Effect of mechanical stretching stimulation on maturation of human iPS cell-derived cardiomyocytes co-cultured with human gingival fibroblasts

Abstract

In the realm of regenerative medicine, despite the various techniques available for inducing the differentiation of induced pluripotent stem (iPS) cells into cardiomyocytes, there remains a need to enhance the maturation of the cardiomyocytes. This study aimed to improve the differentiation and subsequent maturation of iPS-derived cardiomyocytes (iPS-CMs) by incorporating mechanical stretching. Human iPS cells were co-cultured with human gingival fibroblasts (HGF) on a polydimethylsiloxane (PDMS) stretch chamber, where mechanical stretching stimulation was applied during the induction of cardiomyocyte differentiation. The maturation of iPS-CMs was assessed using qRT-PCR, immunocytochemistry, transmission electron microscopy, calcium imaging and contractility comparisons. Results indicated significantly elevated gene expression levels of cardiomyocyte markers (cTnT) and the mesodermal marker (Nkx2.5) in the stretch group compared to the control group. Fluorescent immunocytochemical staining revealed the presence of cardiac marker proteins (cTnT and MYL2) in both groups, with higher protein expression in the stretch group. Additionally, structural maturation of iPS-CMs in the stretch group was notably better than in the control group. A significant increase in the contractility and calcium cycle of iPS-CMs was observed in the stretch group. These findings demonstrate that mechanical stretching stimulation enhances the maturation of iPS-CMs co-cultured with HGF.

Introduction

Regenerative medicine leveraging induced pluripotent stem (iPS) cells has emerged as a transformative approach, with clinical applications spanning various domains, such as retinal therapy¹. In the realm of cardiac medicine, iPS cell-derived myocardial sheets and autologous heart transplantation using iPS cells hold promise for addressing end-stage heart failure². Despite advancements in methods for differentiating iPS cells into myocardial tissues and achieving substantial yields³, challenges remain in achieving complete maturation for clinical applications⁴.

Cardiomyocytes constitute approximately 75% of the myocardial tissue volume but represent only 30%−40% of the total cardiac cell population, with the majority being fibroblasts⁵. Numerous studies have been conducted to enhance myocardial maturation by replicating the cardiac environment. Ieda et al. demonstrated that co-culturing mouse cardiomyocytes (CMs) with cardiac fibroblasts (CFs) promotes maturation through cell-to-cell interactions and paracrine signaling⁶. Additionally, co-culturing mouse embryonic stem cell-derived CMs with mouse embryonic fibroblasts has been shown to improve their differentiated phenotype⁷. Additionally, the inclusion of human foreskin fibroblasts, combined with static stretching, has demonstrated improvements in myocardial tissue maturation⁸. Notably, Giacomelli et al. highlighted the synergistic benefits of combining human iPS cell-derived cardiomyocytes with cardiac fibroblasts and endothelial cells to promote structural, electrical, mechanical, and metabolic maturation⁹.

Human gingival fibroblasts (HGF) represent a particularly advantageous co-culture partner for cardiac differentiation due to their accessibility during routine dental procedures and regenerative potential¹⁰. Prior studies have demonstrated that co-culturing iPS cells with HGF improves cardiomyocyte differentiation and enhances contractility. Matsuda et al. demonstrated that co-culture with human gingival fibroblasts (HGF) facilitates differentiation of iPS cells into cardiomyocytes (iPS-CMs) with higher contractility¹¹. Furthermore, mechanical stretching has been shown to promote cardiomyocyte maturation¹². However, a combined approach utilizing HGF and mechanical stretching to improve myocardial tissue maturation remains unexplored.

This study investigates the effects of mechanical stretching stimulation on the maturation of iPS cell-derived cardiomyocytes co-cultured with HGF. Maturation was assessed through gene and protein expression, sarcomere structure analysis, contractility, and calcium handling properties. Our findings highlight the potential of this approach to advance cardiac regenerative medicine by improving the functional and structural maturity of iPS cell-derived myocardial tissues.

Materials and methods

Isolation of HGF

When extracting teeth from a patient undergoing orthodontic treatment at Okayama University Hospital orthodontic clinic, about 2 mm³ of gingival tissue was collected (Fig. 1b). Patients were at least 16 years of age. Patients with periodontitis were excluded for tissue collection. The tissue pieces were then minced in a tissue culture dish with the following solution: 10% fetal bovine serum (FBS: Sigma Aldrich, MO, USA), 1 mol/l-HEPES buffer solution (Nacalai Tesque, Kyoto, Japan), penicillin-streptomycin, and Dulbecco’s Modified Eagle Low Glucose Medium (DMEM; Cat. #: 11885-084, Thermo Fisher Scientific, MA, USA). Finally, the cells were incubated at 37 °C in a 5% CO₂ humidified incubator. HGF that became confluent in the tissue culture dish was periodically subjected to trypsin treatment and subcultured. All experiments were performed with HGF between passages 3 and 10.

