Experimental details regarding differentiation and enrichment of hESC-CMs in 12–20 days of suspension culture using defined differentiation media supplemented with Wnt-pathway modulators were previously published (Kempf et al., 2014) (see Supplementary Material). Such suspension culture-derived “cardiac bodies” consisting of essentially pure hESC-CMs were enzymatically dissociated and ~118 cells/mm² were plated for long-term cultivation on laminin-coated glass cover-slips. This resulted in a majority of hESC-CMs expressing essentially only the βMyHC isoform at the protein level (Weber et al., 2016). We used hESC-CMs from three different cell batches. In this study, hESC-CM (cf. online Supplementary Material, Video 1) were used after 35–56 days of cultivation on glass cover-slips. hESC-CMs were chemically demembranated using 0.5% Triton-X-100 in the presence of 20 mM BDM as previously described (Weber et al., 2016), and then equilibrated with relaxing-solution containing PIC and 5 mM DTT (without BDM) before the experiment.

hvMFs were isolated from two different non-transplanted donor hearts for which no suitable recipient was found. Long-term cryopreservation of muscular tissues was previously described (Kraft et al., 2013). After quick thawing in Na-solution including 20 mM BDM (~20°C), ventricular fragments were cut in small pieces removing connective tissues and fat traces in cold Na-solution (+BDM), and then demembranated in cold K-solution containing 0.5% Triton-X-100 and 20 mM BDM (~50 min, 5°C). Detergent was rinsed twice using cold K-solution without Triton (~50 min). hvMFs-bundles were prepared freshly before the micromechanical experiments by homogenizing the demembranated ventricular pieces equilibrated with relaxing-solution (+PIC, +DTT 5 mM, on ice) for 5–10 s with a blender (Ultra-Turrax T8, IKA Labortechnik, Germany) at 25,000 rpm. The resulting myofibrillar suspension was filtered through a mesh (pore size 21 μm) to remove the thick myofibrillar aggregates. The homogenate was further centrifuged (400 × g, 10 min, 5°C; Biofuge Primo-R, Thermo-Fisher, Massachusetts, USA) and the pellet re-suspended in fresh relaxing-solution (+PIC, +DTT) or subjected to further treatments.

Approval of Hannover Medical School Ethics Committee was obtained for anonymized use of the different muscle biopsies and the hESC-CMs in our study (approval numbers 2729–2001, 507–2009, 1751–2013). All subjects gave written informed consent at the different institutions in accordance with the Declaration of Helsinki.

Three to five hundred microliters of myofibrillar homogenate in relaxing-solution was pipetted in the pre-cooled chamber of the micromechanical setup and myofibrils allowed to sediment for ~1 h. Then the chamber was filled with relaxing-solution (15°C).

A laminin-coated glass cover-slip containing plated, chemically demembranated d-hESC-CMs was transferred into the chamber of the micromechanical setup, previously filled with relaxing-solution (15°C).

Contractile function of a single d-hESC-CM containing mainly few bundled myofibrils or a small hvMFs-bundle was investigated (Videos 2–4) with a custom-built micromechanical setup mounted on an inverted microscope (Olympus IX-71). Experiments with d-hESC-CMs and hvMFs were performed with solutions from the same batch at 15°C. Working at this temperature allows comparison of the functional results to several biomechanical studies with hvMFs or skinned cardiomyocytes which were previously performed at 15°C (Stehle et al., 2002b; van der Velden et al., 2003b; Piroddi et al., 2007; Walker et al., 2011; Kraft et al., 2013; van Dijk et al., 2014; Pioner et al., 2016; Racca et al., 2016). Some details about the micromechanical setup are described in Supplementary Material.

