RLC phosphorylation amplifies Ca²⁺ sensitivity of force in myocardium from cMyBP-C knockout mice

Hypertrophic cardiomyopathy (HCM) is a chronic heart disease characterized by decreased diastolic function and left ventricle hypertrophy. HCM affects 1 in every 200 people and is the most common cause of sudden cardiac death in athletes and young adults (Virani et al., 2020, Centers for Disease Control and Prevention, 2022). Clinical treatments for patients with HCM remain limited and focus on decreasing heart stress rather than treating the underlying genetic condition (Maron, 2010; Semsarian et al., 2015). The sarcomere, the fundamental structural unit underlying cardiac muscle contraction, contains thick filaments of myosin and thin filaments of actin. Along these filaments, actin–myosin crossbridge interactions generate the force and shortening to power cardiac contraction and blood circulation. It is well known that Ca²⁺ binding to the thin-filament regulatory proteins enable crossbridge binding. The mechanisms by which thick-filament regulatory proteins modulate crossbridge function remain less clear.

Genetic mutations in sarcomeric proteins can often be a preceding factor in HCM development (Marian, 2021). One of the most frequently mutated proteins associated with HCM clinical cases is cardiac myosin-binding protein-C (cMyBP-C). cMyBP-C is a thick-filament regulatory protein located in the c-zone of the cardiac sarcomere. cMyBP-C plays an important role in regulating contractility via interacting with the thick and thin filament with its N-terminal domain, and its C-terminus binds to the thick filament every three myosin crowns (Stelzer et al., 2006a; Stelzer et al., 2006b; Bhuiyan et al., 2016; Mun et al., 2014). Myosin regulatory light chain (RLC) is another thick-filament regulatory protein. RLC binds to the neck of the myosin-heavy chain, just between the catalytic “head” domain and the coiled-coil “tail region.” RLC undergoes posttranslational phosphorylation, which influences myosin head conformation and increases the probability of myosin binding to actin (Muthu et al., 2012; Toepfer et al., 2013; Breithaupt et al., 2019). Myosin heads that are positioned favorably to bind actin occupy the ON conformation, while folded myosin heads that cannot bind actin (i.e., the interacting heads motif) occupy the OFF conformation. Although the individual effects of these thick-filament regulatory proteins on contractile dynamics are starting to become more clear, it is unknown how cMyBP-C and RLC interact with each other to regulate cardiac function.

The cMyBP-C knockout mouse has been a useful transgenic model for characterizing the effects of cMyBP-C on cardiac contractility (Harris et al., 2002). Herein, we measured Ca²⁺-activated force production and myosin crossbridge activity in permeabilized (skinned) myocardial strips as RLC phosphorylation was modulated in the presence or absence of cMyBP-C. We employed a series of biochemical and biophysical assays to further understanding of regulatory interactions between cMyBP-C and RLC, and how these interactions may affect contractile function in cMyBP-C knockout and WT mice.

All procedures were approved by the Institutional Animal Care and Use Committee at the University of Wisconsin-Madison and complied with the National Institutes of Health Guide for the Use and Care of Laboratory Animals. cMyBP-C KO animals were produced by the laboratory of Dr. Richard L. Moss as previously described (Harris et al., 2002). Following euthanasia, hearts were excised, cut along the long axis from base to apex, splayed open, flash-frozen in liquid nitrogen, and stored at the University of Wisconsin-Madison. Hearts were shipped overnight on dry ice to the University of Idaho and stored at −80°C until transported to Washington State University.

Hearts were thawed, dissected, submerged in embedding medium (O.C.T. Compound, 23-730-57; Thermo Fisher Scientific), and flash-frozen in isopentane at −140°C for 30 s and stored at −80°C until sectioned in a cryostat (model CM1860; Leica). Samples were sectioned at 10 µm thickness, placed on slides, and stored at −20°C until stained. Slides were fixed with 4% paraformaldehyde, stained with hematoxylin and eosin as previously described (Fenwick et al., 2019), and covered with glass coverslips. Images were acquired using LAS X imagine software, Leica DFC295 camera, and Nikon Eclipse E200 microscope at 0.4 numerical aperture and 200× total magnification. Image processing was performed using ImageJ.

