Disrupted mechanobiology links the molecular and cellular phenotypes in familial dilated cardiomyopathy

Sarah R. Clippinger; Paige E. Cloonan; Lina Greenberg; Melanie Ernst; W. Tom Stump; Michael J. Greenberg

doi:10.1073/pnas.1910962116

Edited by James A. Spudich, Stanford University School of Medicine, Stanford, CA, and approved July 23, 2019 (received for review June 27, 2019)

Significance

One of the outstanding challenges in understanding familial dilated cardiomyopathy has been connecting mutation-induced changes in sarcomeric protein function with the phenotype seen in cells. Many of the mutation-induced changes in contractility at the molecular scale are subtle, begging the question of how they lead to a devastating progressive disease. Here, we show that disease-causing mutations in sarcomeric proteins affect not only contraction but also how cardiomyocytes sense and respond to changes in their mechanical environment associated with aging and the disease progression. Our results implicate impaired mechanosensing as an underappreciated mechanism in the pathogenesis and progression of dilated cardiomyopathy, and they have important implications for the design of precision therapeutics.

Abstract

Familial dilated cardiomyopathy (DCM) is a leading cause of sudden cardiac death and a major indicator for heart transplant. The disease is frequently caused by mutations of sarcomeric proteins; however, it is not well understood how these molecular mutations lead to alterations in cellular organization and contractility. To address this critical gap in our knowledge, we studied the molecular and cellular consequences of a DCM mutation in troponin-T, ΔK210. We determined the molecular mechanism of ΔK210 and used computational modeling to predict that the mutation should reduce the force per sarcomere. In mutant cardiomyocytes, we found that ΔK210 not only reduces contractility but also causes cellular hypertrophy and impairs cardiomyocytes’ ability to adapt to changes in substrate stiffness (e.g., heart tissue fibrosis that occurs with aging and disease). These results help link the molecular and cellular phenotypes and implicate alterations in mechanosensing as an important factor in the development of DCM.

Dilated cardiomyopathy (DCM) is a major cause of sudden cardiac death in young people, and it is a significant cause of heart failure (1). DCM is phenotypically characterized by dilation of the left ventricular chamber, and it is often accompanied by changes in cellular and tissue organization, including lengthening of individual myocytes and fibrosis-induced stiffening of the myocardial tissue. Genetic studies have demonstrated that familial DCM can be caused by single point mutations, many of which are in sarcomeric proteins responsible for regulating cardiac contractility (1); however, the connection between the initial insult of molecular-based changes in contractile proteins and the development of the cellular disease phenotype is not thoroughly understood. This lack of understanding has hampered efforts to develop novel therapeutics, and there is currently no cure for DCM, with heart transplantation being the only long-term treatment. The goal of this study was to better understand the connection between mutation-induced changes in contractility and the cellular phenotype in DCM.

Understanding the link between point mutations in sarcomeric proteins and the development of the disease phenotype in cells has been challenging for several reasons. One challenge stems from the fact that the clinical presentation and prognosis appear to depend on the specific mutation. In fact, different point mutations within the same molecule, and even different substitutions at the same amino acid site, can lead to different forms of cardiomyopathy (e.g., hypertrophic, dilated, restrictive) with different ages of onset (2 ⇓⇓–5). Another challenge stems from selecting appropriate model systems of the human disease for the specific question being posed. In vitro biochemical studies are excellent for deciphering the molecular consequences of the initial insult (4); however, they are not clearly predictive of how the disease will manifest itself in cells (6). Studies using transgenic mice have significantly advanced our understanding of the disease; however, these mice do not always recapitulate the human disease phenotype due to differences in cardiac physiology between humans and mice (7 ⇓⇓⇓⇓⇓–13). Patient tissue is excellent for studying the disease phenotype in humans (14); however, tissue from patients is difficult to obtain and usually only available from patients in the advanced stages of the disease or post mortem, when compensatory mechanisms can mask the initial insult.

Recent advances in stem cell (15, 16) and gene editing technologies (17) have provided a new avenue for modeling DCM in human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) (18, 19). These hiPSC-CMs can faithfully recapitulate many aspects of the early human disease (20), making them an excellent tool for deciphering how the initial molecular insult leads to the development of the early disease phenotype. Moreover, these cells can be examined in simplified in vitro systems that mimic various aspects of the environment in the heart to dissect how the local environment influences the development of the disease phenotype. It has been shown that cardiomyocytes can adapt their structural organization and contractility in response to their local mechanical and geometric environment (21 ⇓–23). It is possible aberrant responses of cardiomyocytes to disease-related alterations in the local environment such as fibrosis-induced stiffening of the heart tissue could contribute to the development of the disease phenotype.

To uncover the connection between molecular changes and the development of the cellular disease phenotype, we determined the molecular and cellular mechanisms of a DCM-causing point mutation in troponin-T (TNNT2), deletion of lysine 210 (ΔK210) (24) (Fig. 1A). The ΔK210 mutation has been identified in patients as young as 2 y old in at least 4 unrelated families (25 ⇓–27). Troponin-T is a subunit of the troponin complex, which, together with tropomyosin, regulates the calcium-dependent interactions between the force-generating molecular motor myosin and the thin filament. TNNT2 is one of the most frequently mutated genes in DCM (28, 29). There have been several excellent model systems developed to better understand aspects of the disease caused by this mutation (30 ⇓⇓⇓–34); however, the connection between the molecular mutation and the cellular phenotype seen in humans remains unclear.

Fig. 1.

(A) Structure of the troponin core complex (Protein Data Bank ID 4Y99), showing troponin-C (cyan), troponin-I (white), troponin-T (purple), and K210 of troponin-T (red). (B) Speed of thin filament translocation as a function of calcium measured in the in vitro motility assay. Error bars show the SD of n = 3 experiments. Data show a significant shift in the pCa50 toward submaximal calcium activation for ΔK210 (P < 0.0001). The Hill coefficient is not changed between the WT and ΔK210 (P = 0.45).

