Contractility of striated muscle tissue increases with environmental stiffness according to a power-law relationship

Micro-tissues as a model for structural and functional maturation of skeletal muscle

To study the mechanoresponsiveness of skeletal muscle to environmental stiffness during maturation, we generated muscle micro-tissues that formed between two flexible pillars with adjustable spring constant. In total, 5000 C2C12 skeletal muscle cells were suspended in 6 µl of unpolymerized collagen-I/Matrigel solution and added to polydimethylsiloxane (PDMS) chambers containing two flexible pillars (0.5 mm diameter, placed 2 mm apart) (Fig. 1A). Upon matrix polymerization, the cells adhered to the matrix fibers and exerted contractile forces, resulting in matrix remodeling and the formation of a micro-tissue within 24 h (Fig. 1B, Supplementary Fig. 1, Supplementary Fig. 2). The height at which the micro-tissues attach between the pillars was tuned by forming a first layer of cell-free matrix at the bottom of the PDMS chamber with a defined volume and allowing it to polymerize for one hour prior to cell seeding in a second collagen layer on top of the bottom layer. After 24 h, we replaced the culture medium with a differentiation medium, supplemented with 0.5% horse serum and 0.5% ITS (insulin, transferrin, and selenium), to initiate myocyte differentiation.

A Schematic of the experimental protocol for micro-tissues fabrication. In a two-step process, we pipette a layer of cells mixed in an unpolymerized collagen/Matrigel matrix into the PDMS well on top of a pre-polymerized bottom layer without cells. Within 24 h, micro-tissues form in a cell culture medium containing 15% FCS, and we subsequently switch to a differentiation medium containing 0.5% HS and 0.5% ITS for the remainder of the experiment. B Bright-field images of a micro-tissue over several days in culture. Scale bar: 500 µm. C Maximum intensity projections of confocal image stacks of micro-tissues fixed at 0, 1, 3, and 7 days after switching to differentiation medium. Tissues are stained for α-actinin, desmin, and nuclei (DRAQ5). Excitation intensities and detector gains were the same for all samples. The gradual increase in α-actinin and desmin intensities over time demonstrate maturation of the myocytes in culture. Scale bar: 200 µm. D Maximum intensity projections of confocal image stacks over the entire length of a micro-tissue (fixed on day 7) stained as before. The close-up images show parallel orientation of the myocytes along the axis connecting the pillars. Scale bar: 200 µm. Scale bar in close-up: 100 µm.

Over the following days, the myocytes started to exert measurable forces on the pillars, causing further matrix compaction and fiber alignment between the pillars (Fig. 1A, B). Over the course of seven days of differentiation, the myocytes elongated and aligned between the pillars. In addition, they showed increasing expression levels of the muscle differentiation markers α-actinin and the intermediate filament desmin, as revealed by immunofluorescence staining (Fig. 1C, D).

Already 1 day after cell seeding, the collective contractile forces exerted by the myocytes caused the micro-tissues to pull on the pillars, resulting in a pillar deflection that was visible with bright-field imaging. After 4 days of differentiation culture, the muscle tissues started to respond to electrical pacing, as seen by a measurable pillar deflection with every pulse. The force acting on the pillars was then calculated according to Hooke’s law (Eq. (1)) from the pillar deflection (Fig. 2a, b) multiplied by the pillar spring constant (Fig. 2c). We refer to the continuous baseline force as the static force, and we refer to the additional force above the static force that is observed in response to electrical stimulation as the active force (Fig. 2d).

a Bright-field image of a C2C12 micro-tissue focused at the top of the PDMS pillars. The orange arrowhead indicates a high-contrast point that is used for particle image velocimetry analysis. Scale bar: 200 µm. b Kymograph of the region indicated in (a) in response to active tissue contraction. Scale bars: 10 µm and 250 ms. c Calibration curves to determine the spring constants of the PDMS pillar. The spring constants were measured from the force-deflection relationship of pillars at different heights and for PDMS with different crosslinker-to-base ratios. Dots represent slopes of individual force-deflection measurements, and lines represent fits to the data points using the Euler–Bernoulli beam equation. d Static and active contractility of the micro-tissue shown in (a). The stimulation frequency was 1 Hz. e Static contractile forces of C2C12 micro-tissues after 7 days in culture. One group of micro-tissues was prepared as usual (i.e., using ITS differentiation medium), and one group was kept in culture medium for the entire 7 days (instead of switching to ITS medium after 24 h) so that the myocytes did not differentiate. We measured the static contractile forces under control conditions (Ctrl; 1 vol% dimethylsulfoxid), after adding either 10 µM cytochalasin-D (cyto-D), or 100 µM blebbistatin (Blebb) to the medium for 1 h, or after adding Triton X-100 (Tx-100) to the medium for 10 min; n.s., P ≥ 0.05; ★★★, P < 0.001. f Same as (e) for active contractile forces. g Contrast-enhanced brightfield images of exemplary micro-tissues before and after treatment with cyto-D. One micro-tissue was cultured in ITS medium and the other in culture medium before drug treatment. Scale bar: 100 µm.