Cardiac fibrosis was assessed using a hydroxyproline colorimetric assay as previously described (Yuan et al., 2017). Briefly, 15 mg of left ventricle wall from each heart was placed in a glass vial containing 200 µl of 6 M HCl and incubated at 110°C overnight. In triplicate, 5 µl aliquots of the resulting solution were added to 80 µl of isopropanol, then mixed with 40 µl of a solution containing a 1:4 ratio of 7% w/v chloramine-T and acetate citrate buffer (0.695 M sodium acetate, 0.174 M citric acid, 0.435 M NaOH, and 38.5% isopropanol), and allowed to incubate at room temperature for 5 min. After incubation, 0.5 ml of the solution containing a 3:13 ratio of Ehrlich’s solution and isopropanol was added to each sample and incubated at 55°C for 30 min. Samples were placed on ice to quench the reaction, and 200 µl of each sample was aliquoted to a 96-well plate. Absorbance was measured at 558 nm.

Solution recipes were calculated and prepared as follows (Godt and Lindley, 1982), and all values are in mM where not otherwise specified. Relaxing solution: pCa 8.0 (pCa = −log₁₀[Ca²⁺]), 5 EGTA, 5 MgATP, 1 Mg²⁺, 0.3 Pi, 35 phosphocreatine, 300 U/ml creatine kinase (CK), 200 ionic strength, 3% Dextran T-500 (w/v), and pH 7.0. Activating solution was the same as the relaxing solution, but adjusted to a free [Ca²⁺] of pCa 4.8. Dissecting solution: 50 BES, 30.83 K propionate, 10 Na azide, 20 EGTA, 6.29 MgCl₂, 6.09 ATP, 1 DTT, 20 BDM, 50 Leupeptin, 275 Pefabloc, and 1 E-64. Permeabilizing (skinning) solution: Dissecting solution with 1% Triton X-100 wt/vol and 50% glycerol wt/vol. Storage solution: Dissecting solution with 50% glycerol wt/vol. Alkaline phosphatase (AP) solution: relaxing solution with 170 U/ml AP (sc-3716; ChemCruz). Myosin light chain kinase (MLCK) solution: relaxing solution, but adjusted to pCa 6.25, 1.1 µM human skeletal MLCK, and 12 µM Xenopus calmodulin. MLCK sham solution: relaxing solution, but adjusted to pCa 6.25, 12 µM Xenopus calmodulin.

Frozen hearts were placed in a vial with dissecting solution and left on ice for ∼10 min to thaw. Hearts were transferred onto a dissecting dish and papillary muscles were excised from the left ventricle. Papillary muscles were transferred to a dish containing skinning solution, trimmed, and skinned overnight at 4°C. Skinned papillary muscle strips were transferred to a storage solution, and if not used immediately for mechanics experiments, strips were stored at −20°C for up to 1 wk.

For all mechanics experiments, permeabilized left ventricular papillary muscles were trimmed to ∼180 μm in diameter and ∼700 μm in length. All strips were pretreated in AP solution for 10 min at room temperature prior to mounting. Aluminum T-clips were attached to the end of each strip and then mounted between a piezoelectric motor (P841.40; Physik Instrumente) and a strain gauge (AE801; Kronex), lowered into a 30 μl droplet of relaxing solution maintained at 28°C, and stretched to 2.2 μm sarcomere length measured by digital Fourier Transform (IonOptix Corp). Next, the strips were either incubated in MLCK solution or MLCK sham solution for 60 min, then relaxed to pCa 8.0, and the sarcomere length was checked again to confirm 2.2 μm. Following this, strips were Ca²⁺-activated from pCa 8.0 to pCa 4.8. At least three strips from each heart underwent the same mechanics protocol at each condition, with the total number of strips and hearts listed in Table 1.

Sinusoidal length perturbations of 0.125% myocardial strip length (clip-to-clip) were applied at 44 discreet frequencies from 0.8 to 250 Hz to measure the complex modulus as a function of frequency (Kawai and Brandt, 1980; Mulieri et al., 2002; Palmer et al., 2007). The complex modulus represents viscoelastic myocardial stiffness, which arises from the change in stress divided by the change in muscle length that is in-phase (elastic modulus) and out-of-phase (viscous modulus) with the oscillatory length change at each frequency.

