The hiPSCs (American Type Culture Collection, ATCC) cell line present in this study were obtained from Stem Cell Bank of Chinese Academy of Sciences. The hESCs (H9 cell line, WA09, WiCell Research Institute) (Thomson et al., 1998) cell line present in this study were obtained from Shanghai Zhong Qiao Xin Zhou Biotechnology Co. Ltd. hiPSCs and hESCs were routinely cultured at 37 °C and 5% CO₂ in mTeSR™ one complete maintenance medium (Stemcell Technologies, Inc.) on 6-well plates coated with Matrigel (BD Biosciences, Bedford, MA). Cells were passaged every 2–3 days using ACCUTASE™ cell detachment solution (Stemcell Technologies, Inc.). In order to direct differentiation into CMs as our previous work described (Ke et al., 2020), hiPSCs and hESCs were split at a 1:6 ratio and plated onto Matrigel-coated 12-well plates cultured for 3 days. When the hiPSCs and hESCs reached 85% confluence, 6 μM CHIR99021 (MedChemExpress) was added to RPMI/B27 without insulin medium (Gibco, Invitrogen, USA) to initiate the differentiation. On day 2, the medium was changed to RPMI/B27 minus insulin medium supplement with 3 μg/mL IWP2 (MedChemExpress). On day 5, the medium was changed to RPMI/B27 without insulin. On day 7, the differentiating cells were cultured in RPMI/B27 plus insulin medium (Gibco, Invitrogen, USA). On day 15, the cells were cultured with purification medium (Tohyama et al., 2013), which are comprised of glucose-depleted DMEM (Gibco, Invitrogen, USA), 213 μg/mL of l-ascorbic acid 2-phosphate supplemented (Sigma-Aldrich; Merck KGaA), and 500 μg/mL of recombinant human albumin (Oryzogen; Wuhan Healthgen Biotechnology, Corp.) combining with 4 mM L-lactic acid (Sigma-Aldrich; Merck KGaA). The medium was refreshed every 2 days during the purification process. On day 19, cells were maintained in RPMI/B27 plus insulin media. To investigate the effect of DOX (MedChemExpress) in hiPSC-CMs and hESC-CMs, cells at differentiation day 21 were dissociated by using 0.25% Trypsin-EDTA (Gibco, Invitrogen, USA) for 3 min at 37 °C and then centrifuged at 1,000 rpm for 5 min. Cells were replanted in DMEM (Gibco, Invitrogen, USA) containing 20% FBS (Gibco, Invitrogen, USA) for 24 h and changed in a chemically defined medium (CDM3) for maintenance. CDM3 medium consists of DMEM, 500 μg/mL of Oryza sativa–derived recombinant human albumin (Sigma-Aldrich; Merck KGaA), and 213 μg/mL of L-ascorbic acid 2-phosphate (Sigma-Aldrich; Merck KGaA).

For immunocytochemical staining, we fixed cells with 4% PFA (Sigma), permeabilized them with 0.4% Triton X-100 (Sigma), and then incubated them with the primary antibody anti-cTnT (Abcam, USA, 1:400) and anti-Cx43 (Abcam, USA, 1:400) overnight at 4 °C. The nucleus was stained with DAPI (Invitrogen, USA). All samples were imaged with a Carl Zeiss microscope and processed with ZEN software.

For characterization of hiPSC-CMs and hESC-CMs, the differentiated CMs were collected after dissociation with 0.25% Trypsin-EDTA for 3 min. The cells were fixed and permeabilized with the Foxp3 Staining Buffer kit (Invitrogen, USA) for 30 min, followed by incubation with the primary antibody (anti-cTnT, Abcam, 1:100) for 1 h at room temperature. The PE-conjugated secondary antibody (Biolegend, 1:400) was added to the cells for 1 h at 4 °C. Apoptosis was assessed by flow cytometry, using an Annexin V-FITC Apoptosis Detection Kit (BD Pharmingen). The assay was conducted based on the instructions which were provided by the manufacturer. Briefly, after treatment with DOX, 5 × 10⁵ cells were rinsed twice in PBS and incubated in 100 μ L one x binding buffer. Five μ L of Annexin V-FITC was added to the cells and incubated at RT for 15 min in the dark. Propidium iodide was not added because of DOX autofluorescence which may interfere with its detection. All samples were analyzed with a flow cytometer (BD Accuri™ C6, BD Biosciences) and quantified with FlowJo software.