Our results demonstrate that ΔK210 causes a reduction in myosin-based force generation at the molecular and cellular levels, and we used computational modeling to help link the contractile phenotypes seen at these scales. Surprisingly, we found that this disease-causing mutation of a sarcomeric protein not only impairs cardiomyocyte contraction but also causes cellular hypertrophy and impairs the ability of cardiomyocytes to sense and respond to changes in their mechanical environment. Our results suggest a central role for defects in cardiomyocyte mechanosensing in the disease pathogenesis of DCM.

Results

ΔK210 Decreases Calcium Sensitivity in an In Vitro Motility Assay.

We set out to decipher the molecular mechanism of the ΔK210 mutation in vitro. The molecular effects of cardiomyopathy mutations depend on the myosin heavy chain isoform (7 ⇓–9, 35 ⇓–37), and therefore, we used porcine cardiac ventricular myosin (38). Porcine ventricular cardiac myosin (MYH7) is 97% identical to human, while murine cardiac myosin (MYH6) is only 92% identical. Unlike murine cardiac myosin, porcine cardiac myosin has very similar biophysical properties to human cardiac myosin, including the kinetics of the ATPase cycle, step size, and sensitivity to load (38 ⇓⇓–41), making it an ideal myosin for biophysical studies.

Given the role of troponin-T in thin filament regulation, we first determined whether the ΔK210 mutation affects calcium-based regulation of myosin’s interactions with thin filaments using an in vitro motility assay (42). Reconstituted thin filaments, consisting of porcine cardiac actin and recombinantly expressed human troponin complex and tropomyosin, were added to a flow cell coated with porcine cardiac myosin in the presence of ATP. The speed of filament translocation was measured as a function of added calcium. As has been reported previously, the speed of regulated thin filament translocation increased sigmoidally with increasing calcium concentration (43) (Fig. 1B). Data were fit with the Hill equation to obtain the pCa50 (i.e., the concentration of calcium necessary for half-maximal activation). Consistent with previous studies using mouse cardiac, rabbit cardiac, and rabbit skeletal muscle fibers (31, 33, 44), ΔK210 shows a right-shifted curve (pCa50 = 5.7 ± 0.1) compared to the wild type (WT) (pCa50 = 6.1 ± 0.1; P < 0.0001), meaning more calcium is needed for the same level of activation. This suggests that the mutant could show impaired force production during a calcium transient.

Molecular Mechanism of ΔK210-Induced Changes in Thin Filament Regulation.

The changes in calcium sensitivity seen in the in vitro motility assay could be due to changes in the kinetics of myosin attachment and/or detachment from the thin filament. To test whether the mutation affects the kinetics of myosin detachment from thin filaments, we measured the rate of ADP release from myosin, the transition that limits actomyosin dissociation at saturating ATP in the absence of load (45). Consistent with previous studies of other DCM troponin mutations (46), ΔK210 does not change the kinetics of ADP release from myosin (SI Appendix, Fig. S1). Therefore, the primary effects of the mutation are likely due to alterations in the kinetics of myosin attachment to the thin filament.

The kinetics of myosin attachment to the thin filament are regulated by myosin and the thin filament proteins troponin and tropomyosin. Biochemical (47) and structural biological (48, 49) experiments have shown that thin filament regulation is a complicated process where tropomyosin can lie along the thin filament in 3 positions (blocked, closed, and open), and that this positioning is determined by both calcium and myosin binding (Fig. 2A). To determine the molecular mechanism of the observed changes in the calcium sensitivity of myosin-based motility, we determined the equilibrium constants for the transitions between these states using an approach pioneered by McKillop and Geeves (47).

Fig. 2.

Determination of equilibrium constants governing thin filament regulation for ΔK210. (A) The 3-state model for tropomyosin positioning along the thin filament (47). Activation of the thin filament and subsequent force generation requires both calcium and myosin binding. The equilibrium constant for the blocked to closed transition is K_B, and the equilibrium constant for the closed to open transition is K_T. K_W and K_S are the equilibrium constants for myosin weak and strong binding to the thin filament, respectively. (B and C) Normalized stopped-flow fluorescence traces of myosin binding to regulated pyrene-actin. The quenching of pyrene-actin upon binding of S1 occurs at a faster rate at pCa 4 (green) compared to pCa 9 (pink) due to the activating effect of calcium. Traces for (B) WT and (C) ΔK210 are very similar, and the calculated values of K_B are not statistically different (P = 0.998). (D and E) Fluorescence titrations in which increasing amounts of S1 myosin were gradually added to regulated thin filaments to a final concentration of 10 μM S1 in the presence of ADP. Titration curves at 3 different calcium concentrations are shown for both (C) WT and (D) ΔK210. Five repeats were performed for each condition, and error bars show the SD. At low concentrations of myosin, more myosin is bound to the thin filament at higher calcium concentrations (pCa 6.25 [orange] and pCa 3 [green]) than at low calcium (2 mM EGTA, pink).

The equilibrium constant for the transition between the blocked and closed states, K_B, was measured using stopped-flow kinetic techniques (47), where the rate of myosin binding to pyrene-labeled regulated thin filaments was measured at high (pCa 4) and low (pCa 9) calcium concentrations. Myosin strong binding to labeled thin filaments quenches the pyrene fluorescence, leading to a roughly exponential decrease in fluorescence (Fig. 2 B and C). The ratio of the observed rates for myosin binding to the thin filament at high and low calcium can be used to calculate the equilibrium constant, K_B (47):kobs(−Ca2+)kobs(+Ca2+)≈KB1+KB.[1]The values of K_B obtained were 0.36 ± 0.15 for WT (n = 3) and 0.36 ± 0.10 for ΔK210 (n = 3), indicating that ΔK210 does not significantly affect the equilibrium constant for the transition between the blocked and closed states (P = 0.99).