Characteristics of the elastic and viscous moduli responses over the measured frequency range provide a signature of crossbridge binding and cycling kinetics. Shifts in the elastic and viscous moduli are useful for assessing changes in the number of bound crossbridges between experimental conditions. In addition, shifts in the viscous modulus are useful for assessing changes in the work-producing and work-absorbing characteristics of the myocardium that arise from force-generating, cycling crossbridges. Frequencies producing negative viscous moduli represent regions of work-producing muscle function, with the “dip frequency” or frequency of the minimum viscous modulus describing force-generating events and the crossbridge recruitment rate (Awinda et al., 2020; Mulieri et al., 2002; Campbell et al., 2004). Frequencies producing positive viscous moduli represent regions of work-absorbing muscle function, with the “peak frequency” or frequency of the maximum viscous modulus describing crossbridge distortion events and crossbridge detachment rate (Awinda et al., 2020; Mulieri et al., 2002; Palmer et al., 2007, Palmer et al., 2011). These characteristic regions of minima and maxima in the viscous modulus versus frequency relationship were used to assess the effects of MLCK treatment on crossbridge kinetics under maximal Ca²⁺-activated conditions. Given that viscous moduli were only measured at discrete frequencies, polynomials were fitted to regions of minimal viscous modulus (using 0.8–15-Hz data) and maximal viscous modulus (using 16–100-Hz data) using MATLAB (v9.11.0, The Mathworks) to create fitted curves at 0.05-Hz resolution. From these interpolated curves, we extracted the frequency of minimum viscous modulus and frequency of maximum viscous modulus.

After mechanical experiments were performed, permeabilized cardiac strips were homogenized, boiled for 5 min, chilled, and then stored at −80°C until gel electrophoresis. 12% acrylamide SDS-PAGE gels (BioRAD) were stained with Pro-Q diamond phosphostain and Sypro-Ruby total protein stain (Invitrogen). The intensity ratio of Pro-Q diamond to Sypro-Ruby was used to assess RLC phosphorylation levels. Average RLC phosphorylation was 10.3 ± 2.3% in AP-treated strips and 68.3 ± 2.4% in MLCK-treated strips, with no difference in phosphorylation levels between WT and cMyBP-C KO groups.

All statistical analyses were performed using SPSS (IBM Statistics). Unpaired Student’s t test was performed to assess differences of hydroxyproline levels between genotypes. Nested linear mixed models incorporating one effect (treatment) were utilized to compare viscous and elastic moduli at each frequency for pCa 8.0 and 4.8 data, within each genotype. Nested linear mixed models incorporating two main effects (treatment and genotype) and their interaction were utilized for the stress–pCa fit parameters from three-parameter Hill fits, and for the minimum or maximum frequency parameters extracted from the polynomial curve that fits to the viscous moduli. Nested linear mixed model analyses link data from the same hearts, with hearts being a random effect, to optimize statistical power. Post-hoc analyses were performed using Fisher’s least significant difference test, where P values < 0.05 were considered significant.

Histological samples were stained with hematoxylin to visualize the cell nuclei and stained with eosin to visualize the cytoplasm of the cell. Representative images show increased cellular disorganization and fibrotic infiltration in the cMyBP-C KO myocardial samples (Fig. 1, A and B), as previously reported (Harris et al., 2002). Hydroxyproline assays were utilized to measure relative collagen content, or fibrosis levels, in myocardial tissue samples. Hydroxyproline is the most abundant amino acid in collagen, which represents the most abundant fibrotic component of the extracellular matrix. Hydroxyproline concentration was greater for cMyBP-C KO vs. WT samples, indicating increased fibrosis in cMyBP-C KO hearts (Fig. 1 C). Increased extracellular fibrosis due to the loss of cMyBP-C sarcomere contributes to myocardial remodeling that may lead to further contractile deficits.