Fig. 1.

Protocols for myocardial differentiation of iPS cells co-cultured with HGF and mechanical stretching stimulation system. a Myocardial differentiation protocol using iPS cells co-cultured with HGF. b Protocol for the collection of human gingival tissue and isolation of HGF, HGF cultured on day 11 was shown. c Polydimethylsiloxane (PDMS) stretch chamber.

Culture of iPS cells

Human iPS cells (201B7 cells of dermal fibroblast origin) were purchased from RIKEN (Kyoto, Japan). StemFit AK02N medium (Cat. #: RCAK02N, Ajinomoto, Tokyo, Japan) was used for maintenance culture of iPS cells. A 6-well plate was coated with laminin 511-E8 fragment diluted to 0.5 g/ml in PBS (2 ml per well), and the plate was allowed to stand overnight at 4°C. Then the laminin 511-E8 fragment solution was removed from the culture dish of iPS cells, and the iPS cells were washed with PBS. Subsequently, 800 µl of TrypLE Select (Life Technologies, Carlsbad, CA, USA) was added to the wells and incubated for 7 min in a humidified incubator at 37 °C, 5% CO₂. Next, TrypLE Select was removed, the cells were washed again with PBS, and 1 ml of StemFit AK02N medium containing inhibitor of Rho-associated, coiled-coil containing protein kinase, Y-27632 (10µM) was added. Next, the iPS cells adhered to the bottom of the culture dish were peeled off using a scraper, and seeded at a density of 3.0 × 10⁴ cells/well on a 6-well culture plate coated with laminin 511-E8 fragment. The next day, the medium was removed and replaced with StemFit AK02N medium without Y-27632. The medium was changed on days 1, 4, 5, and 6. All experiments were performed with iPS cells between passages 6 and 10.

Induction of differentiation of iPS cells into myocardial tissue

All samples were cultured using a PDMS (polydimethylsiloxane) stretch chamber (size: 20 × 20 mm, Menicon, Aichi, Japan) to apply mechanical stretching stimulation during the induction of differentiation into cardiomyocytes (Fig. 1a and c). First, Matrigel (Cat. #: 354230, Corning, NY, USA) was diluted to 35.5 µl/ml in DMEM. 1.4 ml/well Matrigel solution was added to wells, and the plate was kept overnight at 4 °C for coating. The next day, 4.9 × 10⁵ cells of iPS cells and 2.1 × 10⁵ cells of HGF were mixed and seeded on the wells. From the day after the seeding, the cells were cultured in Essential 8 medium (Cat. #: A1517001, Thermo Fisher Scientific, MA, USA) for 3 days, and the medium was changed every day. Subsequently, differentiation induction of iPS cells was started using PSC cardiomyocytes Differentiation Kits (Cat. #: A2921201, Thermo Fisher Scientific, MA, USA) according to the manufacturer’s protocol. Briefly, the medium was removed and cardiomyocytes differentiation medium A was added. Two days later, the medium was removed, and cardiomyocytes differentiation medium B was added. After incubation for 2 days, the medium was removed and cardiomyocytes maintenance medium was added. The cardiomyocytes maintenance medium was changed every other day thereafter. Cardiomyocyte differentiation was initiated on Day 3 and completed by Day 7, followed by maintenance in cardiomyocyte maintenance medium. Therefore, on Day 15 — corresponding to the cardiomyocyte maturation phase — mechanical stretching stimulation was applied at a 5% elongation and 0.5 Hz frequency for 72 h using ShellPa Pro (Menicon, Aichi, Japan) (Fig. 1a). A sample cultured in a stationary state without applying a stretching stimulus was used as a control group.

Quantitative RT-PCR

RNA was extracted using the High Pure RNA Isolation Kit (Roche, IN, USA) and reverse transcribed into cDNA using the Verso cDNA synthesis kit (Thermo Fisher Scientific, MA, USA). The target genes were cardiac marker cTnT, mesodermal marker Nkx2.5, pluripotent marker Sox2 and Oct4, and the endogenous control was 18 S rRNA. The primers of these genes were mixed with SYBR green reagent (Life Technologies, Warrington, UK) and quantitative RT-PCR was performed to quantify the expression level of the target genes. Table 1 shows the nucleotide sequences of the primers. The expression level of the target genes was normalized by the expression level of 18 S rRNA, and calculated by the ΔΔCt method. All measurements were performed in triplicate for each target gene in three independent samples.

Table 1.

Primers used for quantitative RT-PCR.

