the Assessment and Accreditation of Laboratory Animal Care and Harvard Medical School. Mypbc3t/+, Mypbc3t/t, and WT mice were studied (129SvEv background), with histopathology being previously described in detail (25, 32). The truncated Mybpc3 alleles were created by inserting the neomycin resistance gene, expressed under the phosphoglycerate kinase I promoter, into exon 30, creating a predicted truncation at amino acid 1064 of the 1270 residues of cMyBPC. Homozygous mice express ~10% of the amount of cMyBPC protein in WT myofibrillar extracts (32). It should be noted that a greater proportion of cells in the Mypbc3t/t cohort exhibited fibrillation and were excluded from contractile measures because they could not be reliably paced, making measures in Mypbc3t/t cardiomyocytes more challenging. Mypbc3t/+ and WT mice were administered MYK-461 (2.5 mg/kg per day via drinking water), as described (3). Echocardiograms of mice were obtained at baseline (age, 5 weeks) and every 5 to 20 weeks of age. RNA interference RNAi was delivered at P10 by AAV vector using AAV9 capsid packaging by triple transfection, as described (27). The AAV9 vector contains a short hairpin RNA construct that specifically targets specific 21–base pair sequence targeted to Mybpc3 exon 19 and an enhanced GFP plasmid (Addgene). Vector (5 × 1013 vg/kg) was injected into the thoracic cavity of Mybcp3t/+ neonates (age P1) at day 1 of life, and GFP fluorescence was used to identify isolated cardiomyocytes that had taken up the vector. GFP fluorescence was evident from 48 hours after injection for at least 5 months. RNAi reduced cMyBPC expression to ~10% of normal (27). Human myectomy samples Human myectomy samples were obtained after written informed consent from three patients with HCM with distinct heterozygous frameshift truncating variants (Gln981fs, Leu1014fs, and Lys1209fs) in MYBPC3. Myectomy samples of the septum were flash-frozen and stored in liquid nitrogen, and tissue preparation for Mant-ATP experiments were performed as described below. Cardiomyocyte isolation Cardiomyocytes were isolated from 8- to 20-week old mice by rapid explantation and aortic cannulation on a Langendorff apparatus for perfusion with enzyme buffer [composition: 135 mM NaCl, 4 mM KCl, 0.33 mM NaH 2 PO 4 , 1 mM MgCl 2 , and 10 mM Hepes (pH 7.40), which incorporated collagenase D, collagenase B, and protease XIV] for 10 min. After perfusion, the atria and right ventricle were removed, and the left ventricle was minced in modified Tyrode buffer [composition: 135 mM NaCl, 4 mM KCl, 0.33 mM NaH 2 PO 4 , 1 mM MgCl 2 , and 10 mM Hepes (pH 7.40), which incorporated bovine serum albumin] and passed through a 100-μm filter into a 50-ml conical tube. Tissue was settled for 15 min to allow myocytes to pellet by gravity. The pellet was then sequentially resuspended every 10 min through an increasing calcium gradient (5, 20, 50, and 100% of calcium tyrode) to provide a cell fraction enriched in myocytes with a final experimental buffer (EB; composition: 137 mM NaCl, 5.4 mM KCl, 1.2 mM CaCl 2 , 0.5 mM MgCl 2 , and 10 mM Hepes (pH 7.40), which incorporated glucose). Contractile measures of myocyte function Isolated cardiomyocytes were placed in wells of a sixwell plate that had been precoated with laminin. Laminin coating was performed for 2 hours before cardiomyocyte introduction at a concentration of 10 μg/ml in phosphate-buffered saline (PBS) [composition: 1 mM KH 2 PO 4 , 155 mM NaCl, and 3 mM Na 2 HPO 4 (pH 7.40)]. Laminin coating solution was washed once with PBS before cells were introduced into the wells. Once cells were introduced, they were left to incubate for 10 min to equilibrate to experimental temperature (30°C). Cells were imaged using a Keyence BZ-X710 microscope using a Nikon 40×/0.65 numerical aperture (NA) objective. Cells were kept at 30°C using microscope specific incubation chamber that was also used to deliver 20% O 2 and 5% CO 2 to the experimental chamber. Cells were paced at 1 Hz using custom-built electrodes hooked up to a pacing unit (Pulsar 6i, FHC) delivering 20 V. Movies were acquired at 29 frames/s for 5 s (five contractile cycles). An ImageJ plugin SarCoptiM was used to track SLs