Isometric stress increased as activating [Ca²⁺] increased from pCa 8.0 to pCa 4.8, and these stress–pCa relationships were fit to a three-parameter Hill equation (Table 1). Maximal Ca²⁺-activated stress (i.e., the increase in stress from the relaxed value at pCa 8.0) values were plotted against pCa for myocardial strips from WT and cMyBP-C KO mice. All strips were pretreated with AP, followed by a sham incubation or an MLCK incubation to phosphorylate RLC. AP treatment decreased the average phosphorylation to 10 ± 1% and MLCK treatment increased the average phosphorylation to 68 ± 3% (Fig. 2 C). There were no differences in the average RLC phosphorylation levels between genotypes. MLCK treatment increased Ca²⁺ sensitivity of contraction (as measured by pCa₅₀) and maximal Ca²⁺-activated stress in the WT strips (Fig. 2 A), consistent with our prior findings using WT animals (Pulcastro et al., 2016). In the cMyBP-C KO strips, MLCK treatment increased Ca²⁺ sensitivity but not maximal Ca²⁺-activated stress (Fig. 2 B and Fig. 3 B).

Normalizing stress–pCa curves of both genotypes to maximal stress within each condition helps show the differences in Ca²⁺ sensitivity between groups (Fig. 3 A). As introduced above, MLCK treatment increased Ca²⁺ sensitivity in both the WT and cMyBP-C KO strips, compared with their AP-treated controls (Fig. 3 C). However, pCa₅₀ was most elevated in the MLCK-treated strips from cMyBP-C KO mice, indicating that MLCK treatment has an even greater effect on Ca²⁺ sensitivity when cMyBP-C is absent from the sarcomere. This suggests an interaction and/or coordination between RLC and cMyBP-C that regulates steady-state, isometric force production.

Sinusoidal length–perturbation analysis was used to measure myocardial viscoelasticity and the effect of MLCK treatment on crossbridge cycling rates at both relaxed (pCa 8.0) and maximally activated conditions (pCa 4.8). Under relaxed conditions, there were no significant differences for the viscous or elastic moduli at any single frequency as a result of treatment or genotype (Fig. 4). For both WT and cMyBP-C KO strips at maximal Ca²⁺ activation, elastic moduli values were greater with MLCK treatment compared with AP-treated strips within each genotype (Fig. 5, A and B). Specifically, the elastic moduli values were larger at oscillatory frequencies >16 Hz for WT strips and frequencies >1.5 Hz for cMyBP-C KO strips. Similar to the elastic moduli differences, viscous moduli were greater for MLCK-treated strips than AP-treated strips at frequencies >65 Hz for WT and between 20 and 25 Hz cMyBP-C KO (Fig. 5, C and D). In combination, this reflects more bound, cycling crossbridges or increased crossbridge stiffness in the MLCK-treated strips for both genotypes compared with AP-treated strips.

Frequency shifts in the viscoelastic system response result from changes in enzymatic crossbridge cycling kinetics during Ca²⁺-activated conditions. The frequency of the minimum viscous modulus (i.e., the dip frequency for a single strip) was not different between WT and cMyBP-C strips, nor for MLCK-treated vs. AP-treated strips from cMyBP-C KO mice (Fig. 5 E). Albeit small, the dip frequency was slightly higher for MLCK-treated WT strips than AP-treated WT strips (Fig. 5, C and E), indicating a slightly faster crossbridge attachment rate for the MLCK-treated WT strips. We recognize that the slight increase in the dip frequency for MLCK-treated WT strips appears nearly equivalent to the average value of both treatment groups for cMyBP-C KO, although these latter two groups are not different from AP-treated WT strips (Fig. 5 E). A portion of this may follow from the different data distributions among all four groups and our nested linear mixed model analysis that clusters the technical replicates from each heart to elucidate biological differences more clearly than non-nested tests. With four to seven preparations of each treatment from each heart, the subtle statistical differences can be attributed to an increase in technical variation rather than a false biological difference. Nonetheless, any differences in crossbridge recruitment rate among the four groups are rather small.

Compared with WT strips, the frequency of the maximum viscous modulus (i.e., the peak frequency for a single strip) decreased for cMyBP-C KO strips, which indicates a slower crossbridge detachment rate in the absence of cMyBP-C (Fig. 5 F). Upon MLCK treatment, the peak frequency shifted toward a lower value for both genotypes (Fig. 5, C, D, and F), further suggesting that MLCK treatment slows crossbridge detachment (Pulcastro et al., 2016). Thus, the genotype and the treatment compound to confer the slowest detachment for the MLCK-treated cMyBP-C KO strips. In combination, these kinetics data suggest that removing cMyBP-C from the cardiac sarcomere may slow crossbridge detachment and the MLCK treatment slows the crossbridge detachment rate for WT and cMyBP-C strips.

