Doxorubicin‐Induced Cytotoxicity in Adipose‐Derived Stem Cells Is Associated With Altered SAPK/JNK Signaling

ABSTRACT

Adipose‐derived stem cells (ADSCs) are widely used in regenerative medicine, but their functionality declines under chemotherapeutic stress. Doxorubicin (Dox) is an effective anticancer agent known to induce long‐term toxicity in healthy tissues. Increasing evidence suggests that Dox promotes cellular dysfunction, including apoptosis, promotion of oxidative stress, and premature senescence. The SAPK/JNK signaling pathway is implicated in stress responses and may contribute to Dox‐induced aging in stem cells. However, its role in ADSC senescence and functional decline remains unclear. This study evaluated the effects of Dox on ADSC viability and aging‐associated processes, with a focus on SAPK/JNK signaling. ADSCs exposed to 0.1–100 μM Dox for 24 h showed reduced mitochondrial activity and ATP levels at clinically relevant doses (5 μM), along with disrupted cell cycle progression and cytoskeletal alterations. Dox induced both apoptosis and premature senescence and increased oxidative stress. These effects were accompanied by alterations in SAPK/JNK pathway expression. Overall, Dox promoted ADSC dysfunction, highlighting potential limitations in the therapeutic use of ADSCs during chemotherapy and emphasizing the need for protective strategies. Alterations in the SAPK/JNK signaling pathway were observed in response to Dox‐induced stress, suggesting that this pathway may contribute to the cellular stress response. These findings suggest that the development of approaches to preserve ADSC function and mitigate Dox toxicity is critical for improving the safety and efficacy of regenerative medicine applications.

This study investigated the effects of clinically relevant concentrations of doxorubicin (Dox; 5 μM), commonly used in anticancer therapy, on adipose‐derived stem cells (ADSCs). Dox induced ADSC dysfunction, characterized by reduced mitochondrial activity, apoptosis, oxidative stress, premature senescence, cytoskeletal alterations, and cell cycle disruption. These effects were accompanied by alterations in SAPK/JNK pathway expression, suggesting a role for this signaling pathway in the cellular stress response. Collectively, these findings indicate that Dox‐induced ADSC dysfunction may limit their therapeutic potential and highlight the need to develop protective strategies to preserve ADSC function.

1. Introduction

Adipose‐derived stem cells (ADSCs) are well characterized, safe, and readily accessible for mesenchymal stem cells (MSCs) that exhibit low immunogenicity and pose only a minimal risk of tumorigenesis. The differentiation potential and high proliferative capacity of ADSCs support their application in cell‐based therapies, including the replacement of both soft and hard tissues [1, 2]. ADSCs have shown efficacy in tissue reconstruction following cancer treatment for breast [3] or bone [4] cancer. Moreover, ADSCs display substantial paracrine activity, indicating their potential usefulness in noncellular therapeutic strategies, such as a treatment of tendon disorders using extracellular vesicles (EVs) [5] or treatments targeting ovarian cancer [6]. However, numerous limitations hinder the efficient use of ADSCs in regenerative medicine [7] because many factors adversely affect the viability of these cells. Furthermore, the donor's comorbidities, age, lifestyle, and use of pharmaceutical agents must be considered when using ADSCs as autologous grafts, as our previous work has shown [8]. In addition, culture conditions and the duration of in vitro expansion can reduce the number of viable ADSCs, primarily due to cellular senescence, which can trigger undesirable alterations in cell morphology and proliferation rates, as well as declines in ADSC differentiation potential [9].

Senescence in every type of MSCs can manifest in multiple forms, distinguishable by distinct replicative, oncogene‐induced, stress‐induced, or developmental processes [10]. For this reason, in stem cell research, where cellular alterations occur rapidly, precise characterization of each form of cellular impairment based on genetic and phenotypic profiles is essential. Numerous studies have reported that MSCs can mitigate the side effects induced by chemotherapeutics by acting on the cellular microenvironment [11]. However, if MSC functionality is compromised, the cells cannot exert these protective effects and become poor choices for use in tissue regeneration.

One factor that can modify the therapeutic potential of stem cells is the use of doxorubicin (Dox) as a chemotherapeutic agent. Dox is an anthracycline drug used in the treatment of various cancers, including lung and breast cancers, as well as lymphoma, soft tissue sarcoma, and osteosarcoma [12, 13]. Although Dox shows high efficacy against cancer cells and has a broad action spectrum, it often induces serious long‐term side effects on healthy noncancerous somatic cells, including cardiomyocytes, endothelial cells, and fibroblasts. In the heart, this leads to Dox‐induced cardiotoxicity (DCT) and consequent cellular aging of otherwise healthy heart tissue [14]. The mechanism by which Dox initiates toxicity in healthy somatic cells, such as cardiomyocytes, is likely associated with the generation of reactive oxygen species (ROS), which promote mitochondrial damage and the subsequent activation of apoptosis, autophagy, ferroptosis, and cellular senescence [15]. Likely, the same toxicity mechanism could also adversely affect the functioning of therapeutic ADSCs. However, research on the effects of anticancer therapies on the functional potential of MSCs, including Dox effects on ADSCs, remains insufficient.