To determine the equilibrium constant for the transition between the closed and open states, K_T, we measured the steady-state binding of myosin to pyrene-labeled regulated thin filaments over a range of myosin concentrations (Fig. 2 D and E). Titration relationships obtained for WT and ΔK210 at high (pCa 3) and low (2 mM EGTA) calcium concentrations show 2-state (i.e., no calcium-based blocking) and 3-state processes, respectively, as has been observed previously (47). Five technical replicates were used to define each curve. As described in SI Appendix, Materials and Methods, we modified the fitting and analysis procedure used by McKillop and Geeves to better define the values of the fitted variables (50). We performed an additional titration at an intermediate calcium concentration (pCa 6.25) and used an annealing algorithm to globally fit the binding curves of the fractional change in pyrene fluorescence (a) as a function of myosin concentration for all 3 calcium concentrations (Fig. 2 D and E). The fractional change in the pyrene fluorescence is given by the following (47):a=F0−FF0−F∞=KW[M]Pn−1(KT(1+KS)n+1)(KTPn+Qn+1KB)(1+KS)n−1,[2]where [M] is the concentration of myosin, F is the measured fluorescence, F₀ is the fluorescence in the absence of myosin, F_∞ is the fluorescence at saturating myosin concentrations, n is the size of the cooperative unit, P = 1 + K_W[M] (1 + K_S), and Q = 1 + K_W[M] (Fig. 2 D and E). For the fitting, K_T, K_W (i.e., the equilibrium constant between the open and myosin weakly bound states, sometimes denoted K₁), and n were fit parameters. K_B and K_S (the equilibrium constant between the myosin weakly and strongly bound states, sometimes denoted K₂) were fixed based on the calculated values for K_B (Fig. 2 B and C) and previously measured values of K_S (51). For the fitting at pCa 6.25 and 4, K_B was fixed at 20, since the fit is insensitive to the exact value of K_B at those calcium concentrations, as long as K_B is greater than 5. 95% confidence intervals were calculated from 1,000 rounds of bootstrapping simulations (SI Appendix, Materials and Methods).

We found that while the measured values of K_T for ΔK210 were consistently lower compared to the WT at all calcium concentrations (Fig. 3A), only the difference in K_T at pCa 6.25 was statistically significant (P = 0.03). From the measured equilibrium constants, it is straightforward to calculate the fraction of regulatory units in each state from the partition function (47). At all calcium concentrations, ΔK210 shows a decrease in the fraction of thin filaments in the open, weakly bound, and strongly bound states compared to the WT (Fig. 3B and SI Appendix, Fig. S7), and an increase in the fraction of thin filaments in the blocked and closed states. These data give the biochemical mechanism of the reduced activation at subsaturating calcium seen in the in vitro motility assays (Fig. 1B) and would predict a reduction in the force per sarcomere in mutant muscle due to a reduction in the fraction of myosin cross-bridges in the strongly bound state.

Fig. 3.

Effects of ΔK210 on tropomyosin positioning along the thin filament. (A) The 3-state model for tropomyosin positioning along the thin filament (47) and parameter values obtained from the stopped-flow and fluorescence titration experiments. Reported uncertainties are 95% confidence intervals. K_T for ΔK210 at pCa 6.25 is statistically different from WT (P = 0.03). Error bars are the 95% confidence intervals. (B) Percent change in the occupancy of each state for the mutant compared to the WT. Reds denote increased population of a state, while blues denote decreased population. ΔK210 causes a decrease in the population of the states where myosin is bound to the thin filament and an increase in the population of the inhibited (i.e., blocked and closed) states. Note that the fractional change in the population of thin filaments in the blocked and closed states is similar, since K_B does not change between the WT and the mutant.

Computational Modeling Recapitulates the Shift in Calcium Sensitivity and Predicts a Lower Force per Sarcomere for ΔK210.

To predict how the observed changes in the equilibrium populations of thin filament regulatory units correspond with contractility in a sarcomere, we used a computational model developed by Campbell et al. (52). This model was chosen since it has many similarities to the McKillop and Geeves biochemical model of muscle regulation used to analyze the biochemical experiments (47), although it should be noted that there are some important differences (Discussion and SI Appendix, Materials and Methods). The Campbell et al. model simulates an ensemble of sarcomeric regulatory units that can transition between the blocked, closed, open, and myosin bound states based on the equilibrium constants between these states and a constant describing the coupling between adjacent regulatory units. This model enables the calculation of both the expected steady-state force as a function of calcium and the expected force produced per sarcomere in response to a calcium transient. The details of the implementation of the model are described in Materials and Methods. Using our measured equilibrium constants, the steady-state normalized force was simulated for both WT and ΔK210 over a range of calcium concentrations (Fig. 4A). Simulation of ΔK210 shows a rightward shift toward supermaximal calcium activation, consistent with the shift seen in the in vitro motility assay (Fig. 1B). Therefore, the observed set of changes in the equilibrium constants in the mutant are consistent with the observed changes seen in the in vitro motility assay, validating the approach taken here.

Fig. 4.

Computational modeling of the steady-state and transient force per sarcomere. (A) Using the model developed by Campbell et al. (52), the steady-state normalized force per sarcomere as a function of calcium can be simulated for both ΔK210 and WT. The simulation shows a shift in the pCa50 toward supermaximal calcium activation in the mutant. (B) Simulated twitch forces per sarcomere in response to a calcium transient. The model predicts that ΔK210 should show a lower twitch force per sarcomere.

To predict the effects of the ΔK210 mutation on the force per sarcomere, we used the same computational model to simulate the force per sarcomere in response to a calcium transient. In the modeling, we assumed that the primary effects of the mutation are on thin filament positioning, and that to a first-order approximation, the calcium transient is not significantly changed by the mutation. The consequences of these assumptions are explored in Discussion. The simulation predicts a smaller maximal twitch force per sarcomere in response to a calcium transient for ΔK210 compared to the WT (Fig. 4B). Taken together, these simulations predict that ΔK210 should decrease the force per sarcomere in cardiomyocytes.