In this study, we performed muscle mechanics measurements to investigate the effect of RLC phosphorylation in permeabilized myocardial strips from WT and cMyBP-C KO mice. This enabled a relative assessment of the impact of RLC phosphorylation on the isometric stress–pCa relationship, myocardial viscoelasticity, and crossbridge cycling kinetics when cMyBP-C was either present or absent from the sarcomere. We found that cMyBP-C loss contributed to cellular modifications, such as increased fibrosis and cellular disorganization. RLC phosphorylation increased maximal force in the WT preparations, but not the cMyBP-C KO preparations. This difference could follow from increased fibrosis in the KO preparations, thereby compromising maximal force. RLC phosphorylation increased submaximal force production in strips from both genotypes, although RLC phosphorylation increased Ca²⁺ sensitivity of contraction more in the cMyBP-C KO strips (by roughly twice as much). At maximal Ca²⁺ activation, RLC phosphorylation also increased myocardial stiffness (elastic modulus) for both genotypes, suggesting greater crossbridge binding or increased crossbridge stiffness with phosphorylated RLC. The increased stiffness was not obviously driven by differences in myofilament stiffness because myocardial viscoelasticity of relaxed strips (pCa 8.0) did not differ with genotype or RLC phosphorylation. RLC phosphorylation slowed crossbridge kinetics in WT and cMyBP-C KO strips, although crossbridge kinetics were slightly faster in WT strips. Thus, RLC phosphorylation may prolong the duration of myosin attachment to augment crossbridge contributions to thin-filament activation, and this mechanism was amplified when cMyBP-C was removed (or absent) from the sarcomere. Altogether these findings indicate that RLC and cMyBP-C likely interact with each other, acting in concert to regulate cardiac muscle contraction.

cMyBP-C phosphorylation throughout the cardiac sarcomere defines interactions between the N-terminus of cMyBP-C and the thin or thick filament. Native phosphorylation levels of cMyBP-C have been reported to exist at 50–60% (Previs et al., 2016; McNamara et al., 2019), and phosphorylation increases during β-adrenergic stimulation via the cAMP-dependent enzyme, protein kinase A (Sadayappan et al., 2009). We found that cMyBP-C phosphorylation levels in our WT papillary preparations were 42 ± 0.2% (SDS-PAGE with SYPRO Ruby and Pro-Q Diamond stains, data not shown) in our cardiac tissue samples from the WT mice. cMyBP-C phosphorylation levels did not differ between AP and MLCK treatment, indicating that cMyBP-C phosphorylation is not affected by AP or MLCK. The 42% cMyBP-C phosphorylation level would be consistent with nearly half of the cMyBP-C being released from a thin-filament interaction (as occurs with cMyBP-C phosphorylation) and potentially interacting with the myosin S2 region. RLC also binds the S2 region of myosin, which allows for potential interactions between the N-terminus of cMyBP-C and RLC (Ponnam and Kampourakis, 2022; Pfuhl and Gautel, 2012). This may create an important “regulatory nexus” between cMyBP-C and RLC, given the developments of dual-filament regulation over the past decade (Zhang et al., 2017; Campbell et al., 2018; Kosta and Dauby, 2021; Awinda et al., 2020).

Phosphorylation of RLC is an essential mechanism to regulate cardiac contractility. RLC phosphorylation influences the conformation of the myosin heads, shifting them toward the ON or active state, closer to actin binding sites (Kampourakis and Irving, 2015; Sheikh et al., 2012). This structural shift increases the time acto-myosin crossbridges are bound, the force of contraction, Ca²⁺ sensitivity of contraction, and myocardial stiffness (Pulcastro et al., 2016; Khromov et al., 1998). In vitro motility assays using skeletal muscle myosin have shown that RLC phosphorylation slowed load-dependent crossbridge kinetics, thereby prompting speculation that RLC phosphorylation may increase myosin stiffness (Greenberg et al., 2009). Computational modeling of cardiac contractility showed that increased myosin stiffness with RLC phosphorylation could enhance twitch force and ventricular torsion throughout a heartbeat (Sheikh et al., 2012). While we did not directly measure myosin stiffness, we did measure viscoelastic myocardial stiffness under relaxed and activated conditions (Figs. 4 and 5). As strips are Ca²⁺ activated, the increases in viscoelastic myocardial stiffness are attributed to force-generating myosin crossbridges (Fig. 5). More specifically, the elastic modulus values show a clear increase under maximally activated conditions upon MLCK treatment for both genotypes. The range of frequencies where RLC phosphorylation significantly increased these moduli values (Fig. 5, A and B) was greater for the cMyBP-C KO strips, which supports the idea that RLC phosphorylation may have a greater relative impact on increases in crossbridge binding or crossbridge stiffness. Increases in crossbridge stiffness may alter nucleotide handling rates and “power-stroke” distances to augment crossbridge binding and force production with RLC phosphorylation, and to an even greater degree when cMyBP-C is absent from the sarcomere (Greenberg et al., 2009; Tanner et al., 2014; Wang et al., 2014).