during contractile cycles (50). Sarcomere tracking was then used to calculate cellular shortening (%), relaxed and contracted SLs (μm), contractile cycle, and relaxation durations (seconds). For experiments incorporating MYK-461, drug was applied in concentrations ranging from 0.05 to 0.3 μM in the EB. MYK-461 was incubated with cells for a minimum of 10 min before data acquisition. Mant-ATP experiments Mice were sacrificed by rapid cervical dislocation, atria and right ventricle were removed, and samples were flash-frozen in liquid nitrogen. Mant-ATP protocols were adapted from publications (30, 36). LV human or mouse myocardial samples (20 mg) were thawed in permeabilization buffer [composition: 100 mM NaCl, 8 mM MgCl 2 ; 5 mM EGTA, 5 mM K 2 HPO 4 , 5 mM KH 2 PO 4 , 3 mM NaN 3 , 5 mM ATP, 1 mM dithiothreitol (DTT), 20 mM 2,3-butanedione monoxime (BDM), and 0.1% Triton X-100 (pH 7.0)]. Samples were permeabilized for 6 hours on ice on a rocker solution changes occurring every 2 hours. At the completion of this step, samples were stored overnight at −20°C in glycerinating solution [composition: 120 mM K acetate, 5 mM Mg acetate, 2.5 mM K 2 HPO 4 , 2.5 mM KH 2 PO 4 , 50 mM Mops; 5 mM ATP, 20 mM BDM, 2 mM DTT, and 50% (v/v) glycerol (pH 6.8)] for dissection within 2 days. Once glycerinated ventricular myocardium was dissected into ~90-μm by 400-μm pieces that were held under two pins in a chamber constructed from a slide and coverslip, these samples were permeabilized using the permeabilization buffer for a further 30 min on ice before experimentation. After secondary permeabilization, chambers were flushed with glycerinating buffer. For fluorescence acquisition, a Nikon TE2000-E was used with a Nikon 20×/0.45 NA objective using a Hamamatsu C9100 electron multiplying charge-coupled device. Frames were acquired every 10 s with a 20-ms acquisition and exposure time using a 4′,6-diamidino-2-phenylindole filter set, and images were collected for 15 min. Before acquisition, each chamber was flushed with ATP buffer [composition: 120 mM K acetate, 5 mM Mg acetate, 2.5 mM K 2 HPO 4 , 2.5 mM KH 2 PO 4 , 4 mM ATP, 50 mM Mops, and 2 mM DTT (pH 6.8)] to remove glycerol. This buffer was replaced with two-chamber volumes of rigor buffer [composition: 120 mM K acetate, 5 mM Mg acetate, 2.5 mM K 2 HPO 4 , 2.5 mM KH 2 PO 4 , 50 mM Mops, and 2 mM DTT (pH 6.8)]. Rigor buffer was incubated for 5 min to allow rigor to set in. Initial fluorescence acquisition was simultaneous with the addition of one-chamber volume of rigor buffer with 250 μm Mant-ATP to visualize fluorescent Mant-ATP wash in. At the end of a 15-min acquisition, a chamber volume of ATP buffer (rigor buffer and 4 mM ATP) was added to the chamber with simultaneous acquisition of the Mant-ATP chase. For experiments with MYK-461, all experimental solutions contained 0.3 μM MYK-461. Mant-ATP analyses Similar to protocols previously described for analysis (30), three regions of each myocardial tissue strip were sampled for fluorescence decay using the region of interest manager in ImageJ. The final data point of fluorescence wash in defined the y intercept. Subtraction of nonmyosin-bound Mant-ATP fluorescence signal was made using a correction factor of 52%, as indicated previously (36). All data were plot as a normalized intensity of initial fluorescent intensity from the three sampled regions. These data were fit to an unconstrained double exponential decay using SigmaPlot Where A1 is the amplitude of the initial rapid decay approximating the DRX, with T1 as the time constant for this decay. A2 is the slower second decay approximating the proportion of myosin heads in the SRX, with its associated time constant T2. Each individual experiment was fit using this double exponential decay, with all values determined and plot. Statistical analysis was performed using two-way ANOVA with multiple comparisons tests. Statistical analysis When comparing two treatment groups, Student’s t tests were used. In instances where experimental hypothesis was tested among multiple treatment groups, one-way ANOVA was used. For multiple comparisons, post hoc Bonferroni corrections were used with a significance cutoff of P < 0.05.