The static force of C2C12 micro-tissues increased over time in differentiation culture. In PDMS devices of intermediate stiffness (pillar spring constant ~6 N/m), the static force reached 76 µN on average after 1 week, and the active force was 7.5 µN (Fig. 2e, f). By contrast, undifferentiated C2C12 micro-tissues that were continuously exposed to culture medium but not differentiation medium showed higher static forces (166 µN on average), but almost no measurable active forces (0.9 µN on average).

To further explore the origins of static and active contractility, we measured forces after treatment with inhibitors for different components of the contractile machinery (Fig. 2e, f). Addition of 100 µM of the myosin II inhibitor blebbistatin for one hour completely impaired active contractility in all micro-tissues (active forces were less than 0.1 µN, resulting solely from noise in the pillar deflection measurements). Cell permeabilization with Triton X-100 at a concentration of 0.5% for ten minutes also completely impaired active contractility, confirming that an intact sarcolemma is essential for initiating contractions through electrical stimulation. By contrast, static forces were not completely abolished but strongly reduced following blebbistatin and Triton X-100 treatment. In undifferentiated tissues, static forces fell from 166 µN to 27 µN (blebbistatin) and 24 µN (Triton X-100), and in differentiated tissues, static forces fell from 76 µN to 14 µN (blebbistatin) and 16 µN (Triton X-100). These findings are consistent with the requirement of myosin II activity for contractile force generation in non-muscle cells¹⁷.

Since sarcomeric actin filaments are stably capped at their barbed ends in the Z-disks of myofibers^18,19, we hypothesized that the active forces in differentiated C2C12 muscle-micro-tissues are less susceptible to barbed-end depolymerization than the static forces. To test this, we treated differentiated and undifferentiated micro-tissues with high doses of the F-actin barbed-end depolymerizing fungal toxin cytochalasin D (cyto-D). Treatment with 10 µM cyto-D for one hour reduced the static forces in differentiated and undifferentiated micro-tissues to similar levels as seen after treatment with blebbistatin or Triton-X (26 µN in differentiated and 32 µN in undifferentiated micro-tissues). By contrast, cyto-D treatment had no influence on active contractile forces in differentiated micro-tissues (7.1 µN). In addition, brightfield imaging revealed that after treatment with cyto-D, undifferentiated micro-tissues increased in width, and many cells appeared rounded (Fig. 2g), whereas tissue widening and cell rounding were less pronounced in differentiated micro-tissues. The observation that cyto-D has an inhibitory effect only on static but not on active forces strongly supports our hypothesis that active forces are generated predominantly by differentiated cells where the barbed ends of sarcomeric actin filaments are stably capped, rendering the cells insensitive to cyto-D treatment, while cyto-D can initiate barbed-end depolymerization of the non-sarcomeric F-actin in undifferentiated myocyte precursors, resulting in reduced static forces. The presence of appreciable static forces after treatments with cyto-D, blebbistatin, and Triton X-100 furthermore suggests that a part of the static force originates from stored mechanical tension within remodeled collagen fibers.

Next, we sought to understand the role of integrin-type adhesions to the ECM in the maturation of the micro-tissues. The differentiation of myoblasts into myotubes requires the engagement of integrins or the dystrophin-glycoprotein complex to laminin, a critical adhesive component of the extracellular matrix surrounding skeletal myofibers and cardiac myocytes^{20,21,22,23,24,25,26,27}. Since laminin is a major component of Matrigel²⁸, we tested whether the presence of Matrigel was required for proper C2C12-micro-tissue formation and contraction. C2C12 cells mixed into pure collagen-I (COL), without the addition of Matrigel, did not develop uniformly-compact micro-tissues but over the course of 7 days only formed loose tissues interspersed with denser aggregates (Fig. 3A). Importantly, these micro-tissues were unable to generate active contractile forces (Fig. 3B). These results suggest that laminin promoted C2C12-myoblast differentiation in micro-tissues, in line with our previous results that active contractility requires differentiated cells.