Generation of ΔK210-Induced Stem Cell-Derived Cardiomyocytes.

To examine the effects of the ΔK210 mutation in human cells, we generated hiPSC-CMs. The hiPSC lines were derived from the BJ foreskin fibroblast line by the Washington University Genome Engineering core (SI Appendix, Materials and Methods). Whole-exome sequencing of the stem cell line demonstrated that these cells do not have any genetic variants associated with familial cardiomyopathies (SI Appendix, Fig. S2). Cell lines homozygous for the ΔK210 mutation were generated using the CRISPR/Cas9 system (17) (SI Appendix, Fig. S3 and Materials and Methods). ΔK210 mutant stem cells have normal karyotypes (SI Appendix, Fig. S4) and are pluripotent, as assessed by immunofluorescence staining for pluripotency markers (SI Appendix, Fig. S5). For all experiments with the mutant hiPSC-CMs, 2 separate stem cell lines were used to ensure that the observations are not due to potential off-target cuts introduced during the CRISPR/Cas9 editing. We also used hiPSC-CMs from at least 2 separate differentiations per cell line for both WT and ΔK210. hiPSCs were differentiated to hiPSC-CMs using established protocols (15, 53) involving temporal modulation of WNT signaling using small molecules (SI Appendix, Fig. S5) and aged at least 30 d before use. Using this procedure, >90% cardiomyocytes were obtained, as determined by immunofluorescence staining for cardiac troponin-T (SI Appendix, Fig. S5).

To examine the sarcomeric organization of hiPSC-CMs, cells cultured on glass were fixed and stained for troponin-I and α-actinin to visualize the thin filament and z-disks, respectively (Fig. 5). Unlike adult human tissue-derived cardiomyocytes, which are rectangular with sarcomeres that align with the long axis of the cell, WT hiPSC-CMs cultured on glass orient randomly and display robust sarcomeric staining that is typically not aligned along a single axis (Fig. 5A). Compared to the WT hiPSC-CMs, ΔK210 hiPSC-CMs display more disorganized sarcomeres with patches of punctate staining (Fig. 5B). Similar disorganization and punctate sarcomere structure has been seen with other hiPSC-CM models of DCM cultured on stiff substrates (6, 18, 54), suggesting that it might be a common feature of some forms of DCM.

Fig. 5.

Immunofluorescence images of sarcomeres in hiPSC-CMs. Troponin-I is red and α-actinin is green. All images are z-projections. (A) WT hiPSC-CM on glass. (B) ΔK210 hiPSC-CM on glass, showing sarcomeric disorganization. (C) WT cell on a rectangular pattern on glass. (D) ΔK210 cell on a rectangular pattern on glass. (E) WT cell on a rectangular pattern on a 10-kPa hydrogel. (F) ΔK210 cell on a rectangular pattern on a 10-kPa hydrogel. The sarcomeric organization is significantly improved on the hydrogel with a physiologically relevant stiffness.

ΔK210 hiPSC-CMs Show a Pronounced Increase in Size and Mechanosensitive Alterations in Sarcomeric Structure.

Cardiomyocytes are sensitive to their local physical environment, and it has been shown that providing physical cues that mimic the environment of the heart can improve cardiomyocyte structure and contractility (21, 55). During aging and DCM disease progression, the heart becomes stiffer, and the myocytes become disordered. We hypothesized that ΔK210 hiPSC-CMs would show aberrant responses to changes in their physical environment. To test this hypothesis, we examined the size and structure of cells on glass, which has a high stiffness (approximately gigapascals) and on hydrogel substrates with a stiffness matched to healthy heart tissue (10 kPa).

To examine the role of geometric cues in cellular organization, we micropatterned extracellular matrix for cellular adhesion in rectangular patterns with a 7:1 aspect ratio. It was previously shown that micropatterning of hiPSC-CMs improves cell maturity and the 7:1 aspect ratio optimizes force production (21). We examined live cells using bright-field microscopy, and fixed cells stained for the z-disk marker α-actinin and troponin-I using confocal microscopy (Fig. 5 C–F). Interestingly, the ΔK210 hiPSC-CMs are significantly larger than the WT in fixed cells on glass (Figs. 5 and 6), fixed cells on 10-kPa hydrogels (Figs. 5 and 6), and in live cells on 10-kPa hydrogels (Fig. 7). We analyzed a large number of cells and generated cumulative distributions of cell sizes for single cells, since this methodology does not assume a form for the underlying distribution (Figs. 6 and 7). The cumulative distribution for area shows the fraction of cells with an area less than or equal to the value of the x axis. The increase in size in the mutant is due to increases in both cell width and length (Table 1 and Fig. 6). These data demonstrate that ΔK210 hiPSC-CMs are larger than the WT cells, and that this increase in size is not dependent on the mechanical environment.

Fig. 6.

Quantification of immunofluorescence staining for sarcomeres in hiPSC-CMs on rectangular patterns on (A) glass and (B) 10-kPa hydrogels. Data are plotted as cumulative distributions and parameter values can be found in Table 1. ΔK210 cells are significantly larger on both glass and 10-kPa hydrogels (P = 0.00002), due to increases in both length and width. The orientation order parameter (OOP) is a measurement of sarcomeric organization. For a hiPSC-CM in which all of the sarcomeres are aligned along a single axis, the OOP = 1, and for a hiPSC-CM with no preferred axis, the OOP = 0. On glass, ΔK210 cells have lower OOP values than the WT (P = 0.008). On physiologically stiff hydrogels, there is no significant difference between the OOP of the WT and mutant cells (P = 0.5).

Fig. 7.