Sarcomere length (SL) and cross-sectional area (CSA) of myofibrils within d-hESC-CMs and of hvMF-bundles were determined in bright field (BF) or phase contrast (PhC) at 96-fold magnification (Figures 1A,B), using a CCD camera (Hamamatsu Photonics, Herrsching am Ammersee, Germany) attached to microscope. To calculate specific forces (force/CSA, kPa = nN/μm²) the mean diameter of the myofibrils or myofibrillar bundles within d-hESC-CMs and of entire hvMF-bundles was assessed prior to force measurements as previously described (Stehle et al., 2002b; Weber et al., 2016). In particular, for d-hESC-CMs total CSA of all myofibrils observed within the demembranated cells as darker traces on brighter background (Figure 1A₁) was calculated assuming a cylindrical shape of the myofibrils. As a control for myofibrillar content and diameter in d-hESC-CMs, some cells were immunostained against α-actinin prior to Ca²⁺-activation (Figures 1B₁₋₃), as previously described for isolated myofibrils (Telley et al., 2006). This allowed visualizing myofibrillar thickness and distribution within the cardiomyocytes and to further compare these features with the corresponding ones assigned to the darker traces observed either in PhC (e.g., Figures 1B₁₋₃) within the same cardiomyocyte or in BF (e.g., Figure 1A₁). Such analysis suggested essentially the same thickness for the fluorescently labeled myofibrils at their Z-disks and for the dark fascicles within d-hESC-CMs (in BF or PhC). This is also indicated by Video 5 where visualization of sarcomeres by PhC, fluorescence (FL), or overlapped (PhC and FL) is shown. Based on this we measured myofibrillar CSA directly from the dark fascicles in BF or PhC, and thus, reduced the overestimation of CSA which occurs if we would take the total width of the mounted d-hESC-CMs. An overestimated myofibrillar CSA would yield smaller specific force values.

When activated at saturating [Ca²⁺], kinetics of force development (k_ACT) and re-development (k_TR) of d-hESC-CMs (k_ACT = 0.66 ± 0.14 s⁻¹, n = 11; k_TR = 0.74 ± 0.10 s⁻¹, n = 11) and hvMFs (k_ACT = 0.68 ± 0.14 s⁻¹, n = 37; k_TR = 0.68 ± 0.10 s⁻¹, n = 37) were similar, respectively, with values as previously reported for hvMFs (Stehle et al., 2002b; Piroddi et al., 2007; Walker et al., 2011; Racca et al., 2016). Maximum isometric force (F_ACT,max) for d-hESC-CMs (F_ACT,max = 42 ± 10 kPa, n = 12) was significantly (p < 0.001) smaller than the F_ACT,max generated by hvMFs (F_ACT,max = 94 ± 25 kPa, n = 39) (Figure 1C).

However, we cannot exclude the presence of a very low amount of residual αMyHC in d-hESC-CMs [however, under detection limit in gel analysis; see section Overview of Sarcomeric Proteins by Gel Electrophoresis]. To exclude such CMs from the functional analysis, some d-hESC-CMs were immunostained against αMyHC prior micromechanical measurements. Then, F_ACT,max, k_TR, and k_LIN were determined at pCa 4.18 for d-hESC-CMs exhibiting null fluorescence, i.e., βMyHC positive cells which had no αMyHC-fluorescence (Figure 1D). These values were taken as reference values for all d-hESC-CMs that were characterized micromechanically without preceding immunostaining (Table 1). It has been shown previously that k_TR of d-hESC-CMs which also have αMyHC-positive sarcomeres is significantly faster than for purely βMyHC-positive CMs (Weber et al., 2016). Here we included all d-hESC-CMs measured without preceding immunostaining which had individual k_TR and k_LIN values that were smaller than the upper limit set for k_TR to 0.87 s⁻¹ (= mean + 2 × SD = 0.70 + 0.17) and for k_LIN to 0.47 s⁻¹ (= mean + 2 × SD = 0.29 + 0.18), respectively (Table 1). Therefore, there is a 95% probability that d-hESC-CMs included in the analysis without preceding immunostaining are also negative for αMyHC.

PKA-treatment did not significantly affect F_ACT,max, k_ACT, and k_TR force parameters at pCa 4.18 of either myofibrils within d-hESC-CMs (F_ACT,max = 37 ± 11 kPa, n = 15; k_ACT = 0.62 ± 0.13 s⁻¹, n = 15; k_TR = 0.69 ± 0.11 s⁻¹, n = 16) or hvMFs (F_ACT,max = 95 ± 27 kPa, n = 33; k_ACT = 0.64 ± 0.11 s⁻¹, n = 33; k_TR = 0.67 ± 0.09 s⁻¹, n = 33). This is consistent with previous observations using human ventricular myofibrils and skinned cardiomyocytes (Walker et al., 2011; Kraft et al., 2013).