A Bright-field images of representative micro-tissues for all conditions. Scale bar: 500 µm. B Active contractile forces of C2C12 micro-tissues prepared with ECMs of different compositions: collagen only (COL), collagen and 1 mg/ml Matrigel (COL + MA), collagen and 1 mg/ml growth factor-reduced Matrigel (COL + MA_-gf), collagen and 100 µg/ml laminin (COL + LAM₁₀₀), and collagen and 200 µg/ml laminin (COL + LAM₂₀₀); n.s., P ≥ 0.05; ★★★, P < 0.001.

To test whether the impaired muscle differentiation in the absence of Matrigel was indeed due to a lack of laminin, or due to the lack of growth factors that are also part of Matrigel, we compared C2C12 micro-tissues with collagen-I containing 1 mg/ml of either regular (COL + MA) or growth factor-reduced Matrigel (COL + MA_-gf), or 100 µg/ml (COL + LAM₁₀₀), or 200 µg/ml (COL + LAM₂₀₀) pure laminin isolated from Engelbreth–Holm–Swarm murine sarcoma basement membrane. Micro-tissues formed in the presence of growth factor-reduced Matrigel resembled control COL + MA tissues in terms of morphology and contractility (COL + MA = 25.0 µN; COL + MA_-gf = 29.9 µN; two-tailed t-test: P = 0.42). Similar to pure COL micro-tissues, COL + LAM₁₀₀ micro-tissues showed a low degree of compaction, presence of aggregates, and no active forces. COL + LAM₂₀₀ micro-tissues, however, compacted better and developed low levels of active contractile forces (3.8 µN), indicating that even moderate levels of pure laminin were able to improve micro-tissue formation and contraction. These data show that laminin-1 is important for myotube differentiation, while matrix-bound growth factors present in Matrigel are not. However, addition of 200 µg/ml laminin-1 was not sufficient to restore the full contractility. It is possible that higher laminin-1 concentrations similar to those found in Matrigel (approx. 600 µg/ml²⁸), are required, or that other Matrigel components have also contributed to myotube differentiation.

Together, these data show that C2C12-based micro-tissues recapitulate key features of skeletal muscle, including laminin 1-dependent myoblast elongation and differentiation, electrical excitability, and contractile force generation, suggesting that they can be used as a functional in vitro model for striated muscle tissue.

Neonatal rat cardiomyocytes form micro-tissues with intact electrical coupling

Next, we investigated whether mechanoadaptive behavior is specific to skeletal muscle or applies broadly to striated muscle tissue. For this, we studied neonatal rat ventricular cardiomyocytes (NRVC) that have not yet undergone postnatal terminal differentiation and are lacking key features of adult cardiomyocytes such as a fully organized sarcomere apparatus and a rod-like cell shape. We first tested whether NRVCs were able to form micro-tissue in our system and whether cells and micro-tissues show features of adult cardiomyocytes and myocardial tissue, respectively. Over 3 days in culture, NRVCs had fully compacted the ECM into micro-tissues (Fig. 4A). NRVCs best formed micro-tissues using a lower collagen concentration (0.3 mg/ml) compared to C2C12 cells (0.6 mg/ml). Tissue formation did not require the addition of Matrigel, possibly because NRVCs were already partly differentiated in vivo.

A Bright-field images of a representative micro-tissue on day 2 and 3 after cell seeding. Scale bar: 200 µm. B Maximum projections of confocal image stacks of a micro-tissue fixed 5 days after cell seeding and stained for α-actinin (red) and nuclei (DRAQ5, blue). Scale bar: 20 µm; scale bar inset: 10 µm. C Static and active contractility of a cardiac micro-tissue at different stimulation frequencies. In the insets, the black dashed lines represent the static forces and the orange dashed lines represent the minimum total force between pacing pulses. Percentages indicate the average minimum active force between pacing pulses, relative to the average maximum active force.