Traction force microscopy of single hiPSC-CMs patterned on 10-kPa polyacrylamide gels. (A) Live ΔK210 hiPSC-CMs on hydrogels are significantly larger than WT cells. (B) ΔK210 cells have a lower force per area (i.e., lower force per sarcomere), consistent with the molecular studies; however, since they are larger, (C) their total force production is not significantly different from the WT cells. Reported uncertainties are 95% confidence intervals.

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Table 1.

Analysis of fixed cell sizes on rectangular patterns (reported uncertainties are 95% confidence intervals)

Next, we examined the influence of substrate stiffness on the sarcomeric organization of ΔK210 hiPSC-CMs. We fixed and stained micropatterned hiPSC-CMs for the sarcomeric marker α-actinin on both glass and 10-kPa substrates. Both WT and ΔK210 hiPSC-CM sarcomeres orient preferentially along the long axis of the cells patterned on glass and on 10-kPa hydrogels (Fig. 5). To quantify the degree of sarcomeric organization, we used a program developed by Pasqualini et al. (55). The program analyzes fluorescence images with periodic structure, and it calculates the orientational order parameter (OOP) (i.e., the fraction of z-disks that are oriented along a single axis). For cells with z-disks all oriented along a single axis, OOP = 1, and for cells with z-disks with no preferred axis, OOP = 0.

When patterned on glass (gigapascal stiffness), the ΔK210 mutant hiPSC-CMs show a reduction in the fraction of z-disks aligned along the long axis compared to the WT (P = 0.008) (Fig. 6A and Table 1). This is consistent with what we saw with unpatterned cells on glass (Fig. 5 A and B). Strikingly, when patterned on hydrogels that mimic the stiffness of healthy heart tissue, the ΔK210 cells show significantly improved z-disk organization (Fig. 6B and Table 1). As quantified using the OOP, the organization of the ΔK210 z-disks are indistinguishable from the WT on 10-kPa substrates (P = 0.5). Taken together, these results demonstrate that the ΔK210 mutation affects how the hiPSC-CMs respond to their mechanical environment, with ΔK210 cells showing normal z-disk organization on physiologically stiff substrates, but reduced sarcomeric organization on stiff substrates.

ΔK210 Cells Show Altered Contractility and Features of Hypertrophy on Substrates of Physiological Stiffness.

Our biochemical measurements (Fig. 3B) and computational modeling predict that ΔK210 cells should show a lower force per sarcomere (Fig. 4B); and therefore, we examined whether the ΔK210 mutation affects force production of single hiPSC-CMs. To measure force production of single hiPSC-CMs, we used traction force microscopy. Spontaneously beating cells were micropatterned onto 10-kPa hydrogels prepared with embedded fluorescent microbeads to monitor beating. Spinning disk confocal imaging was used to monitor the bead displacement as a function of time, and the total force of contraction for each cell was calculated using a MATLAB routine developed by Ribeiro et al. (56) (Fig. 7 B and C). For each cell, the force per sarcomere was approximated by calculating the force per area.

The average total force per cell measured by traction force microscopy is the same for both WT [0.27 (−0.03/+0.04) μN] and ΔK210 hiPSC-CMs [0.27 (−0.04/+0.05) μN; P = 0.98] (Fig. 7C). These forces are consistent with previous measurements (21). For each cell analyzed by traction force microscopy, we measured its area from bright-field illumination images (Fig. 7A). Bright-field images of live cells lack the resolution of the immunofluorescence images, making it challenging to accurately measure the cross-sectional area of the live cells; however, ΔK210 hiPSC-CMs are significantly larger than the WT cells (Fig. 7A). Moreover, fixed ΔK210 hiPSC-CMs on 10-kPa hydrogels have a z-disk organization that is indistinguishable from the WT (Fig. 6B). We calculated the force per area for each cell (Fig. 7B). The ΔK210 cells have a lower force per area [0.25 (−0.04/+0.05) μN/μm²] compared to the WT [0.32 (−0.04/+0.04) μN/μm²; P = 0.01]. The lower force per area for ΔK210 is consistent with our biochemical measurements and the computational modeling, which predicted a lower force per sarcomere (Fig. 4B). These data help link the molecular and cellular phenotypes. Moreover, they demonstrate that although the force per sarcomere is reduced in the mutant, the ΔK210 hiPSC-CMs can compensate through an increase in size that results in a total force per cell that is indistinguishable from the WT.

Discussion

Here, we determined the molecular mechanism of a mutation in troponin-T that causes DCM in humans, ΔK210. We found that this mutation affects not only molecular and cellular contractility, but also the ability of the cardiomyocytes to respond to changes in the mechanical environment like stiffening of heart tissue associated with aging and disease in patients. These results implicate defective mechanosensing by cardiomyocytes as an important factor in the pathogenesis of DCM caused by mutations in sarcomeric proteins, and they highlight the importance of multiscale studies in understanding heart disease.

Proposed Molecular Mechanism for ΔK210 and Its Relationship to Previous Studies.

The disease presentation of familial cardiomyopathies and their biophysical manifestation (i.e., contractile and force-dependent properties) can depend on the myosin isoform (7, 8, 57 ⇓⇓–60). Our in vitro motility results (Fig. 1B) using porcine cardiac actin and ventricular β-cardiac muscle myosin (MYH7), which closely mimics the biophysical and biochemical properties of human ventricular β-cardiac muscle myosin (38, 39), demonstrate a shift toward supermaximal calcium activation, consistent with previous experiments studying the ΔK210 mutation using different myosin isoforms (31, 44).

We determined that this shift is due to alterations in the equilibrium constants that govern the positioning of tropomyosin along the thin filament, leading to a reduction in the population of force-generating cross-bridges at submaximal calcium concentrations (Figs. 3 and 4A). The structural basis of these mutation-induced shifts is not well understood; however, it is possible that the mutation affects the interaction between the troponin complex and tropomyosin (61) and/or the allosteric coupling between subunits of the troponin complex (2, 5, 62, 63). These nonexclusive mechanisms could contribute to the observed decrease in force-producing states seen in ΔK210 thin filaments, and future studies should be able to shed light on the structural basis of these changes.