The results show that at saturating [Ca²⁺], cycling βMyHC cross-bridges are able to develop force with similar rate constants, independent of PKA-treatment, for both contractile systems (d-hESC-CMs and hvMFs). Yet, considering the lower maximum force level reached by d-hESC-CMs, the rates by which both contractile systems reach maximum force were significantly different (p < 0.001), being slower for d-hESC-CMs (28.2 ± 8.8 kPa/s, n = 11) than for hvMFs (64.8 ± 26.9 kPa/s, n = 37).

Cardiomyocytes in vivo operate at intermediate [Ca²⁺]. Therefore, it is important also to evaluate and compare myofibrillar contractile function of d-hESC-CMs and hvMFs at submaximal [Ca²⁺]. d-hESC-CMs responded to intermediate [Ca²⁺] with significantly (p < 0.001) higher isometric force when compared to hvMFs (ΔpCa₅₀ = +0.24; Figure 2A). pCa determined at half of the maximum generated force (pCa₅₀) was 6.04 ± 0.08 for d-hESC-CMs and 5.80 ± 0.05 for hvMFs (Figure 2C). The force-pCa curve determined for d-hESC-CMs had similar steepness (n_H) as the curve determined for hvMFs (Figures 2A–C).

Phosphorylation levels of some sarcomeric proteins can persist even after chemical skinning of myocardial samples. In studies with human cardiomyocytes isolated from failing and non-failing hearts (van der Velden et al., 2003b), PKA-treatment shifted the force-pCa curves to the right-side. Here, we found that PKA-treatment also shifted the isometric force-pCa relationships of both d-hESC-CMs (ΔpCa₅₀ = −0.26) and hvMFs (ΔpCa₅₀ = −0.13) distinctly to higher [Ca²⁺], while the steepness (n_H) of the curves was not significantly affected (Figure 2C).

The force response at a given [Ca²⁺] (considering both absolute and relative values) was different in d-hESC-CMs than in hvMFs. We compared at different intermediate [Ca²⁺] the rate constants of force re-development (k_TR) of d-hESC-CMs toward the same final fractional force level F_n as in hvMFs (F_n = F_ACT/F_ACT,max), and plotted k_TR as function of F_n (Figures 3A–D). Relationships between k_TR and F_n obtained for d-hESC-CMs and hvMFs before and after PKA were fitted using the equation k_TR = g_app/(1−F_n·f_app,max/(f_app,max+g_app)), where f_app,max (f_app at F_n = 1) and g_app are the probabilities (rate constants) of cross-bridges to enter and leave the force-generating states, respectively (Brenner, 1991a; Poggesi et al., 2005). Plots of F_n vs. k_TR curves are shown with 95% confidence intervals (95%-CI) to compare separately d-hESC-CMs ± PKA (Figure 3A), hvMFs ± PKA (Figure 3B), d-hESC-CMs with hvMFs (Figure 3C), and d-hESC-CMs + PKA with hvMFs + PKA (Figure 3D).

For d-hESC-CMs at force levels below ~33% of F_ACT,max (and below ~15–20% of F_ACT,max for hvMFs), it was not possible to reliably measure k_TR, because the laminar solutions flow exerted a hydrodynamic pressure on d-hESC-CMs (and on hvMFs) which perturbed the slow exponential rise of the recorded force signal.

We found that for both types of myofibrils receiving or not the PKA-treatment, the F_n vs. k_TR curves (Figures 3A,B) were similar and the 95%-CIs were almost completely overlapped along the entire range of recorded fractional force levels (0 < F_n≤1). At pCa₅₀ (F_n = 0.5), PKA had no effect on k_ACT and k_TR (Figures 4B,D), as at saturating [Ca²⁺] (F_n = 1; Figures 4A,C).

We also compared d-hESC-CMs and hvMFs before and after PKA-treatment (Figures 3C,D), respectively. Despite the fact that at saturating [Ca²⁺] k_ACT and k_TR values were similar when d-hESC-CMs were compared to hvMFs (Figures 4A,C), at submaximal force levels they were different (Figures 3C,D, 4B,D). 95%-CIs of the k_TR vs. F_n curves for d-hESC-CMs and hvMFs did not overlap at intermediate fractional force levels (0 < F_n < 1) before and after PKA-treatment, respectively (Figures 3C,D).

These results indicate that cross-bridge kinetics, determining the force rise toward submaximal force levels, were different for d-hESC-CMs and hvMFs, while PKA-mediated phosphorylation had no significant effect.