Primer	Direction	Sequence	Size (bp)	Ref.
cTnT	forward reverse	5’-GGCAGCGGAAGAGGATGCTGAA-3’ 5’-GAGGCACCAAGTTGGGCATGAACGA-3’	150	13
Nkx2.5	forward reverse	5’-TTCCCGCCGCCCCCGCCTTCTAT-3’ 5’-CGCTCCGCGTTGTCCGCCTCTGT-3’	139	14
Oct4	forward reverse	5’-CAGCGACTATGCACAACGAGA-3’ 5’-GCCCAGAGTGGTGACGGA-3’	196	15
Sox2	forward reverse	5’-GCCGAGTGGAAACTTTTGTCG-3’ 5’-GCAGCGTGTACTTATCCTTCTT-3’	154	16
S18	forward reverse	5’-CTTAGAGGGACAAGTGGCG-3’ 5’-GGACATCTAAGGGCATCACA-3’	71	17

Immunocytochemistry of paraffin sections

Prior to staining, cells were fixed with 4% paraformaldehyde for 8 h at 4 °C, then the paraformaldehyde was replaced with DPBS. The tissue fragment on the membrane was collected and embedded in paraffin. Sections were cut to a thickness of 4 μm and then deparaffinized and rehydrated. Next, the sections were immersed in a Tris/EDTA solution with pH 9.0 for antigen retrieval, boiled for 20 min by microwave heating. Continue with the 1.5% bovine serum albumin (BSA: Sigma Aldrich, MO, USA) blocking for 1 h. Subsequently, the primary antibodies were added and sections were incubated at 4 °C overnight. Next, after washing three times with DPBS, the secondary antibodies were added and incubated at room temperature for 1 h. Nuclei were stained by incubating with NucBlue Fixed Cell ReadyProbes Reagent (Cat. #: R37606, Thermo Fisher Scientific, MA, USA) for 1 h at the same time. The following antibodies were used. Primary antibodies: cardiac troponin T mouse monoclonal (cTnT; Cat. #: MA5-12960, Thermo Fisher Scientific, Rockford, IL, USA), Myosin Light Chain 2 (MYL2; Cat. #: ab79935, abcam, UK), and Vimentin (Cat. #: ab92547, abcam, UK), all antibodies were diluted to 1/100 with 1.5% BSA. Secondary antibodies: goat anti-mouse antibodies Alexa Fluor 488 (Cat. #: A32723, Invitrogen, USA) and donkey anti-rabbit antibody Alexa Fluor 647 (Cat. #: ab150075, Abcam, UK), both were diluted to 1/1000 with 1.5% BSA. Fluorescence images were acquired using a confocal laser microscope (LSM780, Carl Zeiss, Oberkochen, Germany). The mean fluorescence intensity was analyzed by ImageJ software (US National Institute of Health, Bethesda, Maryland, USA). For quantification of fluorescence intensity, images were first converted to 8-bit format. The “Adjust Threshold” function was applied to exclude background and highlight specific signals. A consistent threshold value was used for all samples. As the iPS cells formed a continuous cardiomyocyte tissue structure, the entire fluorescent area of the tissue was selected as the region of interest (ROI), and average intensity was calculated. No individual cell segmentation was performed. Sarcomere directionality was quantified using the Directionality plugin in ImageJ (NIH, USA), which calculates the distribution of fiber angles based on image intensity gradients.

Transmission electron microscope observation

The cells were pre-fixed overnight at 4 °C in a 0.1 M cacodylate buffer containing 2% glutaraldehyde and 2% paraformaldehyde. After washing with a 0.1 M cacodylate buffer, the cells were post-fixed with 2% OsO₄ for 1.5 h at 4 °C, then washed with 0.1 M cacodylate buffer and dehydrated with ethanol. Then, they were embedded in Spurr resin (Polyscience) and thermally polymerized, and an ultra-thin section of 80 nm was prepared using ultramicrosome (LEICA EM UC7) (Leica Microsystems, Wetzlar, Germany). They were double stained with uranium and lead, and observed using a transmission electron microscope (H-7650, Hitachi High-Technologies, Tokyo, Japan). ImageJ software was used for sarcomere length measurement. For sarcomere length analysis, transmission electron microscopy (TEM) images were acquired from multiple fields per sample. A total of four independent biological replicates were analyzed for the control group and three for the stretch group. For each replicate, multiple individual sarcomeres were measured from different fields of view. The number of sarcomeres measured per sample varied depending on image quality and sarcomere visibility. The mean sarcomere length per sample was used for statistical analysis.