Our biochemical data were analyzed using the formalism developed by McKillop and Geeves (47). These data clearly demonstrate a decrease in the population of strongly bound myosin cross-bridges in the mutant (Fig. 3B and SI Appendix, Fig. S7), suggesting that the mutant would show reduced force production. We predicted the effects of the mutations on sarcomeric contractility using a computational model developed by Campbell et al. (52) that is similar, but not identical to the McKillop and Geeves model (47). For example, the McKillop and Geeves model has a calcium dependence for K_T, while this term is calcium independent in the Campbell et al. model. Although these models are not exactly equivalent, the computational modeling was able to recapitulate the shift in calcium sensitivity seen in the motility assay (Figs. 1B and 4A). Other models of thin filament regulation exist (64 ⇓⇓⇓–68), but the McKillop and Geeves model is one of the most frequently used. Regardless of the model used to interpret the results, our biochemical data clearly show molecular hypocontractility in the mutant at micromolar calcium concentrations.

The computational modeling based on our biochemical studies predicts a larger reduction in the force per sarcomere than we observed in hiPSC-CMs (Fig. 7). This is likely due to several necessary simplifying assumptions made in the modeling: 1) We assumed that the mutation does not change the magnitude or time course of the calcium transient, which is not necessarily true given that the knockin ΔK210 mouse shows altered calcium transients (31). 2) We assumed that the troponin in hiPSC-CMs has the same biochemical properties as the adult cardiac troponin complex that we examined in our biochemical experiments. This is not necessarily the case, since 4 troponin-T isoforms can be detected in the human heart at different developmental stages (69, 70), and hiPSC-CMs primarily express a slow skeletal muscle isoform of troponin-I (71), rather than the cardiac isoform used in the biochemical studies. While all isoforms of troponin expressed in hiPSC-CMs would have the ΔK210 mutation (SI Appendix, Fig. S3), different isoforms have different calcium sensitivities (69). That said, even with these simplifying assumptions, the simulations were able to predict the reduction in the force per sarcomere seen in the ΔK210 cardiomyocytes.

Linking the Molecular and Cellular Contractile Phenotypes for ΔK210.

Patients with the ΔK210 mutation show early onset of the disease phenotype and a high incidence of sudden death (26). Aspects of the disease have been replicated in knockin mouse models (31), which have significantly advanced our understanding of the disease phenotype. However, the ΔK210 mouse model shows significant differences with respect to the human phenotype, due to inherent physiological differences between mouse and human hearts, including differences in gene expression, calcium-handling machinery, and ion channel composition (34).

We used human hiPSC-CMs to study how the initial insult of a molecular-based change leads to the early disease pathogenesis in human cells. For these studies, hiPSC-CMs are excellent models, since they can capture cellular changes that occur before major compensatory mechanisms (e.g., fibrosis, activation of neurohormonal pathways, changes in gene expression) mask the initial molecular phenotype (18 ⇓–20, 54, 72). Contractile changes associated with the disease have been observed in cardiomyopathy patients before adverse remodeling of the heart (73 ⇓–75). That being said, hiPSC-CMs are developmentally immature, meaning that they do not necessarily recapitulate all of the aspects of the phenotype in adults or the later stages of the disease progression. All of our molecular and cellular studies modeled the homozygous ΔK210 mutation, so care should be used when extrapolating the results here to the phenotype seen in patients.

ΔK210 hiPSC-CMs on patterned hydrogels show a reduced force per area compared to WT cells (Fig. 7B) without any significant differences in the z-disk organization (Fig. 6B). These data are consistent with the reduced force per sarcomere predicted from our biochemical experiments (Fig. 3B) and computational modeling (Fig. 4B), linking the molecular phenotype with cellular contractility. It should be noted that our studies only examined the z-disk organization and that it is possible that the mutation could affect the expression of other sarcomeric proteins.

Interestingly, while the force per sarcomere is reduced in the mutant, the total force per cell is not different from the WT (Fig. 7C). This can be explained by the fact that the mutant cells are larger (Figs. 6 and 7), a result that would not have been predicted from the molecular studies alone. This increase in size occurs in single cells, independent of endocrine or cell–cell signaling. One possible explanation for this increase in size is that the cells are undergoing compensatory hypertrophy, which allows the cells to counteract the reduced force per sarcomere. While we have not investigated whether the increase in size seen in our cells is due to activation of hypertrophic signaling pathways, future studies should elucidate the mechanism of the increase in cell size seen here. Recent work from Molkentin and coworkers (76) has shown that reduced total tension developed by cardiomyocytes is strongly correlated with the development of dilated cardiomyopathy in both hiPSC-CMs and mice; however, it is also possible that the increase in cell size could be due to alterations in calcium handling.

Defects in ΔK210 Mechanosensing Contribute to the Disease Phenotype.

Fundamentally, troponin serves as a gate that regulates myosin-based tension. As such, one would expect that the primary effects of the mutation would be to reduce force production; however, the ΔK210 cells show several changes beyond altered contractility. While healthy cardiomyocytes adapt their contractility and structure to changes in their mechanical environment, such as age or disease-related stiffening of the heart tissue (21), ΔK210 cells do not adapt in the same way as the WT cells. Similar to other hiPSC-CM models of DCM grown on stiff substrates, ΔK210 hiPSC-CMs cultured on glass show impaired sarcomeric organization (18, 54, 77); however, ΔK210 hiPSC-CMs z-disks can organize normally when given mechanobiological cues that mimic the healthy heart (Figs. 5 and 6). Therefore, the ΔK210 mutation affects not only contractility but also the ability of the hiPSC-CMs to sense and respond to their environment.