Here we determined k_LIN (the rate constant of the first relaxation phase) which is related to tension cost (= ATPase/Force ~ g_app) (Brenner, 1988; Weber et al., 2016), while the second relaxation phase (described by the rate constant k_REL) is related to the rapid drop of force and cardiomyocyte re-lengthening that contributes to the rapidity of the ventricular pressure decay (i.e., during diastole) (Stehle and Iorga, 2010).

k_LIN values corresponding to the relaxation from full Ca²⁺-activation and from pCa₅₀ to pCa > 8 were compared (Figures 4E,F), revealing that k_LIN was the same for both activation levels either in d-hESC-CMs or in hvMFs. This suggests that kinetics of cross-bridges leaving the force-generating states are not linked to the Ca²⁺-dependent occupancy of the previous force levels (Stehle et al., 2003). PKA-treatment accelerated the first relaxation phase of both d-hESC-CMs and hvMFs. k_LIN of d-hESC-CMs treated with PKA had the highest value (Figures 4E,F). Independent of PKA-treatment (± PKA) and of the previous force level (at pCa 4.18 or pCa₅₀), k_LIN for d-hESC-CMs was significantly faster than for hvMFs (Figures 4E,F).

Duration of the first relaxation phase (t_LIN) after rapid Ca²⁺ removal, i.e., the time until the sequential sarcomere lengthening (Videos 2–4) inducing the fast relaxation phase started (Stehle et al., 2002a,b; Poggesi et al., 2005), was similar in d-hESC-CMs and in hvMFs (Figures 4G,H). With PKA-treatment, it became significantly shorter only for hvMFs compared to d-hESC-CMs.

Due to the reduction in strain on the cross-bridges during the second relaxation phase (Figure 1C-right panel, Videos 2–4), the redistribution of cross-bridges toward the non-force-generating states was accelerated by ~10–20 times from the k_LIN values (<0.4 s⁻¹) to the values of k_REL of ~4–5 s⁻¹ for both d-hESC-CMs and hvMFs (Figures 4I,J). Whereas, strained cross-bridges during the first relaxation phase left the force-generating states faster (p < 0.05) after PKA-treatment (higher k_LIN), kinetics of the second relaxation phase (k_REL) were not significantly affected by PKA-treatment for both d-hESC-CMs and hvMFs (Figures 4I,J), as it was previously reported for hvMFs (Walker et al., 2011).

In d-hESC-CMs, we could clearly identify the bands of MyHC, α-actinin, desmin, actin, TnT, α and β isoforms of Tm, TnI, atrial (MLC-1a, MLC-2a) and ventricular (MLC-1v, MLC-2v) isoforms of the MLC-1 and MLC-2 proteins (Figure 6A, lane 1), and compared them to those in atrial (lane 2), ventricular (lane 3), and skeletal (lane 4, M. gastrocnemius) adult human muscles.

In d-hESC-CMs and in the homogenized ventricular tissue from which hvMFs were prepared for functional investigations, αMyHC isoform was below the detection limit (Figure 6B). MLC-1a (~84%; Figure 6A) and MLC-2a (81.5 ± 2.1%, n = 3; Figure 6A, Figure S4D) were the predominant protein isoforms expressed in d-hESC-CMs, similar to the atrial sample (~89% for MLC-1a, ~95% for MLC-2a; Figure 6A), while MLC-2v was 18.2 ± 2.5% (n = 3) (Figure 6A, Figure S4D).

Overall, phospho-staining (ProQ Diamond; Figure 6H) revealed that the following sarcomeric proteins were phosphorylated in d-hESC-CMs (Figure 6H, lane 1; + PKA): cMyBP-C (inset 1; ± PKA), desmin, cTnT (inset 2; ± PKA), αTm, MLC-1a, and MLC-2a. For desmin, contributions of cytoskeletal desmin in d-hESC-CMs to the detected band should be considered, because the cytoskeletal network most likely was not completely removed from detergent-treated d-hESC-CMs. Phospho-stained bands corresponding to the phosphorylated cTnT and αTm isoforms were present in all samples [Figure 6H, lane 1 (+PKA), lanes 2,3 (−PKA)]. In adult ventricular and atrial samples, the phosphorylated form of cTnI was clearly visible. Yet, in d-hESC-CMs (±PKA) this band was missing, likely because the cTnI isoform was below detection limit. In d-hESC-CMs, we observed a weak band corresponding to the phosphorylated MLC-1a, which was absent or under the detection limit in adult atrial and ventricular samples (Figure 6H). MLC-2a isoform was also phosphorylated in d-hESC-CMs and more intense than the MLC-1a isoform.