Aging is known to hinder the proper functioning of ADSCs and their use in clinical practice [16]. Consequently, the senescence‐inducing effects of Dox could be particularly detrimental for new biomaterial engineering anticancer strategies that incorporate chemotherapeutic agents into the tissue‐replacement scaffolds that provide platforms for stem cell cultivation [4]. Perhaps an even greater concern is the possible failure of ADSC‐related stem cell therapy in patients who are undergoing or have undergone Dox chemotherapy.

Apart from its cytotoxicity to noncancerous cells, Dox is also known to induce chemoresistance in cancer cells, as observed in patients with, for example, triple‐negative breast cancer [12]. This chemoresistance appears to arise via the stress‐activated protein kinase/Jun amino‐terminal kinase (SAPK/JNK) pathway, which also promotes the metastasis of cancer cells [17]. Importantly, although the proteins of the JNK pathway promote the self‐renewal and maintenance of cancer stem cells [18], the SAPK/JNK pathway also plays a role in the proper functioning of noncancerous stem cells by promoting their proliferation and maintaining their pluripotent state [19]. Therefore, identifying the mechanisms underlying chemoresistance in cancer cells, and especially in cancer stem cells, may help determine which molecular pathways could be targeted to induce Dox resistance in ADSCs.

In the present study, we hypothesized that the SAPK/JNK pathway is altered in ADSCs following Dox exposure and that modulation of the pathway activity might enhance stem cell survival. Therefore, the aims of this study were to evaluate the cytotoxic effects of Dox on ADSCs and to determine whether SAPK/JNK expression is associated with this cytotoxicity. There is still insufficient evidence supporting this mechanism in ADSCs. Although their use in clinical trials has increased each year, this unresolved aspect may contribute to important limitations.

2. Materials and Methods

2.1. Experiments

The experiments were performed using normal human ADSCs (PT‐5006, Lonza, Basel, Switzerland) originated from single, healthy donor. The effects of Dox were comprehensively assessed by determining the half maximal inhibitory concentration (IC₅₀), defined as the concentration of a compound that reduces cellular activity by 50% and comparing the viability of control ADSCs versus ADSCs exposed for 24 h to Dox (0.1–100 μM). Cell viability was evaluated using mitochondrial dehydrogenase activity (MTT assay), intracellular ATP level (CellTiter‐Glo 2.0 assay), and total protein content measured by the sulforhodamine B (SRB) assay. Based on these results, three Dox concentrations (5, 25, and 50 μM) were selected for subsequent experiments to evaluate apoptosis, cellular senescence, induction of oxidative stress and cytoskeletal organization. Furthermore, activation of the stress‐activated protein kinase/c‐Jun N‐terminal kinase (SAPK/JNK) signaling pathway in Dox‐treated cells was evaluated at both the transcriptional and protein expression levels.

2.2. Cell Culture Conditions

ADSCs were cultured in Dulbecco's Modified Eagle Medium (DMEM, 11965‐092; Gibco, Massachusetts, USA), supplemented with 10% fetal bovine serum (FBS, ECS0180L; EuroClone, Pero, Italy), and a 1% penicillin–streptomycin mixture (17‐602E; Lonza, Basel, Switzerland) at 37°C in a 5% CO₂ incubator (Sanyo MCO‐19M, Osaka, Japan). The culture medium was changed every 3 days. The experiment was performed on cells in the logarithmic phase. The ADSCs used for the experiment were at the fourth passage. The cells were assessed using an Olympus IX73 microscope (Olympus, Shinjuku, Tokyo, Japan), which was also used for photographic documentation.

2.3. Doxorubicin Solution Preparation

Doxorubicin (doxorubicin hydrochloride, D1515‐10MG; Sigma‐Aldrich, St. Louis, MO, USA) was prepared in sterile DMSO (D2650; Sigma‐Aldrich, St. Louis, MO, USA) and serially diluted in complete culture medium to final concentrations ranging from 0.1 to 100 μM, using a twofold dilution series to generate a dose–response curve. Control wells were treated with vehicle alone to account for potential solvent effects.

2.4. Cell Phenotyping

Flow cytometry analysis was used to characterize the cells by identifying the following surface mesenchymal stem cells positive markers: CD73‐CFS, CD90‐APC, and CD105‐PerCP in comparison to negative markers: CD45, CD34, CD11b, CD79A, and HLA‐DR‐PE. The Human Mesenchymal Stem Cell Verification Flow Kit (FMC020; R&D Systems, Minneapolis, MN, USA) was used for all experiments [20] (Figure S2). Cells were differentiate also into mesoderm cells: adipocytes, chondrocytes, and osteocytes using a Human Mesenchymal Stem Cell Functional Identification Kit (Cat. #SC006; R&D Systems, Minneapolis, MN, USA) to assess their multipotency according to the manufacturer's protocol [21] (Figure S2).