Notably, confocal imaging of α-actinin revealed an elongated morphology of NRVCs and a homogenous cross-striation pattern reminiscent of adult myocardial tissue in vivo (Fig. 4B). In addition, NRVC micro-tissues displayed spontaneous as well as electrically induced active contractions already three days after cell seeding. The static forces of NRVC micro-tissues were one order of magnitude lower than those of C2C12 micro-tissues (typically 1–10 µN), while the active forces were comparable (typically 20 µN). The active contraction of NRVC micro-tissues followed the frequency of the electrical stimulation up to about 4 Hz (Fig. 4C). At higher frequencies, single force twitches did not fuse to a tetanic force as seen in skeletal muscle micro-tissues²⁹, but still showed discernible pulses, closely mimicking the contraction dynamics of cardiac tissue in vivo. Consequently, the maximum active forces at higher stimulation frequencies (2, 4, 8, 10 Hz) remained unchanged compared to single pulse stimulation (0.5 and 1 Hz).

These data show that NRVC micro-tissues replicate essential features of terminally differentiated myocardial tissue, such as cardiomyocyte elongation and cross-striation as well as electrically inducible contraction, suggesting that, similar to our C2C12 micro-tissues, they are well suited for in vitro studies of cardiac muscle tissue.

The active contractility of skeletal and cardiac muscle micro-tissues is mechanoadaptive

A key feature of our system is that the environmental stiffness seen by the micro-tissues can be tuned over a wide range, (i) by varying the volume of the bottom layer during the fabrication process so that the micro-tissues form at different heights h between the pillars (Fig. 5a), and (ii) by varying the PDMS crosslinker-to-base ratio, which changes the Young’s modulus E of the pillars.

A Schematic of the relationship between bottom layer volume and tissue height. After 24 h of initial compaction, the micro-tissues are attached to the PDMS pillars approximately at the height of the bottom/cell layer interface. Since the effective spring constant k of the pillars depends on the micro-tissue height, a high/intermediate/low bottom layer (9/6/3 µl) results in a low/moderate/high environmental stiffness. B Scatter plots of active and static forces of C2C12 micro-tissues versus k measured on days 4–7 after initialization of differentiation. Dashed lines represent power-law fits. C Power-law coefficients for the same fits (with days 8 and 9 added). Bar plots show mean ± SE as determined by bootstrapping. D Scatter plot of active deflections over k for day 5 with power-law fit. E Same as (D) but for static deflection. F Scatter plot of times-to-peak (i.e., the time at which the micro-tissues reach maximum deflection after electrical stimulation) versus k at day 5.

To investigate the influence of the mechanical environment and of culture time on the contractile performance, we first fabricated C2C12 micro-tissues with different E and h and measured the active and static contractile forces from day 4 to 9 after switching to differentiation medium (Fig. 5B). We found that the active forces F increased with environmental stiffness k, and that the relationship can be well approximated by a simple power-law relationship of the form \(F\left(k\right)=a\times {(k/{k}_{0})}^{b}\), where the factor a is a measure of contractile force at a nominal spring constant of k₀ = 1 N/m, and the power-law coefficient b is a measure of mechanoadaptation (Fig. 5C). A power-law exponent of zero corresponds to a tissue that does not adapt to environmental stiffness, and an exponent of unity corresponds to a tissue that increases its contractile force linearly with increasing environmental stiffness.

On day 4—the first day with measurable, active contraction—active forces increased with k according to a power-law exponent b = 0.57 ± 0.04 (mean ± SE as determined by bootstrapping; R² = 0.53, ρ = 0.82), i.e., roughly with the square root of environmental stiffness (Fig. 5B). Over the next two days in culture, the degree of mechanoadaptation increased further, reaching a maximum power-law coefficient of 1.03 ± 0.05 on day 6 (R² = 0.59), hence contractility increased nearly linearly with environmental stiffness. By contrast, the static contractile force increased only weakly with k and reached a maximum power-law coefficient of 0.17 ± 0.04 (R² = 0.18) on day 5 (Fig. 5B). These data demonstrate a pronounced mechanoresponse of C2C12 micro-tissues that stems from differentiated myotubes, but not myoblasts.