In conclusion, d-hESC-CMs contain sarcomeric protein isoforms found not only in human adult ventricular samples, but also in the atrial and skeletal muscles of the adult human.

The significantly lower passive force (F_pass) for d-hESC-CMs vs. hvMFs (Figure S1), could be due to the presence of the more compliant and longer fetal-N2BA titin isoform (Kruger and Linke, 2009). This is supported by other studies which showed that the fetal-N2BA isoform predominated in hiPSC-CMs (Hinson et al., 2015). F_pass for hvMFs was slightly, but significantly (p < 0.05) diminished following PKA-treatment, while for d-hESC-CMs, PKA had no effect (Figure S1). It was previously shown that adult donor ventricular tissue contains the shorter compliant N2BA and the stiffer N2B titin isoforms in the ratio N2BA:N2B~0.5 and only N2B is target for PKA-mediated phosphorylation (Kruger and Linke, 2009). To determine F_ACT we subtracted F_pass from the recorded active force.

We found that maximum isometric force was significantly smaller for d-hESC-CMs (~42 kPa) than for hvMFs (~94 kPa), presumably due to less well-aligned myofibrils within d-hESC-CMs compared to more compact hvMFs-bundles (e.g., Figures 1A, 5D), but differences in sarcomeric protein isoform composition (Figure 6, Figure S4) could also contribute to the lower isometric force.

Upon sudden increase of [Ca²⁺] from relaxation (>pCa 8) to pCa 4.18, force raised mono-exponentially without lag (Figure 1C) and with very similar k_ACT in both d-hESC-CMs and hvMFs. This suggests that myofibrils within d-hESC-CMs, as previously shown for hvMFs (Stehle et al., 2002b; Poggesi et al., 2005), rapidly equilibrate with the surrounding Ca²⁺-defined solution and that the measured kinetics are not rate-limited by diffusional events. Because k_ACT in both types of myofibrils is similar to k_TR, kinetics of the force rise upon Ca²⁺-activation of d-hESC-CMs, as shown before for hvMFs (Stehle et al., 2002b), seem to be primarily governed by cycling cross-bridges rather than by the much faster Ca²⁺-controlled switch on-and-off of the regulatory proteins of the thin filaments. k_TR for d-hESC-CMs and hvMFs was very similar to k_TR previously determined for hvMFs (Stehle et al., 2002b; Piroddi et al., 2007). This suggests that cross-bridge cycling kinetics at full calcium-activation seems not to be affected by the observed differences in thin filaments regulatory protein- and myosin light chain-isoforms.

Cardiomyocytes in the living heart and in cell culture are operating at submaximal intracellular [Ca²⁺]. At intermediate [Ca²⁺], development-dependent isoform differences of some sarcomeric proteins may influence the steady-state force and cross-bridge cycling kinetics (Metzger et al., 2003; Siedner et al., 2003; Racca et al., 2016).

Ventricular myofibrils and cardiomyocytes expressing the ssTnI isoform exhibit an increased Ca²⁺-sensitivity of force compared to those containing cTnI, and even a partial expression of ssTnI, co-expressed with cTnI, seemed to have a dominant Ca²⁺-sensitizing effect (Metzger et al., 1994, 2003; Siedner et al., 2003; Elhamine et al., 2016; Racca et al., 2016). Therefore, it is very likely that the ssTnI, solely detected in the d-hESC-CMs, contributes to the observed higher Ca²⁺-sensitivity of force when compared to adult hvMFs (Figures 2A,C) containing only cTnI (Figure 6E, Figure S4C). In addition, cTnI was phosphorylated in hvMFs (Figure 6H), which reduces Ca²⁺-sensitivity and thus, this may add to the large difference in Ca²⁺-sensitivity of force between d-hESC-CMs and hvMFs (ΔpCa₅₀ = +0.24) (Figures 2A,B). ssTnI does not have the PKA-dependent phosphorylatable N-terminal extension of cTnI. Therefore, effects of PKA-treatment of d-hESC-CMs on pCa₅₀ cannot be mediated through TnI phosphorylation.