Colson et al. (2010) also showed that RLC phosphorylation moved myosin heads away from thick filaments, closer to thin filaments, and that RLC phosphorylation simultaneously decreased thick-to-thin filament spacing. Eliminating cMyBP-C from the sarcomere did not affect thick-to-thin filament spacing (Colson et al., 2007). Therefore, it is possible that the RLC phosphorylation-dependent reduction in lattice spacing (∼2%, at 2.15 µm sarcomere length; Colson et al., 2010) saturates or decreases at longer sarcomere lengths, such as the 2.2 µm sarcomere length utilized in this study. In addition, we utilized 3% Dextran T-500 to restore native thick-to-thin filament spacing that was disrupted during skinning (Kawai and Schulman, 1985; Kawai et al., 1993), which may overwhelm any subtle changes in lattice spacing due to RLC phosphorylation. To our knowledge, no previous studies have investigated the combined effects of increasing RLC phosphorylation and Dextran T-500 levels on lattice spacing per se. Nonetheless, it is likely that RLC-dependent decreases in thick-to-thin filament spacing and myosin head relocation toward the thin filament (away from the thick-filament backbone) could combine to increase the probability of crossbridge binding and decrease crossbridge detachment rate to augment force production.

We did not observe any clear increases in the crossbridge attachment rate (frequency of minimum viscous modulus; Fig. 5 E) with RLC phosphorylation or cMyBP-C loss, as previously reported (Stelzer et al., 2006a; Sweeney and Stull, 1990). Increases in crossbridge attachment rate have been associated with RLC phosphorylation repelling myosin heads away from the thick-filament backbone (Colson et al., 2010; Levine et al., 1998), which relocates them closer to their actin-binding sites along thin filaments. Alternatively, it is possible that RLC phosphorylation-dependent relocation of myosin heads could stabilize or promote strong crossbridge binding, thereby allowing crossbridges to remain bound longer and generate greater force (Tyska and Warshaw, 2002). The latter interpretation is consistent with the slower crossbridge detachment rates (frequency of maximum viscous modulus; Fig. 5 F) in the MLCK-treated strips from both genotypes.