We next explored how muscle shortening and hence pillar deflection changed with the spring constant, since the active force of striated muscle may decrease with increasing amount of shortening as the internal resistance against filament sliding increases³⁰. To this end, we correlated pillar deflection for active and static forces with the corresponding spring constant. Consistent with the absence of a clear mechanoresponse of static force generation, the pillar deflection induced by static forces decreased over several orders of magnitude with increasing pillar stiffness. By contrast, the pillar deflection due to active forces was nearly constant, regardless of pillar stiffness. For example, on day 5, the static pillar deflection was strongly correlated with k (R² = 0.76) and had a power-law coefficient of −0.85 (Fig. 5E), while the active pillar deflection was only weakly correlated with k (R² = 0.18) and had a power-law coefficient of −0.20 (Fig. 5D). Together with the observation that the maximum amount of shortening was only 2% of the total tissue length, this finding suggests that the active forces of C2C12 micro-tissues in a softer microenvironment are not limited by filament sliding constraints or by a limited capacity to shorten. Consistent with this, the time required for the micro-tissues to reach maximum active contraction (time-to-peak) was independent of the spring constant (Fig. 5F).

We next tested whether cardiomyocyte micro-tissues prepared from NRVCs were also adapting to pillar stiffness. We observed a less pronounced but still strong mechanoadaptation response (Fig. 6A, B). These micro-tissues first exhibited active contraction at day 3 after initial cell seeding, with a power-law coefficient of the active forces versus k relationship of 0.48 ± 0.04 (R² = 0.59, ρ = 0.73). Thereafter, the degree of mechanoadaptation decreased slightly with culture time, and the power-law coefficient was between 0.35 and 0.39 on days 4–6 and decreased to 0.25 on day 7. These data demonstrate that the mechanoadaptation is not limited to skeletal muscle micro-tissues, suggesting that the underlying mechanisms may be general to different types of striated muscle tissues.

A Scatter plots of NRVC micro-tissue active forces versus k measured on days 3–6 after cell seeding. Dashed lines represent power-law fits. B Power-law coefficients for the same fits (with day 7 added). Shown are means ± SE derived from bootstrapping.

Only the contractile force but not the stiffness of muscle micro-tissues is mechanoadaptive

We next explored whether the stiffness of muscle micro-tissues adapts to the pillar stiffness. To measure the stiffness of micro-tissues, we stretched or compressed the PDMS device (Fig. 7A) and analyzed the relationship between the external force F, tissue shortening δ_tissue, and pillar deflection δ_pillar, as described in “Methods”. Briefly, we assume a serial arrangement of three springs representing the micro-tissue and two pillars with the spring constants k_tissue and k_pillar, respectively (Fig. 7B). We measure δ_tissue and δ_pillar using bright-field imaging and determine k_pillar from calibration measurements (Fig. 7C). The tissue stiffness k_tissue can then be calculated using Eq. (3).

A Schematic of a pre-stretched PDMS device. B Three-spring-model of a micro-tissues between two pillars during stretch/compression. Note that during uniaxial stretch/compression, one end of the serial spring assembly is fixed while the external force acts on the other end. C Scatter plot of tissue length versus total length (tissue length + pillar deflections) and tissue length versus deflection of one pillar. Dashed Lines represent linear fits. D Scatter plot of the calculated values of k_tissue versus the corresponding values of k_pillar. E Three-springs-model of a micro-tissues between two pillars during active/static contraction and equivalent circuit of one half of this arrangement (from one wall to the center of the micro-tissues). F Line plot of the analytical expression F_pillar/F versus k_pillar/k_tissue. Dashed lines represent power-law fits of F_pillar/F in different decades of k_pillar/k_tissue. The line with vertical endpoints represents the stiffness range of the PDMS pillars used in previous experiments.

We performed these experiments with C2C12 micro-tissues for varying pillar stiffness on days 4 and 6 after switching to differentiation medium (the days on which we had measured the lowest and highest degree of mechanoadaptation, respectively). On both days, k_tissue was independent of k_pillar, demonstrating that the stiffness of the micro-tissues is not mechanoadaptive. Furthermore, k_tissue was not significantly different between days (two-tailed t-test: P = 0.750), with an average spring constant of all measured micro-tissues of 0.36 ± 0.04 N/m (mean ± SE; Fig. 7D), corresponding to a Young’s modulus of 9.2 kPa (for an effective tissue radius of 150 µm).