In contrast to the Ca²⁺-sensitizing effect of ssTnI, the ssTnT isoform, detected together with cTnT in d-hESC-CMs (Figure 6F), was shown to reduce the Ca²⁺-sensitivity of force in cardiac fibers (Pinto et al., 2012). However, the effect of ssTnT apparently does not dominate the Ca²⁺-sensitivity of force for d-hESC-CMs compared to hvMFs.

αTm is the predominant tropomyosin isoform expressed in both d-hESC-CMs and hvMFs. βTm and κTm isoforms were shown to exert increasing and decreasing effects on Ca²⁺-sensitivity of force, respectively (Jagatheesan et al., 2010; Rajan et al., 2010). Therefore, the effects may be compensatory in both, d-hESC-CMs and hvMFs. d-hESC-CMs seem to express additionally the γTm isoform (Figure 6G). Co-expression of γTm in cardiac muscle decreases Ca²⁺-sensitivity of force, and importantly, γTm apparently has a functional dominance over the other Tm isoforms in the regulation of striated muscle performance (Pieples et al., 2002; Jagatheesan et al., 2010). Therefore, the presence of γTm in d-hESC-CMs would argue for a decrease of Ca²⁺-sensitivity of force compared to hvMFs.

Isoform differences of TnI, TnT and Tm observed in d-hESC-CMs vs. hvMFs may affect the fast switch-on/-off equilibrium of the thin filament, determining in d-hESC-CMs a particular thin filament status which might be different from that in hvMFs. γTm is more negatively charged and less flexible than the αTm (Jagatheesan et al., 2010) and, together with the ssTnT/cTnT and ssTnI isoforms in d-hESC-CMs, may alter dynamics of the thin filaments to expose their strong binding sites to myosin heads and to the other actin-interacting partners, e.g., N-terminus of MLC-1 (Rarick et al., 1996) and of cMyBP-C (Craig et al., 2014). All these may provide a feedback to the dynamical status of thin filaments, especially at lower [Ca²⁺], as previously proposed considering the cross-bridge-mediated cooperative activation of thin filaments to further recruit new cross-bridges (Campbell, 1997).

MLC-1 interacts in an isoform dependent manner with the lever arm of MyHC (via non-covalent interactions) and with actin (via electrostatic interactions), providing a possible tether for the myosin head between thick and thin filaments. MLC-1a has a higher affinity for the MyHC lever arm and a lower affinity for actin than the MLC-1v isoform (Morano and Haase, 1997; Hernandez et al., 2007). Therefore, the MyHC·MLC-1a may leave the force-generating states faster than MyHC·MLC-1v. It was shown that a higher MLC-1a content by ~20% in human ventricular fibers increases Ca²⁺-sensitivity of force (pCa₅₀) by +0.36 pCa-units (Morano and Haase, 1997; Schaub et al., 1998; Hernandez et al., 2007). MLC-1 can be phosphorylated (Arrell et al., 2001), and this is believed to increase cross-bridge detachment rates, thus shifting the equilibrium of cross-bridges more toward the non-force-generating states, i.e., increasing g_app. Altogether, a predominant expression of MLC-1a in d-hESC-CMs compared to hvMFs (Figures 6A,D, Figure S4B), which seems to be partially phosphorylated (Figure 6H), may contribute to the higher Ca²⁺-sensitivity of force (Figure 2A) and to a distinct modulation of cross-bridge cycling kinetics at submaximal force levels. This is supported here by significantly faster k_LIN (Figures 4E,F) and k_TR (Table 2, Figures 3C, 4D) at pCa₅₀ for d-hESC-CMs compared to hvMFs.

Phosphorylation of MLC-2 was shown to increase k_TR and Ca²⁺-sensitivity of force (van der Velden et al., 2003a,b). In d-hESC-CMs, both MLC-2a/v isoforms co-exist, with MLC-2a being predominant (Figure 6A, Figure S4D) and phosphorylated (Figure 6H). This may contribute to the higher pCa₅₀ (Figure 2) and faster k_TR at pCa₅₀ (Table 2, Figures 3C, 4D) or at a given [Ca²⁺] (e.g., at pCa 5.80 shown in Table 2) for d-hESC-CMs compared to hvMFs.