2.5. Cell Viability Assays

ADSCs were plated at a density of 5 × 10³ per well in 96‐well plates, treated with Dox at the concentrations of: 0.1, 1, 2.5, 5, 10, 25, 50, and 100 μM and incubated for 24 h before conducting viability assays. Cell viability assays were performed as eight biological replicates (N = 8).

2.5.1. MTT Assays for Mitochondrial Activity Analysis

After the 24 h of incubation with Dox, the viability of cells related to mitochondrial activity was evaluated using the MTT assay (Thiazolyl Blue Tetrazolium Bromide, M2128; Sigma‐Aldrich, St. Louis, MO, USA), according to the manufacturer's protocol. MTT solution was added, and cells were incubated for 3 h at 37°C in 5% CO₂ (incubator Sanyo MCO‐19M, Osaka, Japan). The medium was aspirated, and DMSO was added to the cells for 1 h at room temperature (RT) to dissolve the formazan crystals. Untreated cells served as a control (0 μM), whereas cells treated with 0.1% Triton X100 served as a negative control (c neg). The formazan absorbance was measured at 570 nm using a microplate reader (VICTOR Nivo; PerkinElmer, Shelton, CT, USA).

2.5.2. CellTiter‐Glo 2.0 Assay for Quantifying ATP

Cells were plated at a density of 5 × 10³ cells per well in 96‐well black plates. The effect of Dox was analyzed using the CellTiter‐Glo 2.0 assay kit (G9241; Promega, Madison, WI, USA) according to the manufacturer's protocol. The bioluminescent CellTiter‐Glo 2.0 assay determines the number of viable, metabolically active cells in culture by quantifying ATP. It is based on a luciferase enzymatic reaction and uses ATP from viable cells to generate photons of light. After 24 h incubation with Dox, the plate was equilibrated to RT for 30 min. The CellTiter‐Glo 2.0 Reagent (100 μL) was then added to each well of the cell culture plate. Control wells containing medium without cells were prepared to determine background luminescence. The luminescence (which is directly proportional to the number of viable cells in culture) was recorded using a microplate reader (VICTOR Nivo; PerkinElmer, Shelton, CT, USA).

2.5.3. Sulforhodamine B Assay for Total Cellular Protein Content

The cytotoxic effect of Dox on ADSCs was assessed using the Sulforhodamine B (SRB) assay (TOX6; Sigma‐Aldrich, St. Louis, MO, USA), which quantifies cellular biomass based on protein staining. Cells were seeded at 5 × 10³ per well in 96‐well plates and treated with Dox at the selected concentrations. The SRB assay was performed according to the manufacturer's protocol. Cells were fixed with trichloroacetic acid (TCA) and plates were incubated at 4°C for 1 h and then washed with deionized water to remove TCA. After air‐drying, the cells were stained with 0.4% SRB solution in 1% acetic acid for 30 min at RT. Unbound dye was removed by repeated washing with 1% acetic acid. After air drying, the incorporated SRB dye was solubilized with 10 mM Tris base for 5 min at RT with shaking. The absorbance was measured spectrophotometrically at 565 nm using a microplate reader (VICTOR Nivo; PerkinElmer, Shelton, CT, USA). The background absorbance at 690 nm was subtracted from each measurement for nonspecific signals.

2.6. Morphological and Cytoskeletal Analysis of Cells

ADSCs were cultured under standard conditions until they reached the desired confluence and characterized (as shown in Section 2.4). Prior to the initiation of experiments, the expression of MSCs surface markers CD73, CD90, and CD105 was analyzed (Figure S1) and the multipotency potential of ADSCs was confirmed (Figure S2).

The cells were then treated with 5, 25, or 50 μM Dox for 24 h, followed by microscopy examination at 200× and 400× magnification to observe the cell morphology and actin filament arrangement.

Morphological Analysis of the Cells Was Performed Using Phase‐Contrast Bright‐Field Imaging at 200× Magnification. ADSCs Were Evaluated Based on Their Shape, Size, Adhesion Properties, Nuclear‐Associated Granularity, and the Presence of Cytoplasmic Vacuolation.

For this purpose, ADSCs were fixed in 4% formaldehyde for 15 min at RT. Following fixation, the cells were rinsed three times in phosphate‐buffered saline (PBS, 21‐040‐CV, Corning, NY, USA) for 5 min each to remove residual fixative. Immunostaining was performed according to the Cell Signaling Technology protocol. Actin filaments were then stained with Alexa Fluor 555 coupled phalloidin (8953; Cell Signaling, Danvers, MA, USA; diluted in PBS at a 1:100 ratio) applied to the cells. After a 15 min incubation at RT, the cells were rinsed with PBS to remove unbound dye. Cell nuclei were stained with DAPI dye (VECTASHIELD Vibrance Antifade Mounting Medium, H‐1700; Vector Laboratories, Newark, CA, USA). The specimens were examined under a fluorescence microscope at 400× magnification using the excitation wavelength 553 nm.