Video analysis of iPS-CMs

Videos of the contraction of the iPS-CMs were recorded for 10 s before and after mechanical stretching stimulation using a phase contrast microscope (BZ-X710, KEYENCE, Osaka, Japan) at 37 °C, 5% CO₂. Video analysis was done as described previously¹⁸. All videos were captured at a fixed position using a 4×objective lens at 20 frames per second. The displacement vector fields were calculated between frame 1 and all subsequent frames (frame 1 vs. 2, frame 1 vs. 3, frame 1 vs. 4, etc.). The displacement vector fields were calculated by using the particle image measurement plug-in¹⁹ of ImageJ (National Institutes of Health²⁰, and the displacement vector D (x, y) was obtained for each 16 × 16 pixel. The maximum displacement vector M (x, y) was defined as follows for each (x, y) pair:

Here, k represents the frame number at which | D_k (x, y) | = max [| D₂ (x, y) |, | D₃ (x, y) |, …, | D_n (x, y) |]. The number n indicates the last frame number (n = 200 in this study). The contractility C was calculated in arbitrary units as follows:

The rates of change in contractility before and after mechanical stretching stimulation were determined as follows:

Calcium imaging

iPS-CMs was stained with 5 µM calcium indicator Cal-520 (AAT Bioquest, USA) after stretching and incubated at 37 °C for 1 h. Then calcium transients were recorded using a fluorescence microscope (ECLIPSE TE2000-U, Nikon, Japan) at room temperature (24 °C) for 60s. Calcium imaging videos were analyzed using ImageJ software, peak detection and quantification were performed automatically using Spiky plugin²¹ and the data were averaged from multiple detected peaks.

Statistical analysis

All statistical analyses were performed using GraphPad Prism 9.5.0 (GraphPad Software, CA, USA). Data obtained from experiments were expressed as mean ± standard error of the mean (SEM). Two-way ANOVA or unpaired t-test were used for statistical analysis. P-values less than 0.05 were considered significant.

Results

Gene expression analysis in iPS-CMs

To compare expression levels of differentiation marker genes in response to mechanical stimulation, we performed quantitative reverse transcription polymerase chain reaction (qRT-PCR). As shown in Fig. 2a, cardiomyocyte differentiation was successfully achieved, as evidenced by the significant downregulation of pluripotent markers (Oct4 and Sox2). In Fig. 2b, the expression levels of cardiac-specific markers (cTnT and Nkx2.5) were higher in the stretch group compared to the control group. mRNA expression level of cTnT increased 2.5-fold in the stretch group compared to the control group (4.818 ± 0.737 vs. 1.939 ± 0.350, P < 0.01). Similarly, mRNA expression of Nkx2.5 increased 2-fold in the stretch group compared to the control group (2.601 ± 0.341 vs. 1.232 ± 0.147, P < 0.01).

Fig. 2.

mRNA expression of cardiac and pluripotent markers. qRT-PCR was employed to analyze the mRNA expression levels of a pluripotent markers (Oct4 and Sox2) between day 0 (before) and day 18 (after) and b cardiac markers (cTnT and Nkx2.5) on day 18 in iPS-CMs between the control and stretch groups. The data are presented as means ± S.E.M., n = 3 (Oct4, Sox2) and n = 5 (cTnT, Nkx2.5) for each condition. Statistical analysis was conducted using Two-way ANOVA (Oct4, Sox2) and an unpaired t-test (cTnT, Nkx2.5). **: P < 0.01, ***: P < 0.001.

Protein expression in iPS-CMs

The expression of the cardiac muscle marker cTnT and MYL2 were evaluated by fluorescent immunocytochemical staining. Mechanical stretching promotes the maturation of myocardial tissue. As shown in Fig. 3a, expression of these cardiac marker proteins were evident in both stretch and control groups. Striated pattern of cTnT was observed in the stretch group, implying developed sarcomere structure. In Fig. 3b, the fluorescence intensity of cardiac markers cTnT (16.98 ± 2.752 vs. 6.491 ± 0.649, P < 0.01) and MYL2 (14.02 ± 0.970 vs. 3.713 ± 0.576, P < 0.0001) respectively increased 2.6-fold and 3.8-fold in the stretch group compared to the control group, indicating enhanced sarcomere development.

Fig. 3.

Protein expression of cardiac markers and direction of sarcomere. a Immunofluorescence imaging of iPS-CMs in the control and stretch groups. Blue: Nuclei, green: cTnT, red: MYL2, arrows indicate sarcomere structure. b Mean fluorescence intensity of cTnT and MYL2 was compared between the control and stretch groups. The data was presented as means ± S.E.M., n = 6 represents independent biological replicates. c Frequency of directionality analysis of sarcomere in both groups, n = 5 represents independent biological replicates. Statistical analysis was performed using an unpaired t-test. **: P < 0.01, ****: P < 0.0001.

Sarcomere alignment in iPS-CMs

To investigate the impact of mechanical stretching on sarcomere arrangement, the direction of sarcomere was analyzed. Results demonstrated that mechanical stretching promotes patterned arrangement of cardiomyocyte sarcomere. In the control group, the arrangement of the fibers is random with no specific direction. In contrast, in Fig. 3a, there is a clear trend in the direction of the fibers in the stretch group. And as shown in Fig. 3c, the frequency of fiber orientations in the range of −20° to 20° was 1.5-fold higher in the stretch group compared to the control group (0.418 ± 0.031 vs. 0.284 ± 0.008, P < 0.01), The alignment of sarcomere fibers observed in the stretch group mirrors the orderly organization in functional myocardial tissue. This improved alignment is critical for synchronized contraction, a hallmark of mature cardiomyocytes.