The connection between the reduction in myosin-generated tension in the mutant cells and an impaired ability to respond to mechanical forces warrants further investigation; however, a possible link between cellular tension and sarcomeric organization in DCM was revealed by live-cell imaging of sarcomerogenesis in hiPSC-CMs (54). It was shown that pharmacological inhibition of myosin contractility or genetic disruption of titin leads to an impaired ability of cells to generate well-organized sarcomeres. Chopra et al. proposed that the transduction of myosin-driven tension is necessary for the proper assembly of sarcomeres in hiPSC-CMs. We propose that a similar mechanism could be relevant in ΔK210 hiPSC-CMs, which show reductions in myosin-based contractility and z-disk disorganization when patterned on glass. Consistent with this idea, treatment of hiPSC-CMs containing a different DCM mutation with omecamtiv mecarbil, a myosin-based thin filament activator (41, 78, 79), leads to an improvement in sarcomeric organization (77). We speculate that some of the salutary effects of the drug are due to the restoration of tension necessary for proper mechanosensing.

The result that ΔK210 hiPSC-CMs show altered responses to changes in their mechanical environment has important implications for the mechanism of the disease pathogenesis. Previous studies of cardiomyocytes have shown that the organization of sarcomeres varies depending on substrate stiffness (21, 77), with peak contractility occurring on substrates of physiological stiffness. ΔK210 hiPSC-CMs can organize properly and they have normal force production on substrates that match the stiffness of the healthy heart (Figs. 6 and 7); however, heart tissue becomes stiffer with the disease progression and aging. We propose that as the heart tissue becomes stiffer, ΔK210 cells would become more disordered, effectively causing a progressive loss of function, and potentially contributing to myocyte disarray. Thus, the inability of the mutant cells to adapt to changes in their mechanical environment may contribute to the disease pathogenesis and progression.

The Potential Importance of Mechanosensing in DCM Pathogenesis.

Our results suggest that reduced force at the molecular scale due to a mutation in a sarcomeric protein can lead to impaired mechanosensing in cardiomyocytes. While the disease presentation in DCM depends on the exact mutation (80), we propose that disruption of mechanosensing in cardiomyocytes could be a common mechanism in the disease pathogenesis of other sarcomeric and nonsarcomeric DCM mutations. It has been proposed that DCM can be caused by molecular hypocontractility (4, 76, 81); however, there are also many DCM-causing mutations in nonsarcomeric genes with no known roles in contractility; suggesting that other mechanisms might be involved (1, 3, 82). Most of these genes are located along proposed mechanosensing pathways (83, 84) used to sense and transduce mechanical forces. For example, some of these genes link the cytoskeleton to the extracellular matrix (e.g., dystrophin [DMD], vinculin [VCL]), others link the cytoskeleton to the nucleus (e.g., nesprin-1 [SYNE1], lamin-A [LMNA], centromere protein F [CENP-F]), and others are mechanosensitive transcription factors (e.g., TAZ). Consistent with this idea, altered mechanobiology has been identified in DCM cells with mutations in the nonsarcomeric protein lamin A/C (82). Moreover, patients with mutations in these genes, such as those with Duchenne muscular dystrophy and Emery–Dreifuss muscular dystrophy, often develop DCM. It is intriguing to speculate that impaired mechanosensing by cardiomyocytes in the heart could link some of these seemingly different diseases and provide a common pathway that could be targeted by therapeutic interventions.

Conclusions

Using a combination of biochemical, computational, and cell-biological techniques, we reveal several features of the early disease pathogenesis of a DCM mutation that would have been missed by studying the molecular or cellular phenotypes alone. We demonstrate that the ΔK210 mutation in human cardiac troponin-T causes a change in the equilibrium positioning of tropomyosin, leading to alterations in the calcium sensitivity of thin filament activation and a reduction in the number of strongly bound myosin cross-bridges at all calcium concentrations. Computational modeling predicts that these changes should lead to a reduction in the force generated per sarcomere in mutant cells, and we demonstrate that this is indeed the case in hiPSC-CMs. We demonstrate that the ΔK210 hiPSC-CMs can increase in size to normalize their force production. Moreover, our results demonstrate that the ΔK210 mutation affects the structural organization of hiPSC-CMs and alters how they respond to their mechanical environment. This impaired mechanosensing likely contributes to the DCM disease progression and the development of myocyte disarray as the heart stiffens. Taken together, these results demonstrate that disease-causing mutations of sarcomeric proteins affect not only contraction but also how cardiomyocytes sense and respond to changes in their mechanical environment associated with aging and disease. These data also suggest that disrupting mechanosensing pathways can contribute to the disease phenotype in DCM.

Materials and Methods

Investigation of the Molecular Mechanism Using Recombinant Proteins.

Human troponin and tropomyosin were expressed recombinantly in Escherichia coli, and cardiac actin and myosin were purified from porcine ventricles. In vitro motility assays were conducted as previously described (36). The equilibrium constants that govern thin filament activation were determined using stopped-flow and steady-state fluorescence techniques (47, 50). Full details are provided in SI Appendix, Materials and Methods.

Computational Simulations.

Simulations of force production by sarcomeres were conducted using the model developed by Campbell et al. (52). Full details are provided in SI Appendix, Materials and Methods.

Differentiation of hiPSCs to hiPSC-CMs.

The parent stem cell line, BJFF.6, was generated from the human BJ fibroblast line (CRL-2522, ATCC) by the Genome Engineering and iPSC Center at Washington University in St. Louis. Two independent stem cell lines homozygous for the ΔK210 deletion were generated using the CRISPR/Cas9 system (17, 85). Differentiation to hiPSC-CMs was accomplished using the method of Lian et al. (53). Full details are provided in SI Appendix, Materials and Methods.

Analysis of Sarcomeric Structure in Cardiomyocytes.

hiPSC-CMs were fixed in 4% paraformaldehyde (Electron Microscopy Sciences), and sarcomeric structure was visualized via immunofluorescence using confocal microscopy (Washington University Center for Cellular Imaging). Analysis of the organization of the sarcomeres was conducted using software developed by Parker and coworkers (55). Full details are provided in SI Appendix, Materials and Methods.