Altogether, in d-hESC-CMs, functional effects of MLC-1a and ssTnI (both increasing pCa₅₀) seem to dominate the effects of ssTnT and γTm (both decreasing pCa₅₀) at submaximal forces (Table 2), because d-hESC-CMs had a higher Ca²⁺-sensitivity of force compared to hvMFs, when both were either treated (ΔpCa₅₀ = +0.11) or not treated (ΔpCa₅₀ = +0.24) with PKA, respectively (Figures 2A,B). However, possible differences between the phosphorylation status of other sarcomeric proteins than cTnI of d-hESC-CMs compared to hvMFs may also affect Ca²⁺-sensitivity.

cMyBP-C is a target for PKA and it is expressed in both types of myofibrils studied here. Even though phospho-staining analysis showed a weak signal for phosphorylation of cMyBP-C (Figure 6H, inset 1), the force-pCa relation of d-hESC-CMs was clearly shifted to higher [Ca²⁺] upon PKA-treatment (ΔpCa₅₀ = −0.26; Figure 2C). It would be possible that a change in cMyBP-C phosphorylation contributes significantly to this right-shift of the force-pCa relation, because cTnI was undetected in d-hESC-CMs. It was previously shown that in cardiomyocytes lacking cTnI, the phosphorylation of cMyBP-C is able to decrease Ca²⁺-sensitivity of force (Chen et al., 2010). PKA-treatment reduced less, but significantly Ca²⁺-sensitivity of force in hvMFs vs. d-hESC-CMs (Figure 2C), suggesting that hvMFs were already more phosphorylated than d-hESC-CMs.

For both d-hESC-CMs and hvMFs at any given submaximal fractional force level (0 < F_n < 1, F_n = F_ACT/F_ACT,max), PKA-treatment had no significant effect on measured k_TR (Figures 3A,B, 4D) and thus, not on f_app+g_app (= k_TR). f_app (which depends on [Ca²⁺] modulating the thin filament status) and g_app (which is independent of [Ca²⁺]) are the rate constants (probabilities) of cross-bridges entering and leaving the force-generating states, respectively (Brenner, 1988). At the same given [Ca²⁺], small differences could eventually exist between k_TR values determined before and after PKA-treatment, because f_app is Ca²⁺-sensitive and PKA-treatment decreased Ca²⁺-sensitivity of force. For example, at pCa₅₀ (F_n = 0.5) or at a given [Ca²⁺] (pCa 5.80), k_TR values are shown in Table 2. It was previously reported with adult hvMFs, involving the same investigation method to determine k_TR as in the present study, that a shift in Ca²⁺-sensitivity of force following the PKA-treatment did not significantly affect k_TR at different fractional force levels (Walker et al., 2011). Using the same methodological approach with human donor cardiomyocytes, k_TR at different [Ca²⁺] was not significantly affected by PKA-treatment (van Dijk et al., 2014). In studies with murine ventricular skinned myocardium, which may have lower endogenous phosphorylation levels than human donor samples, PKA-treatment accelerated k_TR and this was attributed mainly to cMyBP-C phosphorylation (Stelzer et al., 2006). It was suggested that the acceleration of k_TR occurred by increasing the rate constants of both cross-bridges detachment and recruitment (Stelzer et al., 2006). In d-hESC-CMs expressing cMyBP-C and lacking cTnI, PKA-treatment enhanced the probability of strained cross-bridges to leave the force-generating states, as the differences (p < 0.01) in k_LIN for d-hESC-CMs ± PKA suggest (Figures 4E,F), but it had no significant impact on less strained cross-bridges, as k_REL values were similar (Figures 4I,J). k_LIN was accelerated by PKA-treatment also in hvMFs, but apparently less than in d-hESC-CMs (Figures 4E,F). Therefore, during mechanically loaded cross-bridge turnover at submaximal F_n, the k_TR, which was essentially unaffected by PKA-treatment in d-hESC-CMs (Figures 3A, 4D), would likely result from a relatively small increase of g_app and decrease of f_app (to keep f_app+g_app = k_TR relatively constant). This could be due to a subtle alteration of thin filaments turning-on/off dynamics in the context of different protein isoform pattern (e.g., lack of cTnI) in d-hESC-CMs compared to hvMFs. However, the potential effects caused by PKA-treatment on g_app (increasing) and f_app (decreasing) could not be accurately predicted (large 95%-CI) from k_TR vs. F_n dependencies determined at submaximal force levels for d-hESC-CMs (Figure 3A), but such predictions can be used to compare d-hESC-CMs with hvMFs (see below Figures 7A,B).