2.7. Assessment of Apoptosis Using the RealTime‐Glo Annexin V Apoptosis Assay

Apoptosis in ADSCs was assessed using the RealTime‐Glo Annexin V Apoptosis Assay (JA1000; Promega, Madison, WI, USA) following the manufacturer's instructions. The positive apoptosis control treatment was 0.05% Triton X100 (93443‐100ML; Sigma Aldrich, St. Louis, MO, USA). Cells were cultured in DMEM medium supplemented with 10% FBS and 1% penicillin–streptomycin under standard conditions at 37°C in a humidified atmosphere containing 5% CO₂. Prior to treatment, ADSCs were seeded at a density of 3 × 10⁴ cells/well in white tissue culture‐treated 96‐well plates and allowed to adhere for 24 h. Immediately before use, the 2X Detection Reagent was prepared by diluting each assay component in pre‐warmed complete medium containing 10% FBS. The reagent mixture included Annexin V‐LgBiT, Annexin V‐SmBiT, Annexin V NanoBiT Substrate, and CaCl₂, each at a final dilution of 1:500. All components were mixed by gentle inversion to avoid bubble formation and maintained at RT until added to the experimental wells. Dox and 2X Detection Reagent were added to the plate and incubated at 37°C in a humidified CO₂ incubator. Luminescence was measured at predefined time points (0, 3, 6, 24, and 48 h) using a microplate reader (VICTOR Nivo; PerkinElmer, Shelton, CT, USA). Luminescence, recorded in relative luminescence units (RLU), reflected the binding of Annexin V fusion proteins to externalized phosphatidylserine. Assay was performed as six biological replicates (N = 6).

2.8. Assessment of Senescence With β‐Galactosidase Identification

Cellular senescence in ADSCs was evaluated using β‐galactosidase staining with the Senescence Cells Histochemical Staining Kit (CS0030; Sigma Aldrich, St. Louis, MO, USA) and CellEvent Senescence Green Flow Cytometry Assay Kit (C10841; Thermo Fisher Scientific, Waltham, MA, USA). After a 24 h incubation in Dox at three concentrations, the growth medium was aspirated from each well, and the cells were washed with PBS.

In β‐galactosidase staining with the Senescence Cells Histochemical Staining Kit, fixation buffer was added, and the plate was incubated at RT for 5 min to fix the cells. After fixation, the cells were rinsed three times with PBS to remove fixative. Freshly prepared staining mixture was added to the wells, and the plates were incubated at 37°C in a CO₂‐free atmosphere for 3 h. The cells were observed by light microscopy, and the blue‐stained cells expressing β‐galactosidase were counted as senescent cells. The number of senescent cells and total number of cells were quantified across six images per well to determine the percentage of β‐galactosidase‐positive cells.

In CellEvent Senescence Green Flow Cytometry Assay Kit ADSCs after exposure to Dox were harvested, washed with PBS, and incubated with CellEvent Senescence Green Probe at a dilution 1:1000 for 2 h at 37°C without CO₂. After incubation, cells were washed with PBS, resuspended in assay buffer, and analyzed by flow cytometry. Fluorescence of the activated probe was detected in the FITC channel (excitation/emission: 490/514 nm). A minimum of 10 000 events per sample was acquired. Cell populations were gated based on forward and side scatter characteristics to exclude debris, and senescent cells were analyzed based on median fluorescence intensity (MFI) quantification. Unstained cells and cells treated with Dox were used to establish background fluorescence and define the positivity threshold.

2.9. Determination of Oxidative Stress Parameters

2.9.1. ROS‐Glo H₂O₂ Assay

The ROS‐Glo H₂O₂ Assay (G8820; Promega, Madison, WI, USA) was used to measure hydrogen peroxide (H₂O₂) levels. Cells were seeded at 5 × 10³ cells/well in white culture‐treated 96‐well plates and allowed to adhere for 24 h. The cells were treated with Dox at three concentrations for 24 h. The ROS‐Glo H₂O₂ assay was performed following the manufacturer's instructions. Plates were incubated at 37°C, 5% CO₂, for 6 h, then ROS‐Glo detection solution was added, and the plates were incubated for 20 min at RT. Luminescence was measured using a microplate reader (VICTOR Nivo; Perkin Elmer, Shelton, CT, USA). Control wells included untreated cells and wells without cells to determine background luminescence. The assay was performed as six biological replicates (N = 6).