Microstructural observations of iPS-CMs

To examine the microstructural differences between iPS-CMs in the control and stretch groups in greater detail, transmission electron microscopy was performed, as shown in Fig. 4a. Sarcomere structures were observed in both groups, with evidence suggesting that mechanical stretching enhances sarcomere maturation. In Fig. 4b, the sarcomere length in the stretch group (1.547 ± 0.029 μm) was significantly longer than that in the control group (1.219 ± 0.038 μm), closely approaching the range typical of mature myocardium (1.6–2.2 μm). Additionally, cells in the stretch group exhibited more well-defined Z-lines, distinct I-bands and prominent gap junctions. Moreover, iPS-CMs in the stretch group demonstrated a higher degree of alignment compared to those in the control group.

Fig. 4.

Transmission electron microscopy images of iPS-CMs. a Transmission electron microscopy images depict sarcomere structures of iPS-CMs in both control and stretch groups. Z-line and I-band (Z, I, black arrow), gap junction (GJ, white arrow) was shown. b A comparison of sarcomere lengths between the control and stretch groups. The data are presented as means ± S.E.M., a total of 72 sarcomeres were measured from four independent biological replicates in the control group, and 128 sarcomeres were measured from three independent replicates in the stretch group. Sarcomeres were measured from multiple fields of view per sample, and the number of sarcomeres per replicate varied depending on image quality. Statistical comparisons were performed using the mean sarcomere length per replicate. Statistical analysis was conducted using an unpaired t-test. **: P < 0.01.

Evaluation of contractility of iPS-CMs

Contractility is an important indicator of cardiomyocyte maturation. To compare contractile function of the iPS-CMs, we recorded the cultured cells under microscope. Spontaneous contraction of iPS-CMs was analyzed, and changes in contractility before and after stretching were visualized using vector field. The results indicated that the application of mechanical stretching during the induction of cardiomyocytes differentiation can improve the contractility of myocardial tissue. As shown in Fig. 5a, an increase in contractility was observed in the stretch group. Further, the change in the contractility before and after stretching was compared in both groups. Significant increase in contractility was observed in the stretch group compared to control group (−0.034 ± 0.061 and 0.411 ± 0.076, respectively) which was shown in Fig. 5b. The observed increase in contractility following mechanical stretching supports the hypothesis that mimicking physiological strain promotes functional maturation.

Fig. 5.

Contractility analysis of iPS-CMs. a Myocardial displacement vector field plot illustrating the direction and magnitude of displacement resulting from contraction, represented as vectors. b Comparison of the change ratio of contractility before and after stretching between the control and stretch groups. The data are presented as means ± S.E.M., n = 6 represents independent biological replicates. Contractility was measured from one representative field of view per sample. Representative spontaneous contraction videos from each group are provided as Supplementary Videos (https://figshare.com/s/387998937fa617645d62). Statistical analysis was conducted using an unpaired t-test. **: P < 0.01.

Ca²⁺ handling in iPS-CMs

Figure 6a illustrates a representative calcium transient waveform in iPS-CMs and Fig. 6b shows a schematic diagram of calcium transient analysis. The results indicate that mechanical stretching enhances the functional maturation of iPS-CMs in terms of calcium handling, a key hallmark of cardiomyocyte maturation. As shown in Fig. 6c, the amplitude of calcium transients did not differ significantly between the control and stretch groups (7.744 ± 0.841 vs. 9.274 ± 0.955, respectively). However, the cycling speed of calcium (peak-to-peak interval) was 1.8 times faster in the stretch group compared to the control group (2.374 ± 0.239 vs. 4.465 ± 0.299, P < 0.0001). Additionally, both the calcium transient peak time (time to peak) and the time to 50% transient decay were reduced by 30% following stretching (0.356 ± 0.019 vs. 0.519 ± 0.034, P < 0.001 and 0.253 ± 0.009 vs. 0.390 ± 0.028, P < 0.001, respectively). These findings highlight the role of mechanical stretching in accelerating calcium handling dynamics, contributing to the functional maturation of iPS-CMs.

Fig. 6.

Calcium transient analysis of iPS-CMs. a Representative intracellular calcium transient from iPS-CMs in control and stretch groups. b Schematic diagram of calcium transient analysis. c Values of amplitude, peak to peak, time to peak and time to 50% transient decay (time at 50% of amplitude from peak to next baseline at right). The data are presented as means ± S.E.M., with n = 12 represents independent biological replicates. Statistical analysis was conducted using an unpaired t-test. ns: no significance, ***: P < 0.001, ****: P < 0.0001.