Conflicting results have been reported regarding the role of cMyBP-C loss on crossbridge cycling kinetics, showing increases in crossbridge cycling rates (Harris et al., 2002; Korte et al., 2003; Stelzer et al., 2006b; Tong et al., 2008) and decreases in crossbridge cycling rates (Carrier et al., 2004; Pohlmann et al., 2007). A portion of the discrepancy may follow from the type of muscle preparation being studied. For example, de Lange et al. (2013) only observed increased crossbridge cycling kinetics in either single myocyte preparations derived from a cMyBP-C KO mouse model or human-derived engineered heart tissues with a cMyBP-C deletion. Neither of these preparations showed any hypertrophic remodeling and fibrosis. de Lang et al. (2013) did, however, observe decreased crossbridge cycling kinetics in multicellular preparations from cMyBP-C KO mice, in which hypertrophic remodeling and fibrosis were present. We also found slower detachment rates in our multicellular strips from cMyBP-C KO mice with demonstrated fibrosis (Fig. 1), consistent with prior studies using multicellular preparations with hypertrophic remodeling (Carrier et al., 2004; Pohlmann et al., 2007). It is well established that increased cardiac fibrosis results in relaxation deficits and dysregulated shortening velocities, likely due to increased myocardial stiffness (Burlew and Weber, 2002; Detterich, 2017; Moreo et al., 2009). Consequently, greater fibrosis may negatively impact muscle shortening and relaxation. The increased fibrosis observed in the multicellular preparations may alter how forces are transmitted throughout the myocardial strip and sarcomere, which may contribute to slowed crossbridge detachment rate (Fig. 5 F). A recent study by Hanft et al. (2021) showed slower maximal shortening velocity in permeabilized cardiomyocytes from cMyBP-C KO at long sarcomere lengths (compared with controls), but similar maximal shortening velocities at short sarcomere length . Another study showed faster unloaded shortening velocity and faster time-to-peak twitch, but slower twitch relaxation and slower crossbridge kinetics in permeabilized myocardial strips from a cMyBP-C truncation mouse model (Palmer et al., 2004). Similarly, a recent study observed faster time-dependent unloaded shortening velocities in permeabilized myocardial strips from a cMyBP-C KO mouse model during submaximal activation (Giles et al., 2021). We speculate that the preparation type (myocyte vs. multicellular), permeabilization (skinned vs. intact), disease phenotype (pre-remodeling vs. hypertrophic remodeling and fibrosis), sarcomere length (short vs. long), and load (unloaded vs. loaded shortening) may all independently contribute to the differing contractile kinetics reported among these studies. Future investigation into cMyBP-C’s regulatory role within the sarcomere will likely further elucidate its effects on each variable listed above in the context of contractile kinetics. RLC phosphorylation also led to decreased detachment rates, consistent with prior findings (Kampourakis and Irving, 2015; Pulcastro et al., 2016). In conjunction, both RLC phosphorylation and cMyBP-C loss contributed to slower detachment rates, and the combination of both effects may play a role in the increased Ca²⁺ sensitivity observed in cMyBP-C KO strips during RLC phosphorylation.

Altogether, our data show that (1) RLC phosphorylation enhances Ca²⁺ sensitivity of contraction in both genotypes, and (2) the effect of RLC phosphorylation on Ca²⁺ sensitivity is amplified when cMyBP-C is absent from the sarcomere. These findings suggest that RLC and cMyBP-C likely act in concert to contribute to contractility in healthy hearts. Prior studies have reported an interaction, or perhaps binding, between the C0 domain of cMyBP-C and RLC (Ratti et al., 2011; Bunch et al., 2018; Kampourakis and Irving, 2015; Pfuhl and Gautel, 2012; Ponnam et al., 2019), although the specific residues or number of RLC molecules with which cMyBP-C can interact are not well-defined. A plausible explanation for the amplified effects observed herein is the suppression of RLC activity by cMyBP-C’s N-terminal binding to RLC. If the binding of the N-terminus of cMyBP-C and RLC does occur, this may directly alter RLC structure and/or function to mimic activity similar to decreased phosphorylation of RLC without altering the capacity for RLC to be phosphorylated (as demonstrated by the equal ability of WT and cMyBP-C KO preparations to be phosphorylated; Fig. 4). It may also be possible that cMyBP-C and RLC interact through a more indirect mechanisms that influence conformational transitions of either or both regulatory proteins, thereby influencing myocardial contractility. Future studies clarifying these interactions and how they may be disrupted with decreased RLC phosphorylation and/or common cMyBP-C mutations associated with heart disease may enlighten future therapeutic avenues to improve dysfunctional cardiac contraction.

Henk L. Granzier served as editor.

This work was supported by grants from the National Institutes of Health (R01 HL149164 to B.C.W. Tanner) and the American Heart Association (19TPA34860008 to B.C.W. Tanner).

The authors declare no competing financial interests.

Author contributions: K.L. Turner, D.P. Fitzsimons, and B.C.W. Tanner conceptualized the study. D.P. Fitzsimons procured and distributed the tissue samples. K.L. Turner, H.S. Morris, and P.O. Awinda performed the experiments. K.L. Turner, H.S. Morris, and P.O. Awinda analyzed and curated the data. K.L. Turner and B.C.W. Tanner wrote the original draft of the paper. K.L. Turner, D.P. Fitzsimons, and B.C.W. Tanner reviewed, edited, and revised additional versions of the manuscript, and all authors approved the final version submitted for review.