2.9.2. GSH/GSSG‐Glo Assay

The GSH/GSSG‐Glo assay (V6611; Promega, Madison, WI, USA) was used to measure the levels of reduced glutathione (GSH), oxidized glutathione (GSSG), and total glutathione (GSH + GSSG) and the GSH/GSSG ratio in ADSCs treated with Dox at concentrations 5, 25, and 50 μM. GSH and GSSG both exist in healthy cells, but most glutathione is in the reduced (GSH) form, with only a small percentage in the oxidized (GSSG) form. GSSG is an indicator of cell health and oxidative stress. Usually, a lower ratio GSH/GSSG is related to disease, aging, and the reaction of cells to chemical compounds. Cells were seeded at a density of 5 × 10³ cells/well in white culture‐treated 96‐well plates and allowed to adhere for 24 h, and then treated with Dox at three concentrations for 24 h. After the treatment, the cells were lysed with total or oxidized glutathione reagent and shaken for 5 min. Luciferin generation reagent was added to the wells and incubated for 30 min, followed by the addition of luciferin detection reagent and incubation for 15 min at RT. Luminescence was measured using a microplate reader (VICTOR Nivo; PerkinElmer, Shelton, CT, USA). Controls were untreated cells and wells with medium without cells to determine background. Assay was performed as three biological replicates (N = 3).

2.9.3. MitoSOX Red

MitoSOX Red (M36007; Thermo Fisher Scientific, Waltham, MA, USA) was used for identification of mitochondrial superoxide. It is a fluorogenic dye specifically targeted to mitochondria in live cells. Oxidation of the MitoSOX reagent by mitochondrial superoxide produces red fluorescence. The production of superoxide by mitochondria was visualized in fluorescence microscopy. The assay was performed as three biological replicates (N = 3).

2.10. Cell Cycle Analysis

FxCycle PI/RNase Staining Solution (F10797; Thermo Fisher Scientific, Waltham, MA, USA) was used for flow cytometry analysis of the cell cycle and recognition of the G0/G1, S, and G2/M mitotic phases in the cell populations. After 24 h culturing with Dox at 5, 25, or 50 μM or 0 μM (standard culture medium) for the control group, the cells were fixed in 70% ethanol and incubated with staining solution for 30 min at RT. The samples were analyzed in three biological replicates (N = 3) at 488 nm and 532 nm excitation wavelengths.

2.11. Gene Expression

Relative expression was determined for the following genes related to apoptosis, senescence, oxidative stress, and SAPK/JNK pathway were assessed (Table S1).

Total RNA was extracted from cells using the High Pure RNA Isolation Kit (11828665001; Roche, Basel, Switzerland). RNA concentration was determined using a Nanodrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA).

The relative expression of genes was determined using real‐time RT‐qPCR with SYBR Green chemistry using a GoTaq 1‐Step RT‐qPCR Kit (A6020; Promega, Madison, WI, USA) and a Light Cycler 96 instrument (Roche, Basel, Switzerland). All samples were tested in triplicate. β‐actin was included as the housekeeping gene. Oligonucleotide primers were obtained from Sigma‐Aldrich (Sigma‐Aldrich, St. Louis, MO, USA). Melt curve analysis confirmed the specificity of the primer pairs used for RT‐qPCR (Figure S5). The assay was performed as three biological and six technical replicates (N = 6).

2.12. Protein Expression

Expression of proteins related to apoptosis and the SAPK/JNK pathway was detected in ADSCs in all groups using the Western blot method. The proteins were isolated from cell pellets using RIPA lysis buffer (R0278, 10X; Merck Millipore, Missouri, USA), together with cOmplete Mini Protease Inhibitor Cocktail (11836170001; Roche, Basel, Switzerland) and Protease Inhibitor Cocktail (P1860; Sigma‐Aldrich, St. Louis, MO, USA). Lysates were centrifuged at 14 000 rpm for 15 min at 4°C. The supernatants were mixed with Laemmli Sample Buffer (1610747; Bio‐Rad, Warsaw, Poland) and β‐mercaptoethanol (1610710; Bio‐Rad, Warsaw, Poland) and incubated at 95°C for 5 min.

Protein electrophoresis was performed using 12% polyacrylamide gel, and the separated proteins were transferred to PVDF membrane by wet electroblotting. The membrane was treated for 1 h with non‐fat milk solution in TBST (TBST‐10X, #9997; Cell Signaling, Danvers, MA, USA) as a blocking buffer, followed by overnight incubation at 4°C with the following primary antibodies related to apoptosis: anti‐BAX (50599‐2‐IG; Proteintech, Rosemont, IL, USA), anti‐BCL‐2 (12789‐1‐AP; Proteintech, Rosemont, IL, USA), anti‐cleaved Caspase 3 (9661S; Cell Signaling, Danvers, MA, USA), anti‐cleaved Caspase 9 (7237; Cell Signaling, Danvers, MA, USA), and specific for SAPK/JNK pathway: anti‐c‐Jun (60A8) (9165S3; Cell Signaling, Danvers, MA, USA), anti‐JNK2 (56G8) (9258S; Cell Signaling, Danvers, MA, USA), anti‐phospho‐c‐Jun (Ser63) II antibody (p‐c‐JUN, 9261S; Cell Signaling, Danvers, MA, USA) and anti‐Phospho‐SAPK/JNK (p‐JNK, 9261S; Cell Signaling, Danvers, MA, USA). GAPDH antibody (anti‐GAPDH, G9545; Sigma Aldrich, St. Louis, MO, USA) and β‐actin antibody (anti‐β‐actin, A2066; Sigma Aldrich, St. Louis, MO, USA) were used as reference controls (Table S2). The membranes were then incubated with secondary antibodies (goat anti‐rabbit, ab205718; Abcam, Cambridge, UK) diluted in 5% nonfat milk solution for 1 h at RT. Proteins were detected using chemiluminescence with SuperSignal West Pico PLUS Chemiluminescent Substrate (34578; Thermo Fisher Scientific, Waltham, MA, USA).