Stretching increases the expression of vimentin in iPS-CMs co-cultured with HGF

To understand the response of cells to stretching, the expression of fibroblast marker vimentin was evaluated by fluorescent immunocytochemical staining. As shown in Fig. 7a, expression of vimentin was evident in both stretch and control groups. In Fig. 7b, protein expression was analyzed by quantifying mean fluorescence intensity, the protein expression of vimentin (6.476 ± 0.546 vs. 4.453 ± 0.405, P < 0.05) was 1.5-fold higher in the stretch group compared to control group, which demonstrated that mechanical stretching promotes vimentin expression in cells.

Fig. 7.

Protein expression of fibroblast marker. a Immunofluorescence imaging of cells in the control and stretch groups. Blue: Nuclei, red: vimentin. b Mean fluorescence intensity of vimentin was compared between the control and stretch groups. The data was presented as means ± S.E.M., with n = 6 represents independent biological replicates. Statistical analysis was performed using an unpaired t-test. *: P < 0.05.

In summary, mechanical stretching significantly enhanced the structural and functional maturation of iPS-CMs co-cultured with HGF. The observed improvements in sarcomere alignment, contractility, and calcium handling underscore the potential of combining mechanical stimulation with co-culture techniques for myocardial tissue engineering.

Discussion

These results show that the application of mechanical stretching stimulation in the process of inducing differentiation of iPS cells into myocardium can improve the maturation of iPS-CMs.

In this experiment, we applied mechanical stretching to iPS-CMs co-cultured with HGF for the first time and determined the maturation of differentiated myocardial tissue. Unlike previous studies that relied on static co-culture conditions, our findings demonstrate that dynamic mechanical stretching enhances both structural and functional maturation of iPS-CMs.

Gene and protein expression changes

The results of qRT-PCR showed a decrease in pluripotent markers Oct4 and Sox2 after differentiation induction in both the stretch and control groups, indicating that the iPS cells were successfully differentiated. In addition, compared to the control group, the myocardial marker cTnT and the mesodermal marker Nkx2.5 were elevated in the stretch group. Nkx2.5 is expressed early during differentiation into cardiomyocytes, while cTnT is a pan-cardiomyocyte marker that is widely used to confirm cardiomyocyte identity. Although its expression may increase during differentiation and early maturation, it is not considered a specific marker of mature myocardial tissue^22,23. The significant increase in Nkx2.5, a key cardiac transcription factor, may indicate enhanced activation of cardiomyocyte-related gene expression programs. In contrast, cTnT, as a structural protein, more likely reflects the outcome of differentiation rather than contributing directly to gene regulation.

The sarcomere is the basic structural and functional unit of the myocardium. cTnT is one of the three subunits that form troponin, which regulates the binding of actin to myosin, thereby controlling muscle contraction²⁴. MYL2 is a contractile protein belonging to the myosin family, and it interacts with the neck/tail region of the muscle thick filament protein myosin to regulate myosin motility and function²⁵. MYL2 plays a crucial role in early embryonic heart development and function²⁶ and is one of the earliest markers of ventricular specialization²⁷. Immunocytochemical staining of iPS-CMs showed that cTnT-positive cells (green) and MYL2-positive cells (red) were observed in both groups, with higher expression in the stretch group compared to the control group. Furthermore, experiments confirmed the presence of more prominent sarcomere structures in the stretch group, indicating the maturation of myocardial tissue.

However, it remains unclear whether the observed increase in cardiac marker expression reflects upregulated expression per cell or a higher proportion of differentiated cardiomyocytes in the stretch group. In this study, we did not employ cell population quantification methods, such as flow cytometry, to distinguish between these possibilities. Nonetheless, the combined evidence from immunostaining—namely, elevated cTnT and MYL2 protein levels and enhanced sarcomere structure—suggests that mechanical stretching may contribute to both differentiation and structural maturation. Future studies incorporating flow cytometry or single-cell RNA sequencing will be needed to clarify whether mechanical stimulation increases the proportion of cardiomyocytes or enhances gene expression within already differentiated cells. Additionally, commonly used gene expression ratios such as MYL2/MYL7 and TNNI3/TNNI1 are valuable indicators of cardiomyocyte maturation^28,29. Although these were not evaluated in the present study, they should be included in future analyses to more comprehensively assess the maturation status of iPS cell-derived cardiomyocytes under mechanical stimulation.

Sarcomere maturation and structural organization

At the same time, the stretch group exhibited a more organized arrangement of fibers in the myocardium (Fig. 3c). It is well established that the fiber arrangement in healthy, mature cardiomyocytes is highly ordered, and studies have confirmed that changes in the arrangement of cardiomyocyte fibers affect the contractility of myocardial tissue^30,31. According to these reports, cardiomyocytes with a normal arrangement beat synchronously, whereas cardiomyocytes with a disordered arrangement lose synchronized beating, with different parts of the cells contracting in different directions³². Therefore, the arrangement of fibers in cardiomyocytes is crucial for generating myocardial tissue that exhibits highly synchronized contractions.