SUPPLEMENTARY MATERIALS www.sciencetranslationalmedicine.org/cgi/content/full/11/476/eaat1199/DC1 Materials and Methods Fig. S1. Representation of Mybpc3 mouse models. Fig. S2. In vivo cardiac function and proteomic characterization in Mybpc3 mouse models. Fig. S3. Resting SLs of animal models studied. Fig. S4. The protein and function effects of increasing MyBPC-RNAi titers. Fig. S5. Analysis of Mant-ATP video files. Fig. S6. Mant-ATP assays in cardiac tissues for Myh6764/764 mice with DCM and Myh6764/764 + Mybpc3 RNAi. Table S1. Echocardiographic parameters with MYK-461 administration. Movie S1. The fluorescent decay during dark ATP chase of Mant-ATP in permeabilized myocardium. Reference (51)

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Acknowledgments: We thank R. Cooke for assistance with the Mant-ATP assays. Funding: This work was supported in part by grants from a Sir Henry Wellcome Postdoctoral Fellowship from the Wellcome Trust (206466/Z/17/Z to C.N.T.), the Sarnoff Foundation (to A.C.G.), National Medical Research Council (NMRC) and Ministry of Education, Singapore to D.L. and J.J., American Heart Association Midwest Predoctoral Fellowship (15PRE22430028 to T.L.L.), American Heart Association Postdoctoral Fellowship (17POST33630095 to J.W.M.), National Institutes of Health grants (R01HL130356, R01HL105826, and K02HL114749 to S.S.), and Research council of Norway (221707 to I.G.L.), the British Heart Foundation (program grant RG/12/16/29939) and the British Heart Foundation Centre of Research Excellence (Oxford) to H.W. and C.S.R, the National Heart Blood and Lung Institutes and Leducq Foundation (HL084553 and HL080494 to J.G.S. and C.E.S.), and the Howard Hughes Medical Institute (to C.E.S.). Author contributions: C.N.T. and H.W. performed experiments on mouse cardiomyocytes. H.W. performed in vivo characterization of mice and RNAi injection. D.L. and J.J. produced the RNAi virus. A.C.T. and J.M.G. performed quantification of Mybpc3 expression. T.L.L., J.W.M., and S.S. performed phosphorylation analyses. H.W. and M.L. performed mouse genotyping. B.M. performed the preparation of human myectomy samples. C.N.T. and A.C.G. performed Mant-ATP experiments. Blots of cMyBPC were performed by H.W., T.L.L., J.W.M., and S.S. C.N.T., H.W., J.J., I.G.L., H.C.W., C.S.R., C.E.S., and J.G.S contributed to the intellectual design of experiments. All authors participated in the editing and preparation of the manuscript. Competing interests: C.E.S. and J.G.S. are founders and own shares in Myokardia Inc., a startup company that is developing therapeutics that target the sarcomere. All other authors declare that they have no competing interest. Data and materials availability: All data associated with this study are present in the paper or the Supplementary Materials.