Western blot band intensities were quantified using ImageJ software. Equal amounts of total protein were loaded into each lane for all samples. Protein expression was quantified by densitometric analysis (Figure S4). The intensity of each target protein was normalized to the corresponding GAPDH signal and expressed relative to the untreated control group (control = 1).

2.13. Statistical Analysis

Statistical analysis was performed using Statistica 13.3 software. Different groups were compared using the Kruskal–Wallis test for non‐normally distributed data and ANOVA with the post hoc Tukey test for normally distributed data. The level of significance was set at p < 0.05. Values were expressed as median (Me) with the interquartile range (25th–75th percentile) and range (min–max) for non‐normally distributed data, and as mean and standard deviation for normally distributed variables.

3. Results

3.1. Effect of Different Doxorubicin Concentrations on ADSCs Viability

Control cells exhibited the highest viability of ADSCs, as determined by the MTT assay. A significant, dose‐dependent decrease in cell viability was observed with increasing Dox concentrations (ANOVA with post hoc Tukey test). Mitochondrial activity, which correlates with cell viability, was significantly reduced at Dox concentrations of 1 μM (Me = 66.8% ± 12.2%) and higher compared to untreated control cells (Me = 100%). Lower ATP activity was observed at 5 μM Dox (Me = 73.4% ± 3.15%) and higher. The total protein content only decreased at Dox concentrations of 50 μM (Me = 81.9% ± 3.75%) or higher (Figure 1). These findings indicate that cellular metabolic activity, as assessed by two independent mitochondrial‐based assays, declines at relatively low Dox concentrations, whereas a reduction in cell number becomes evident only at higher concentrations. Based on these cytotoxicity results, subsequent experiments were conducted using Dox concentrations of 5, 25, and 50 μM.

FIGURE 1.

Effect of different Dox concentrations (0.1–100 μM) on mitochondrial activity, ATP content, and total protein content in ADSCs, expressed as a percentage of the control cell values (0 μM); means ± SD, *p < 0.05 versus control cells (0 μM); c neg—death cells treated with 1% of Triton X, N = 8. The circles indicate the threshold concentrations above which cellular activity changes significantly.

3.2. Effects of Doxorubicin on the Cell Cycle

Flow cytometry assessment of the ADSC populations in the G0/G1, S, or G2/M phases of the cell cycle (Figure 2) revealed Dox dose‐dependent increases in the percentage of cells in the G0/G1 phase and decreases in the percentage of cells in the G2/M phase.

FIGURE 2.

Flow cytometry analysis of the cell cycle in ADSCs after exposure to Dox at 5, 25, or 50 μM for 24 h, *p < 0.05 versus control (0 μM), ^# p < 0.05 versus 5 μM. (A) Mean percentage of ADSCs representing different phases of the cell cycle; mean ± SD, N = 3. (B) Histograms showing the distributions of ADSCs in different phases of the cell cycle.

At lower concentrations (5 μM), only moderate shifts in cell cycle distribution were observed. The percentage of cells in the G2/M phase in the control group (Me = 15.61% ± 1.4%) was three times higher than in cells treated with Dox at a concentration of 5 μM (Me = 5.36% ± 0.4%). Higher concentrations (25–50 μM) led to pronounced G1/G0 accumulation and substantial depletion of cells in G2/M phase, for 25 μM (Me = 0.55% ± 0.1%) and for 50 μM Dox (Me = 0.33% ± 0.4%). Treatment with 50 μM Dox resulted in the highest percentage of cells in the S phase (Me = 7.8% ± 2.7%) compared to other groups (ANOVA, post hoc Tukey).

3.3. Morphological and Structural Changes in ADSCs

3.3.1. Dox Changes ADSC Morphology

Bright field microscopy examination of unstained ADSCs after a 24 h Dox incubation revealed notable changes in ADSC morphology (Figure 3A). Most ADSCs preserved their fibroblast‐like shape, and they adhered to the culture plates, but some cells showed vacuolization and detachment. In cells treated with Dox shrinking was observed, leading to a decrease in cell volume especially in cells cultured in Dox at 25 and 50 μM. Some cells may also increase in size due to swelling prior to membrane rupture and cell death. A similar effect was observed in ADSCs treated with the highest Dox (50 μM) concentration.

FIGURE 3.