To observe the intracellular structure in more detail, a transmission electron microscope was used. Sarcomere maturation is characterized by changes in microstructural organization³³. In our experiments, when comparing the sarcomere length between the two groups, the control group exhibited a length of 1.219 ± 0.038 μm, while the stretch group showed a greater value of 1.547 ± 0.029 μm. The sarcomere length of mature myocardium ranges from 1.6 to 2.2 μm³⁴, and the sarcomere length in the stretch group was closer to this range. A more mature sarcomere structure was observed in the stretch group, including regular Z-lines and well-defined I-bands. Gap junctions between cells also indicated the maturation of iPS-CMs³⁵. These findings suggest that mechanical stretching stimulation can promote the maturation of sarcomere structures.

Contractility and calcium handling

In the experiment, a video of contraction was analyzed and recorded, and changes in contraction before and after stretching were visualized and quantified. As a result, an increase in contraction was observed in the stretch group. The experiment also compared the change rate of the average maximum contractility before and after stretching between the two groups. It was observed that the myocardial tissue in the control group showed decreased contractility in some cases. On the other hand, all samples in the stretch group exhibited no decrease in contractility, suggesting consistent improvement across the group. Contractility is one of the most important indicators of cardiomyocyte maturation³⁶, indicating that mechanical stretching stimulation during the induction of myocardial tissue differentiation can enhance the contractility of myocardial tissue.

Changes in cell contractility were accompanied by alterations in Ca²⁺-handling behaviors. We found that the cycling speed of calcium was significantly faster in the stretched iPS-CMs, with a notably shorter time to peak and a faster 50% transient decay compared to the control iPS-CMs. However, there was no significant change in the transient amplitude. These results provide evidence that mechanical stretching enhances the calcium handling capabilities of iPS-CMs. This is important because efficient calcium cycling is critical for synchronized contraction, and defects in calcium handling are commonly observed in immature or diseased cardiomyocytes.

The faster calcium cycling observed in the stretched group may reflect more efficient sarcoplasmic reticulum (SR) function and better coupling between excitation and contraction. It is well known that calcium ions regulate the contraction and relaxation of cardiomyocytes through their interaction with the contractile proteins actin and myosin³⁷. By promoting faster calcium cycling, mechanical stretching may help establish more synchronized and forceful contractions, contributing to the maturation of the cardiomyocytes. Interestingly, no significant change in transient amplitude was observed, suggesting that the mechanical stretching might not have impacted the calcium influx into the cells. This could imply that the stretching primarily influenced the speed and dynamics of calcium handling, rather than the overall amount of calcium available for contraction.

Role of fibroblasts in cardiac maturation

Myocardial tissue is composed of cardiomyocytes and non-cardiomyocytes, including fibroblasts, which play essential roles in cardiac structure and function. Vimentin protein is a widely used marker for cardiac fibroblasts³⁸. Fibroblasts are involved in synthesizing the extracellular matrix (ECM) of the heart, providing mechanical scaffolding for cardiomyocytes, and influencing their function³⁹. Previous research has suggested that mechanical stretching stimulation induces the proliferation of cardiac fibroblasts, including human cardiac fibroblasts (HGF)^40,41. Furthermore, vimentin protein intermediate filaments function as shock absorbers, providing cellular mechanical protection, and act as scaffolds that help maintain cell shape and cytoskeletal organization⁴². Increased vimentin protein expression is associated with enhanced cell motility, adhesion to the ECM, and collagen deposition^43,44. In fibroblasts, mechanosensitive actin stress fibers, together with vimentin protein filaments, integrate into ECM adhesions to regulate the mechanical integrity of cells and tissues⁴⁵.

In our study, we observed a 1.5-fold increase in vimentin protein expression in the stretch group. This finding alone cannot distinguish whether the change represents a direct mechanotransducive effect on fibroblasts, increased proliferation, or extracellular matrix (ECM) remodeling. Therefore, while we propose that fibroblast responses to mechanical stimuli could influence the microenvironment and support iPS-CMs maturation, these effects remain hypothetical based on the current data. Vimentin is a general marker of mesenchymal cells and may reflect cellular activation rather than proliferation. Similar observations have been made in fibroblasts exposed to biomechanical stress^46,47 where increased vimentin expression correlated with ECM remodeling, but did not confirm causality. Future studies using proliferation markers (e.g., Ki67, EdU) and quantification of ECM components (such as collagen I/III or fibronectin) will be necessary to verify whether mechanical stretch indeed induces HGF proliferation or alters ECM composition in a way that promotes cardiomyocyte maturation.