The effective spring constant of a C2C12 micro-tissue is on the same order of magnitude as the spring constant of the softest PDMS pillars used in this study. This creates a potential source of misinterpretation of mechanoadaptation responses, as the deflection of a soft pillar (which corresponds to the shortening of the tissue) may be so large that the internal tissue stiffness causes a substantial counter-force upon tissue compression that limits further shortening. To distinguish mechanoadaptation of the tissue from such passive force limitation, we calculated how tissue stiffness influences the relationship between contractile force and environmental stiffness for a tissue that is not mechanoadaptive (Fig. 7E, F, see “Methods”). Accordingly, for a non-mechanoadaptive tissue generating a contractile force F, the force measured from the pillar deflection (F_pillar) increases approximately linearly with environmental stiffness k_pillar if k_tissue ≫ k_pillar. For k_tissue ≪ k_pillar, the measured F_pillar approaches the true tissue force F.

The pillar stiffness was between 2-fold and 150-fold higher than tissue stiffness, as indicated by the horizontal bar in Fig. 7F, hence k_tissue ≪ k_pillar. In this case, there is only a weak dependency between pillar force and tissue stiffness for non-mechanoadaptive model tissues, with power-law exponents on the order of 0.1 and lower. Therefore, the strong power-law dependency of active forces with increasing pillar stiffness that we see in our measurements, is a signature of true mechanoadaptation and cannot be explained by a tissue compression that limits shortening, even on days where the power-law exponent was lowest (day 4 for skeletal muscle tissues, day 7 for cardiac muscle tissues) (Supplementary Fig. 3).

Mechanoadaptation of C2C12 micro-tissues does not require maximum force magnitude

One strategy for microtissues to probe the stiffness of their environment is to exert force and quantify the resulting change in length. We therefore investigated whether the mechanoadaptation of micro-tissues is affected by the maximum force magnitude that they can generate. Previously, we have identified a role of the focal adhesion protein β-parvin in regulating myocyte shape, sarcomere assembly, and contractility in cardiac cells in response to mechanical load¹⁶. Micro-tissues prepared from Parvb-siRNA-treated NRVCs exhibited reduced active forces compared to micro-tissues prepared from control-siRNA-treated NRVCs¹⁶. Since β-parvin is also highly expressed in skeletal muscle¹⁶, we tested whether β-parvin knockdown affected the contractility and mechanoadaptation of C2C12 micro-tissues.

To this end, we fabricated C2C12 micro-tissues between PDMS pillars with different spring constants as described above, using cells treated with either control- (siControl) or Parvb- (siParvb) siRNA 24 h prior to tissue formation. SiControl micro-tissues were morphologically similar to untreated C2C12 micro-tissues (Fig. 8A) and started to actively contract on day 4. By contrast, siParvb micro-tissues were not able to fully compact the collagen matrix. Consistent with our observations in NRVC micro-tissues¹⁶, absolute active forces were strongly reduced by β-parvin knockdown on day 5, but recovered to the level of siControl micro-tissues by day 7 (Fig. 8B, C) likely due to siRNA-degradation over time. The static forces were also significantly decreased by β-parvin knockdown but did not recover to the levels of siControl micro-tissues after day 7. The distribution of pillar spring constants in siParvb and siControl experiments was comparable (Supplementary Fig. 4).

A Bright-field images of micro-tissues prepared from control- and Parvb-siRNA-treated C2C12 cells. Images were taken 5 and 7 days (D5 and D7) after switching to differentiation medium. Scale bar: 200 µm. Scale bar in close-up: 100 µm. B Scatter plots of active and static forces of control-siRNA and knockdown micro-tissues at D5 and D7. Dashed lines represent power-law fits. C Bar plots of the same data. Bootstrapping; n.s., P ≥ 0.05; ★★, P < 0.01. D Bar plots of the power-law coefficients of fits of active forces and spring constants from 4 to 5 independent experiments on D5–7. Two-tailed t-test; n.s., P ≥ 0.05. E Example time course of normalized active contractile force of a control siRNA-treated micro-tissue (dots) fitted by twofold low-pass filtered (with time constant τ) Delta-Dirac pulses applied at time points when the force exhibits local minima (dotted line). F Bar plots of the time constants τ of active contractions in control- and Parvb-siRNA-treated micro-tissues at D5–7. Two-tailed t-test; n.s., P ≥ 0.05; ★★, P < 0.01.