Morphological and structural changes in ADSCs. (A) Morphology of ADSCs after 24 h exposure to Dox (5, 25, or 50 μM) compared to control cells (0 μM), (magn. 100×; magn. in the inset: 200×, scale bar = 50 μm). (B) F‐Actin filament arrangement ADSCs stained with Alexa Fluor 555 coupled phalloidin (red) after 24 h exposure to Dox at 5, 25, or 50 μM compared to control cells (0 μM). Cell nuclei were visualized with DAPI (blue), magnification 400×, scale bar = 20 μm. (C) Quantitative analysis of ADSCs size (μm) after Dox treatment. Data were derived from 6 independent photos. Mean ± SD, *p < 0.05 versus control (0 μM), N = 6.

Quantitative analysis, analyzed on cells after phalloidin staining, demonstrated a reduction in cell size with increasing concentrations of Dox (Figure 3C, ANOVA, post hoc Tukey). This observation suggests that cells treated with 50 μM Dox may have undergone apoptosis, as cellular shrinkage is a characteristic morphological feature of this process (Figure S3).

3.3.2. Disruption of Cytoskeleton Organization

F‐actin filaments determine the structure of the cytoskeleton in cells. F‐actin staining revealed long, well‐organized filaments in the control cells, with stress fibers continuously distributed throughout the cytoplasm. Cells treated with 5 μM Dox showed relatively well‐organized F‐actin filaments; however, some cells contained actin aggregates, suggesting early cytoskeletal degradation. The filaments in the cells treated with 5 μM Dox also appeared shorter and were more prominently localized at the cell periphery rather than uniformly distributed across the entire cell. The ADSCs incubated with higher Dox concentrations (25 and 50 μM) exhibited pronounced actin disorganization characterized by the presence of clumps of degraded actin. F‐actin filaments were predominantly detected in the perinuclear region and appeared highly fragmented and shortened. Cell boundaries were irregular, and cellular degradation, including loss of membrane continuity, was evident (Figure 3B).

3.4. Assessment of Apoptosis in ADSCs

Apoptosis was examined in ADSCs treated with Dox using detection of Annexin V, the expression of genes, and proteins related to apoptosis.

3.4.1. Dox Induces Time‐Dependent Apoptosis in ADSCs

Annexin V‐based measurements of phosphatidylserine externalization revealed a clear temporal and concentration‐dependent induction of apoptosis in ADSCs treated with Dox. During the early phase (0–6 h), no significant increase in apoptotic signal was detected at any tested concentration, indicating that Dox does not trigger immediate apoptotic responses (Figure 4A,B). At 24 h, apoptosis was significantly elevated in cells exposed to higher Dox concentrations (25 and 50 μM), as evidenced by increased luminescence values compared to control (ANOVA, post hoc Tukey). Lower concentration (5 μM) showed only a modest, non‐significant increase at this time point. After 48 h, all Dox‐treated groups, including 5 μM, exhibited increased apoptotic activity, confirming a delayed but progressive effect.

FIGURE 4.

Assessment of apoptosis in ADSCs (A) Kinetic profiles of phosphatidylserine translocation related to apoptosis in ADSCs after exposure to 5, 25, or 50 μM Dox for 0, 3, 6, 24, and 48 h, *p < 0.05 versus control (0 μM), N = 6. (B) Comparison of phosphatidylserine translocation related to apoptosis in ADSCs after 24 h and 48 h of exposure to Dox. Means ± SD, *p < 0.05 versus control (0 μM). The circles indicate the threshold concentrations above which cellular activity changes significantly. (C) Expression of Bax, Bcl‐2, TP53, CASP3, and CASP7 genes in ADSCs after exposure to Dox in comparison to control (0 μM). Means ± SD, *p < 0.05 versus control (0 μM), N = 6. (D) The Bax/Bcl‐2 ratio. (E) The expression of cleaved Caspase 3, cleaved Caspase 9, BAX, and BCL‐2 proteins in ADSCs after exposure to Dox versus control cells. Means ± SD, *p < 0.05 versus control (0 μM), N = 4. (F) Representative Western blot results for the analyzed proteins.

Because Annexin V detects phosphatidylserine exposure without distinguishing between apoptotic pathways, complementary analyses (Western blotting and RT‐qPCR) were performed to further characterize the underlying mechanisms (Figure 4A,B).

3.4.2. Doxorubicin Changes the Expression of Apoptosis‐Related Genes

Bax expression was greater in cells treated with Dox at 25 and 50 μM (ANOVA, post hoc Tukey) than in control cells, and Bcl‐2 expression was lower in cells treated with Dox at all three Dox concentrations (5, 25, and 50 μM).

The Bax/Bcl‐2 ratio, which indicates the susceptibility of cells to apoptosis, was 3.5, 2.9, and 1.8 in cells treated with 5, 25, and 50 μM, respectively, compared to the control cells, suggesting a greater tendency toward apoptosis at lower Dox concentrations than at higher concentrations. A statistically significant decrease in the expression of TP53 and CASP3 in cells treated with all three Dox concentrations was also observed. No changes were detected in CASP7 mRNA levels (Figure 4C,D).