The viability of cells was evaluated by CCK8 assay (Beyotime Institute of Biotechnology, Jiangsu, China). Briefly, 1 × 10⁴ cells/well CM cells were seeded into 96-well plates (Gibco, Invitrogen, USA) and cultured for 3 days. The cells were then treated with the different doses (0, 0.1, 0.25, 0.5, 1, 1.5, 2.5, 5.0, 10, 25, 50, 100 μM) of DOX for 24 h. 24 h later, the medium was exchanged with a fresh medium containing CCK8 reagent and incubated for an additional 4 h at 37 °C. By using an enzyme-linked immunosorbent assay reader (Thermo Fisher Scientific, Inc.), the data were read at an absorbance of 450 nm wavelength.

Cells seeded on a 96-well culture plate were treated with DOX for 24 h. The LDH release assay was performed to detect cell cytotoxicity, following the manufacturer’s instructions (Beyotime Institute of Biotechnology, Jiangsu, China). Briefly, the culture medium was collected and incubated with the working mixture (lactate, INT solution and diaphorase) at 37 °C for 30 min in dark. The absorbance of the sample was measured by an enzyme-linked immunosorbent assay reader at 490 nm. First, cardiomyocytes were incubated with 5 mM 3-MA (MedChemExpress) (Hou et al., 2012) or 2.5 μM Rapamycin (MedChemExpress) (Zhou et al., 2007) for 12 h, then stimulated with 0.25 μM, 0.5 μM and 1 μM DOX, after that both were maintained for 24 h. The LDH release assay was conducted to detect cell cytotoxicity, following the manufacturer’s instructions as previously described.

Intracellular ROS generation was analyzed using a ROS Assay Kit (Beyotime Institute of Biotechnology, Jiangsu, China). In accordance with instructions, after treatment with different concentrations (0, 0.25, 0.5, 1 μM) of DOX for 24 h, cells were incubated with DCFH-DA for 30 min at 37 °C and then washed three times with the medium. The fluorescence intensity was measured using the fluorospectrophotometer at 488 nm excitation and 525 nm emission wavelengths (PerkinElmer, Canada). The ROS fluorescence intensity was normalized relative to the mean Hoechst fluorescence intensity. Meanwhile, the cells were detected by flow cytometry. For further analysis of mitochondrial ROS in hiPSC-CMs and hESC-CMs, the live cells were stained with MitoSOX Red dye (ThermoFisher, Life Technologies, USA) according to the manufacturer’s instructions. The images were obtained with Fluorescence microscopy, and analyzed with Image J software.

hiPSC-CMs or hESC-CMs were seeded on gelatin-coated coverslips and treated with or without DOX in 12-well plates. After 24 h, cells were washed twice with PBS, fixed in 4% formaldehyde in PBS at RT for 30 min, and then permeated with 0.1% Triton-100X at RT for 15 min. Next, the coverslips were washed, and blocked with 10% goat serum for 1 h at RT and then incubated with the first antibody (γ-H2AX, 1:200) overnight at 4 °C. The cells were then washed thrice for 5 min and stained with Alexa Fluor™ 555-conjugated antibody (1:500; Beyotime Institute of Biotechnology). After staining for 10 min with DAPI, cells were analyzed by an Optiphot-2 microscope (Nikon Corporation) equipped with a CCD video camera system (Optronics Engineering, Ltd.) or a confocal laser scanning microscopy (Optronics Engineering, Ltd.).

To analyze autophagic flux, hiPSC-CMs or hESC-CMs were replanted on Laser confocal dishes (JingAn Biological Technology Co., Ltd, Shanghai, China). Cells were washed twice with PBS and infected with adenovirus expressing mCherry-GFP-LC3B fusion protein (Ad-mCherry-GFP-LC3B; Beyotime Institute of Biotechnology, Jiangsu, China) at an MOI of 20 for 24 h. After infection, the cells were treated with 0.5 μM of DOX for 6, 12 and 24 h. Then, cells were fixed with 4% paraformaldehyde, and then visualized with a confocal laser scanning microscopy (Olympus). We measured autophagic flux at different time points by observing the color change of mCherry/GFP.

hiPSC-CMs or hESC-CMs prepared for TEM were treated with or without DOX for 24 h. The cells were harvested, fixed with 5% glutaraldehyde (Sigma-Aldrich; Merck KGaA) at 4 °C overnight, and then postfixed with 1% osmium tetroxide (Sigma-Aldrich; Merck KGaA) at RT for 1 h. After fixation, the cells were dehydrated through a graded series of ethanol (Sinopharm Chemical Reagent Co., Ltd). Subsequently, the samples were treated with Spurr embedding agents mixed with various specifications of acetone (Sigma-Aldrich; Merck KGaA). Then, the ultrathin sections were mounted on nickel grids, followed by staining with lead citrate, uranyl acetate and 50% ethanol solution (Sigma-Aldrich; Merck KGaA). Thin sections of each sample were observed and analyzed under a transmission electron microscope (Hitachi H −7,650).