Strikingly, the mechanoresponsiveness of the active forces at day 5 and 7 were almost unaffected by β-parvin knockdown (Fig. 8B, D). The average power-law coefficient of the active force versus spring constant relationship for siControl micro-tissues was slightly but not significantly higher than that of siParvb micro-tissues for all days (siControl: 0.64–0.71; siParvb: 0.52–0.64, n = 5 independent experiments each with >100 tissues). This indicates that micro-tissues remain fully mechanoresponsive even when the active and static forces are markedly reduced. A plausible explanation is that the expression levels of β-parvin after transient siRNA knockdown recover inhomogeneously over time across the cell population, so that the active tissue force, although it is reduced, is generated by differentiated myotubes that may be fewer in number but are still fully mechanoresponsive.

To test the idea that a temporary β-parvin knockdown delayed muscle differentiation in C2C12 micro-tissues, we first analyzed the time course of micro-tissue contraction. The active force response after a single pulse (twitch force) can be modeled as a Dirac-pulse that is low-pass filtered twice with the same time constant τ, as explained in “Methods”. We find that τ of siControl and siParvb tissues decreased over time after initiating muscle differentiation, consistent with the notion that τ reports on the degree of muscle differentiation (Fig. 8F). The twitch time constant was significantly longer in siParvb micro-tissues compared to siControl micro-tissues (day 5; siControl: 137 ms; siParvb: 161 ms; two-tailed t-test: P < 10⁻¹⁰), but converged to similar levels over time (day 7; siControl: 82 ms; siParvb: 81 ms; two-tailed t-test: P = 0.73). This result supports the hypothesis that β-parvin knockdown suppressed muscle differentiation.

To further test whether β-parvin knockdown delayed muscle differentiation, we differentiated C2C12 cells in 2-D cell culture and repeatedly subjected them to transfection with control- or Parvb-siRNA for a sustained β-parvin knockdown over the entire culture time. Confocal microscopy of the cultures after immunofluorescence-staining for desmin as differentiation marker and phalloidin to label F-actin revealed a significantly reduced desmin/F-actin fluorescence-intensity ratio in siParvb- compared to siControl-cultures (Fig. 9a, b). In addition, the ratio of the nuclear muscle differentiation marker myogenin to the DNA-marker DRAQ5 was considerably reduced in the absence of β-parvin (Supplementary Fig. 5). These data demonstrate that β-parvin is required for the differentiation of C2C12-myoblasts. However, a fraction of myotubes in siParvb-cultures showed regular desmin fluorescence intensity, suggesting that the recovery from β-parvin depletion, and hence the initial β-parvin knockdown, was not homogenous across the culture (Fig. 9a, insets).

a Confocal micrographs from control- and Parvb-siRNA-treated C2C12 cultures in 2-D after immunofluorescence staining for desmin as differentiation marker, phalloidin-staining to label F-actin and DRAQ5-staining to label nuclei. Images were taken 8 days after repeated β-parvin knockdown, and 6 days after switching to differentiation medium, respectively. Note the reduced fraction of desmin-positive myotubes (inserts) in siParvb compared to siControl-samples. Scale bar: 50 µm. b Box plot of desmin/F-actin fluorescence intensity ratio computed from confocal micrographs shown in (a); Paired two-tailed t-test; n = 5 independent biological experiments; ★, P = 0.014. c RT-PCR analysis of the time course of Parvb-mRNA (194 bp) depletion and recovery after knockdown in 3-D cardiac microtissues. Actb (183 bp) was used as positive control.

To characterize the time course of β-parvin knockdown and recovery, we performed RT-PCR analysis of siControl and siParvb micro-tissues on days 0, 2, 4, and 6 after onset of differentiation (Fig. 9c). SiControl micro-tissues displayed a gradual increase in Parvb-mRNA from day 0 to day 6, in line with the requirement of β-parvin for C2C12- myoblast differentiation. By contrast, siParvb micro-tissues showed a complete loss of Parvb-mRNA between days 0 and 2, followed by a recovery of Parvb-mRNA between days 4 and 6 (Fig. 9c), demonstrating that the recovery of siParvb micro-tissues in active force generation between days 5 and 7 (Fig. 8B) coincides with a recovery of Parvb-mRNA expression. Together, these data show that β-parvin is required for C2C12-myoblast differentiation, and that β-parvin knockdown was efficient, but also transient and incomplete with a heterogeneous recovery, suggesting that the lower contractile forces in siParvb micro-tissues after 5 days of differentiation reflects a lower percentage of differentiated muscle fibers that were nonetheless similarly mechanoadaptive as muscle fibers under control conditions.