From the curves (Figure 3) fitting the k_TR vs. F_n data, the apparent rate constants g_app at F_n = 0 and f_app,max at F_n = 1 can be predicted (Figure 7A) (Brenner, 1988). In d-hESC-CMs, f_app,max and g_app are essentially equal, different from the hvMFs where we calculated 2 to 3-fold larger values for f_app,max compared to g_app (Figure 7A). Such a difference between f_app,max and g_app, and even larger ones, were also previously shown for hvMFs (Stehle et al., 2002b; Poggesi et al., 2005; Piroddi et al., 2007). Based on these calculations, the decrease in F_ACT,max generated by d-hESC-CMs (~42 kPa) vs. hvMFs (~94 kPa), partly results from a decreased occupancy of force-generating cross-bridge states determined by a reduced duty-ratio f_app,max/(f_app,max+g_app) (Brenner, 1988), because f_app,max is lower and g_app is higher in d-hESC-CMs compared to corresponding parameters in hvMFs. Yet, if F_ACT,max is estimated considering only the duty-ratio from the predicted values of f_app,max and g_app (Figure 7A), F_ACT,max for d-hESC-CMs would be ~67% of F_ACT,max for hvMFs, while the measured force of d-hESC-CMs (42 kPa) was ~45% of the force generated by hvMFs (94 kPa). Therefore, other factors may contribute to the estimated differences, for instance, a less ordered and less compact sarcomere alignment within d-hESC-CMs than that observed in hvMFs-bundles (Figures 1A, 5D) and in healthy cardiomyocytes (Kraft et al., 2013; van Dijk et al., 2014). In developing hESC-CMs, structural maturation of sarcomeres could play an important role for force generation, as previously described in human fetal cardiac muscles (Racca et al., 2016).

While at full Ca²⁺-activation k_TR is similar for both contractile systems, at submaximal force levels (0 < F_n < 1), k_TR was significantly faster for d-hESC-CMs than for hvMFs (Figures 3C, 4D). This is mostly due to a higher g_app in d-hESC-CMs than in hvMFs (Figure 7A). Higher g_app of d-hESC-CMs vs. hvMFs may determine an increase of the tension cost (= ATPase/force) that is proportional to g_app (Brenner, 1988). At lower [Ca²⁺], f_app/g_app may modulate contractile function in addition to modulation through changes within the regulatory proteins (Brenner, 1988). Predicted curves fitting k_TR vs. F_n data (Figure 3C) and g_app parameters (Figure 7A) were further used to calculate f_app and then f_app was plotted against F_n (Figure 7B). Independent on g_app, f_app decreases monotonically with decreasing F_n (Figure 7B). Even if predicted f_app vs. F_n curves seem to be similar below F_n~0.75 for both contractile systems (Figure 7B), each fractional force level F_n is reached at lower [Ca²⁺] in d-hESC-CMs than in hvMFs, because d-hESC-CMs are more Ca²⁺-sensitive than hvMFs (Figure 2). For example, at a given [Ca²⁺] of pCa 5.8, d-hESC-CMs generated 75% of maximum force F_ACT,max (Table 2) and predicted f_app = 0.22 s⁻¹ (Figure 7B), while hvMFs generated 50% of their maximum force F_ACT,max (Table 2) and predicted f_app = 0.10 s⁻¹ (Figure 7B). Independent on [Ca²⁺], predicted g_app was 0.37 s⁻¹ for d-hESC-CMs and 0.18 s⁻¹ for hvMFs (Figures 7A,B). Therefore, k_TR (= f_app+g_app) is larger for d-hESC-CMs (0.59 s⁻¹) than for hvMFs (0.28 s⁻¹) at pCa 5.8.

The aspects discussed here resulted solely from the comparison of hESC-CMs [differentiated and cultivated as previously described (Kempf et al., 2014; Weber et al., 2016)] with hvMFs (isolated from adult human ventricles). Variability in the outcomes cannot be excluded if d-hPSC-CMs would be used which were differentiated in another way and subjected to different maturation conditions.