Proteins were extracted from hiPSC-CMs or hESC-CMs with RIPA lysis buffer then centrifuged at 120,00× g for 30 min. The concentration was quantified with Bradford assay (Thermo-Fisher Biochemical Co. Ltd, Beijing, China). For immunoblotting analysis, 30 μg of total proteins were loaded into the PAGE-SDS gels. The proteins were separated and transferred to a nitrocellulose membrane (Millipore, Billerica, MA, USA). Next, the membrane was blocked in 5% Nonfat dry milk in PBST and then incubated overnight at 4 °C followed by primary antibodies against GAPDH (1:5000), mTOR (1:500), p-mTOR (1:500) and LC3B (1:1,000) purchased from Cell Signaling Technology. Subsequently, the membrane was rinsed with PBST and incubated with horseradish peroxidase-conjugated goat anti-rabbit or mouse immunoglobulin G (IgG) antibody (Proteintech) for 1 h at RT. Afterward, the membrane was then washed with PBST three times and visualized with enhanced chemiluminescence (ECL) reagent (Amersham; GE Healthcare Life Sciences), and images were captured with the ECL Tanon 5500 system (Tanon Science and Technology Co., Ltd.).

In all experiments, data were detected by three or four independent biological replicates (n = 3–4) and presented as means ± standard deviation (SD). Grayscale analysis was quantified using ImageJ software. Prior to parametric testing, the Shapiro–Wilk test confirmed normal distribution and the Brown–Forsythe test confirmed homogeneity of variances for all datasets. Data Statistical significance was analyzed by using the unpaired Student’s t-test or one-way ANOVA followed by Dunnett’s post hoc multiple comparison test. Two-way ANOVA was employed to analyze the effects of two independent variables. Differences were considered statistically significant when p < 0.05.

hiPSCs and hESCs were differentiated to CMs as described previously (Ke et al., 2020) with slight modifications (Figure 1A). Immunocytochemical staining results showed that the cardiomyocyte-specific marker cTnT and the mature gap-junction marker Cx43 were expressed in hiPSC-CMs and hESC-CMs (Figure 1B). Flow cytometry analysis revealed that over 97% of the hiPSC-CMs and hESC-CMs were expressing the cTnT, indicating that these cells represent a highly pure cardiomyocyte population (Figure 1C). By day 10 post-differentiation, both hiPSC-CMs and hESC-CMs exhibited robust spontaneous contractions, providing qualitative evidence of electrophysiological activity (Supplementary Videos S1, S2).

In clinical studies, DOX can induce cardiomyocyte death within hours of intravenous administration in some patients at relatively low cumulative doses of 200–250 mg/m2 (Unverferth et al., 1983; Swain et al., 2003). The CCK8 results showed that DOX treatment resulted in a dose-dependent decrease in cell viability (Figure 2A). The cardiotoxicity was induced at a low concentration of DOX (0.1 μM for 24 h). We next investigated the cell damage caused by different concentrations of DOX (0–1 μM), which were chosen based on prior studies and are considered clinically relevant, as they fall well within the plasma concentration range observed in patients undergoing DOX therapy (Burridge et al., 2016; Frost et al., 2002). As shown in Figure 2B, LDH release significantly increased in a dose-dependent manner (0.25, 0.5 and 1 μM) following DOX treatment. Subsequently, we investigated the DOX-induced apoptosis in hiPSC-CMs and hESC-CMs. At a high concentration of 1 μM, DOX caused the most significant cell damage, prompting its use in the present study for apoptosis analysis. Flow cytometry analysis demonstrated that exposure of CMs to 1 μM DOX for 24 h resulted in a significant increase of Annexin V-FITC⁺ cells (an indicator of total apoptotic cells) compared to vehicle-treated control cells (Figure 2C). Moreover, morphologies of nuclear shrinkage and apoptotic bodies, indicative of typical apoptotic features, were observed by TEM in the DOX-treated group (Figure 2D). These data demonstrate that DOX treatment decreases cell viability and induces apoptosis in hiPSC-CMs and hESC-CMs. Collectively, these results from our study align with previous cellular and human biopsy reports (Burridge et al., 2016; Maillet et al., 2016; Zhao and Zhang, 2017; Cui et al., 2019), confirming the cardiotoxic effects of DOX and establishing a reliable in vitro model for assessing DIC in hiPSC-CMs and hESC-CMs.