Analysis of Single Cardiomyocyte Contractility.

Single hiPSC-CMs were seeded onto rectangular patterns on hydrogels for traction force microscopy according to the protocol of Ribeiro et al. (21). All traction force microscopy measurements were performed on patterned polyacrylamide hydrogels with a stiffness of 10 kPa with hiPSC-CMs that were at least 30 d postdifferentiation. Single, spontaneously beating cells were imaged in an environmentally controlled chamber (Tokai Hit) on a Nikon spinning disk confocal microscope (Washington University Center for Cellular Imaging). Traction force movies were analyzed by generating displacement maps of the fluorescent beads using a MATLAB program developed by the Pruitt and coworkers (56). Full details are provided in SI Appendix, Materials and Methods.

Acknowledgments

We thank Stuart Campbell and Francesco Pasqualini for sharing their code for the simulations and sarcomeric structure analysis, respectively. We acknowledge Mike Ostap and Ken Margulies for extremely helpful discussions during the early planning phases of this project. We also acknowledge Drew Braet for technical assistance and Samantha Barrick for assistance with biochemical experiments. We acknowledge the Washington University Institute of Materials Science and Engineering for the use of microfabrication instruments and staff assistance, and we thank the Alvin J. Siteman Cancer Center at Washington University School of Medicine and Barnes-Jewish Hospital (St. Louis, MO) for the Genome Engineering and Induced Pluripotent Stem Cell Center, which provided the genome editing service (National Cancer Institute Cancer Center Support Grant P30 CA091842). Confocal microscopy was performed through the Washington University Center for Cellular Imaging supported by Washington University School of Medicine, The Children’s Discovery Institute of Washington University and St. Louis Children’s Hospital (CDI-CORE-2015-505), and the Foundation for Barnes-Jewish Hospital (3770). Exome sequencing was performed by the McDonnell Genome Institute. Funding for this project was provided by a pilot grant from the Children’s Discovery Institute of Washington University and St. Louis Children’s Hospital, the Washington University Center for Cellular Imaging (CDI-CORE-2015-505), the National Institutes of Health (R00HL123623 and R01HL141086 [to M.J.G.]; T32EB018266 [to S.R.C.]), and the March of Dimes Foundation (FY18-BOC-430198 [to M.J.G.]).

Footnotes

↵¹S.R.C. and P.E.C. contributed equally to this work.
↵²To whom correspondence may be addressed. Email: greenberg{at}wustl.edu.

Author contributions: S.R.C., P.E.C., L.G., W.T.S., and M.J.G. designed research; W.T.S. contributed new reagents/analytic tools; S.R.C., P.E.C., L.G., M.E., and M.J.G. performed research; S.R.C., P.E.C., L.G., M.E., and M.J.G. analyzed data; and S.R.C. and M.J.G. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1910962116/-/DCSupplemental.

Published under the PNAS license.

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Disrupted mechanobiology links the molecular and cellular phenotypes in familial dilated cardiomyopathy

Significance

Abstract

Results

ΔK210 Decreases Calcium Sensitivity in an In Vitro Motility Assay.

Molecular Mechanism of ΔK210-Induced Changes in Thin Filament Regulation.

Computational Modeling Recapitulates the Shift in Calcium Sensitivity and Predicts a Lower Force per Sarcomere for ΔK210.

Generation of ΔK210-Induced Stem Cell-Derived Cardiomyocytes.

ΔK210 hiPSC-CMs Show a Pronounced Increase in Size and Mechanosensitive Alterations in Sarcomeric Structure.

ΔK210 Cells Show Altered Contractility and Features of Hypertrophy on Substrates of Physiological Stiffness.

Discussion

Proposed Molecular Mechanism for ΔK210 and Its Relationship to Previous Studies.

Linking the Molecular and Cellular Contractile Phenotypes for ΔK210.

Defects in ΔK210 Mechanosensing Contribute to the Disease Phenotype.

The Potential Importance of Mechanosensing in DCM Pathogenesis.

Conclusions

Materials and Methods

Investigation of the Molecular Mechanism Using Recombinant Proteins.

Computational Simulations.

Differentiation of hiPSCs to hiPSC-CMs.

Analysis of Sarcomeric Structure in Cardiomyocytes.

Analysis of Single Cardiomyocyte Contractility.

Acknowledgments

Footnotes

References

Citation Manager Formats

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Main menu

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Search

Significance

Abstract

Results

ΔK210 Decreases Calcium Sensitivity in an In Vitro Motility Assay.

Molecular Mechanism of ΔK210-Induced Changes in Thin Filament Regulation.

Computational Modeling Recapitulates the Shift in Calcium Sensitivity and Predicts a Lower Force per Sarcomere for ΔK210.

Generation of ΔK210-Induced Stem Cell-Derived Cardiomyocytes.

ΔK210 hiPSC-CMs Show a Pronounced Increase in Size and Mechanosensitive Alterations in Sarcomeric Structure.

ΔK210 Cells Show Altered Contractility and Features of Hypertrophy on Substrates of Physiological Stiffness.

Discussion

Proposed Molecular Mechanism for ΔK210 and Its Relationship to Previous Studies.

Linking the Molecular and Cellular Contractile Phenotypes for ΔK210.

Defects in ΔK210 Mechanosensing Contribute to the Disease Phenotype.

The Potential Importance of Mechanosensing in DCM Pathogenesis.

Conclusions

Materials and Methods

Investigation of the Molecular Mechanism Using Recombinant Proteins.

Computational Simulations.

Differentiation of hiPSCs to hiPSC-CMs.

Analysis of Sarcomeric Structure in Cardiomyocytes.

Analysis of Single Cardiomyocyte Contractility.

Acknowledgments

Footnotes

References

Citation Manager Formats

Article Classifications

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