Optimization of mechanical stretching

Many studies have explored the effects of different stretching rates in mechanical stimulation^12,48. According to research by Lux et al., mechanical stretching at 2%, 5%, 10%, and 20% rates was applied to cardiomyocytes. They found that stretching at 10% and 20% significantly reduced the contractile function of cardiac structures, while a 2% stretching rate did not induce any significant change in contractility compared to the control group⁴⁹.Moreover, a 20% stretching rate caused cellular damage, leading to apoptosis of the cardiomyocytes⁵⁰. Although most mechanical stretching protocols use 1 Hz to simulate the human heart rate^51–54, experiments have shown that 0.5 Hz is sufficient to induce cardiomyocyte differentiation^55,56. Therefore, in this experiment, under co-culture conditions with HGF, mechanical stretching with a 5% stretching rate and 0.5 Hz frequency was applied during the differentiation of iPS cells into myocardial tissue. This approach more closely mimics the in situ conditions of cardiomyocytes compared to standard culture dish conditions and does not induce excessive cellular stress.

Ronaldson-Bouchard et al. proposed a strategy of embedding early-stage differentiated iPS-CMs and fibroblasts into fibrin hydrogels, followed by the application of progressively increasing electromechanical stimulation to accelerate cardiomyocyte maturation⁵⁷. This approach successfully achieved near-adult myocardial structure and functional characteristics under in vitro culture conditions. However, the method relies on highly specialized equipment (such as flexible pillar electromechanical stimulation devices) and optimized culture conditions, which could present technical challenges for clinical translation and large-scale application. In contrast, our study introduces a simpler approach. We co-cultured iPS cells with HGF and induced cardiomyocyte differentiation in this co-culture system. By employing an external stretching device, we were able to explore the effects of mechanical stretching on the functional and structural maturation of iPS-CMs. Mechanical stretching was applied using a PDMS chamber, a device that is easy to operate and does not require complex electrical stimulation equipment.

Study limitations

This study has several limitations that should be acknowledged. First, the in vitro co-culture system, although controlled and reproducible, does not fully replicate the complex multicellular and biochemical environment of the native myocardium. Second, we did not perform long-term follow-up to assess whether the observed structural and functional improvements are sustained over extended periods. Finally, no in vivo validation was conducted to evaluate the therapeutic efficacy or integration potential of the matured iPS-CMs. Future studies should address these limitations to better translate these findings into clinical applications.

From a translational perspective, the co-culture system used in this study—based on human iPS cells and gingival fibroblasts—offers certain practical advantages such as accessibility of cell sources and relatively simple mechanical stimulation protocols. However, the current setup is not yet suitable for large-scale production or therapeutic use. To move toward clinical application, future work should aim to scale up this system using bioreactors, develop xeno-free and GMP-compliant culture protocols, and evaluate batch-to-batch reproducibility. In addition, adaptation for high-throughput screening or preclinical testing will require further engineering to ensure uniform mechanical stimulation and integration with automated platforms.

Conclusion

Our study highlights the potential of mechanical stretching as a practical and effective strategy for promoting the maturation of iPS-CMs. By focusing on the fibroblast response to mechanical forces, we present a novel approach to improving the efficiency and quality of myocardial tissue engineering. As such, future research into the mechanical regulation of fibroblasts and their role in heart tissue regeneration is essential for advancing the field and overcoming current challenges in clinical translation.

Author contributions

Conceptualization, Y.M., K.T., H.K. and K.N.; validation, M.W. and H.I.; data curation, M.W.; writing—original draft preparation, M.W., H.I., C.W., Y.L. (Yin Liang), Y.L. (Yun Liu); writing—review and editing, M.W., K.T.; supervision, K.T.; project administration, K.T.; funding acquisition, K.T. All authors have read and agreed to the published version of the manuscript.

Funding

This research received support from the Japan Society for the Promotion of Science (JSPS) through Grant-in-Aid for Scientific Research (B) (No. 20H04518) and Grant-in-Aid for Scientific Research (A) (No. 21H04960).

Data availability

Availability of data and material (data transparency): All datasets generated and/or analysed during the current study are available in the Figshare repository. 1-Quantitative RT-PCR: https://figshare.com/s/2fc5d0d57e7b0467ad37, 2-Immunocytochemistry of paraffin sections: https://figshare.com/s/bd29bbac4c8da3e375dc, 3-Transmission electron microscope observation: https://figshare.com/s/dfc8138d5cea4a4a0788, 4-Evaluation of contractility of iPS-CMs: https://figshare.com/s/387998937fa617645d62, 5-Calcium imaging: https://figshare.com/s/fb376bd736f0b3e2f5db. Code availability (software application or custom code): The ImageJ macro code used for contractility analysis in this study is available at: https://dx.doi.org/10.3791/61104.

Declarations

Competing interests

The authors declare no competing interests.

Ethics approval and consent to participate

All HGF handling and experimental procedures in this study were approved by the Ethics Committee of the Graduate School of Biomedical Sciences, Okayama University (approval number 1612-007-002). Written informed consent was obtained from all participants and/or their legal guardian(s). All methods were performed in accordance with the relevant guidelines and regulations.