To investigate the damage role of DOX in hiPSC-CMs and hESC-CMs, we first measured intracellular ROS production following DOX treatment using the probe DCFH-DA. As shown in Figure 3A, compared with the control, DOX induced a significant increase in intracellular ROS in hiPSC-CMs at 0.25, 0.5, and 1 μM, whereas in hESC-CMs significance was observed only at 0.5 and 1 μM. Similarly, the results were confirmed by flow cytometry analysis (Figure 3B). Additionally, a dose-dependent increase in MitoSOX red fluorescence was observed in cells exposed to DOX (Figure 3C). Next, the level of double-stranded DNA damage was assessed through staining for phosphorylated H2A histone family member X (γ-H2AX) on serine 139. As shown in Figure 3D, DOX treatment induced γ-H2AX expression in hiPSC-CMs in a dose-dependent manner, with significant DNA damage evident at a concentration of 0.5 μM. Therefore, we evaluated γ-H2AX expression in hiPSC-CMs and hESC-CMs treated with 0.5 μM DOX using laser confocal microscopy. The results demonstrated that DOX induced an increase in γ-H2AX expression in both hiPSC-CMs and hESC-CMs groups (Figure 3E). These findings indicate that DOX increases ROS levels and triggers DNA damage, leading to apoptosis, which represents one of the mechanisms of DIC.

Autophagy has dual functions in both physiology and pathology. To evaluate the changes in autophagy, we first observed ultrastructure changes and autophagosome formation in hiPSC-CMs and hESC-CMs by TEM. The results revealed an accumulation of autophagosomes and autolysosomes in DOX-treated groups (Figure 4A). Western blots were performed to detect the changes in the ratios of LC3 II/LC3 I in hiPSC-CMs and hESC-CMs treated with or without DOX. We found that DOX significantly upregulated the ratio of LC3 II/LC3 I in the hiPSC-CMs and hESC-CMs (Figure 4B). The most pronounced changes in autophagy-related proteins were noted at a DOX concentration of 0.5 μM. Therefore, the concentration of 0.5 μM was utilized in subsequent studies to investigate the effects of DOX on autophagy and its underlying mechanisms. To further study DOX-induced autophagic flux, we detected autophagy in hiPSC-CMs and hESC-CMs infected with adenovirus expressing a dual-fluorescence reporter mCherry-GFP-LC3B fusion protein (Ad-mCherry-GFP-LC3B) at a different time point (6 h, 12 h, and 24 h). Treatment with DOX resulted in an increase of autophagic flux within 6 h, as indicated by the number of both yellow and red puncta, with the maximum effect observed at 24 h (Figure 4C). Moreover, the DOX-induced increases in the LC3 II/LC3 I ratio were attenuated by 3-MA, an autophagy inhibitor, as shown in Figure 4D. In addition, to assess the role of autophagy in DOX-induced cardiomyocyte death, we evaluated the effects of 3-MA on LDH release. In Figure 4E, compared with the control, DOX induced an increase in LDH level, indicating cellular damage. However, pretreatment with the autophagy inhibitor 3-MA led to a significant yet partial reduction in DOX-induced LDH release at both 0.5 and 1 μM, resulting in a partial rescue effect (Figure 4E; Supplementary Table S1). Collectively, these results suggest that DOX upregulates autophagy in both hiPSC-CMs and hESC-CMs, which partially contribute to cardiotoxicity.

The kinase mTOR functions as a key signaling ‘station’ in the regulation of cellular metabolism, promoting protein synthesis while inhibiting the induction of autophagy (Liang, 2010). Although other pathways such as PI3K/AKT and AMPK also play roles in autophagy regulation, mTOR is widely recognized as one of the most pivotal and extensively studied mediators of autophagic processes (Shackebaei et al., 2024). Given the gatekeeper role of mTOR in autophagy regulation, we investigated whether DOX alters autophagy in hiPSC-CMs and hESC-CMs through mTOR signaling. We found that DOX treatment notably decreased mTOR activation in hiPSC-CMs and hESC-CMs, as evidenced by the reduced phosphorylation at Ser-2248 across various concentrations (Figure 5A). To further confirm the role of mTOR signaling in autophagy induced by DOX, we next treated hiPSC-CMs and hESC-CMs with rapamycin (RAPA), an mTOR inhibitor. In hiPSC-CMs, RAPA alone increased LDH release and exacerbated DOX-induced cell death, although without significant additive cytotoxicity (Figure 5B; Supplementary Table S1). In contrast, in hESC-CMs co-treatment with RAPA and DOX (0.25 and 0.5 μM) resulted in a significant additive increase in LDH release, suggesting the inhibition of mTOR enhances the cytotoxic effects of DOX (Figure 5B; Supplementary Table S1). Collectively, these experiments demonstrate that DOX stimulates autophagy in hiPSC-CMs and hESC-CMs through inhibiting mTOR signaling.