3.4.3. Dox Upregulates the Expression of Apoptosis‐Related Proteins

Based on the above results we conducted Western blot analysis of proteins associated with apoptosis, expression of cleaved caspase 3 and cleaved caspase 9 after treatment with Dox at 5, 25, 50 μM concentrations (ANOVA, post hoc Tukey). The expression of the p21 isoform, proapoptotic form of the Bax protein, was higher in cells treated with Dox at 5 and 25 μM, while the relative expression of isoform p18 and Bcl‐2 was lower (Figure 4E,F).

All three methods described above confirmed that Dox at a concentration of 5 μM or above induces apoptosis in ADSCs.

3.5. Assessment of Senescence in ADSCs

The induction of senescence following Dox treatment was assessed by histochemical detection of senescence‐associated β‐galactosidase (SA‐β‐gal), flow cytometric analysis of β‐galactosidase activity using the CellEvent Senescence Green assay, and evaluation of the expression of the senescence‐related gene RB1.

3.5.1. β‐Galactosidase Analysis Indicates That Dox Promotes Senescence

Histochemical analysis demonstrated that Dox treatment significantly increased the proportion of SA‐β‐gal‐positive cells compared with the untreated control (16.55% ± 4.95%). The percentage of positive cells increased to 37.97% ± 11.81%, 41.16% ± 3.06%, and 61.44% ± 13.95% following treatment with 5, 25, and 50 μM Dox, respectively (Figure 5A,B, ANOVA, post hoc Tukey).

FIGURE 5.

Assessment of senescence and oxidative stress in ADSCs (A) β‐galactosidase staining in ADSCs after exposure to 5, 25, and 50 μM Dox; Representative images. Magn. 200×, scale bar = 50 μm. (B) Quantification of SA‐β‐gal‐positive cells expressed as percentage of total number of cells, mean ± SD, *p < 0.05 versus control (0 μM), N = 8. (C) Senescence‐associated β‐galactosidase activity after exposure to 5, 25, and 50 μM Dox analyzed with flow cytometry. (D) Expression of RB1 gene in ADSCs after exposure to Dox, median and quartiles; *p < 0.05 versus control (0 μM), N = 6. (E) ROS generation in ADSCs after exposure to 5, 25, and 50 μM Dox, means ± SD, *p < 0.05 versus control cells (0 μM), ^# p < 0.05 versus 5 μM, N = 8. (F) Expression of CAT, SOD1, SOD2, and GPX4 genes in ADSCs after exposure to Dox, medians and quartiles; *p < 0.05 versus control cells (0 μM), N = 8. (G) GSH/GSSG ratio in ADSCs after exposure to 5, 25, and 50 μM Dox. Means ± SD, *p < 0.05 versus control cells (0 μM), N = 3. (H) Mitochondrial superoxide identification in ADSCs after exposure to Dox. Representative images. Magn. 100×, scale bar = 20 μm.

Changes in β‐galactosidase activity were further evaluated by flow cytometry using the CellEvent Senescence Green assay. Probe fluorescence was analyzed in the fluorescein (FITC) channel. Because adipose‐derived stem cells (ADSCs) exhibit intrinsic autofluorescence and doxorubicin itself emits fluorescence that contributes to background signal in the FITC channel, fluorescence measurements were corrected using the corresponding unstained control for each experimental condition. The CellEvent Green signal was expressed as the background‐subtracted geometric mean fluorescence intensity (ΔgMFI), calculated according to the following equation: ΔgMFI = g MFI stained sample − g MFI corresponding unstained sample. Calculations were performed separately for the untreated control and for each doxorubicin concentration. The relative change in fluorescence signal was expressed as fold change relative to the untreated control and calculated as follows: Fold change = ΔgMFI untreated control/ΔgMFI treated sample (Figure 5C, Figure S3).

Both assays consistently demonstrated that Dox induces senescence in ADSCs. Histochemical staining revealed a dose‐dependent increase in the proportion of SA‐β‐gal‐positive cells, whereas flow cytometric analysis confirmed enhanced β‐galactosidase activity following Dox treatment, with the strongest fluorescence signal observed at 25 μM Dox (Figure 5C).

3.5.2. Dox Do Not Change the Expression of Senescence‐Related Genes

The expression of RB1 gene was higher only in cells treated with 5 μM Dox, but the difference was not statistically significant (Figure 5D, Kruskal–Wallis with multiple comparisons, medians and quartiles).

3.6. Assessment of Oxidative Stress in ADSCs

Our analysis included the level of ROS, the reduced glutathione (GSH)/oxidized glutathione (GSSG) ratio, and the expressions of oxidative stress‐related genes (CAT, SOD1, SOD2, and GPX4).

3.6.1. Dox Induces ROS Production

Dox promoted a statistically significant increase in ROS production only at 25 μM (Me = 7907 ± 349) and 50 μM (Me = 7907 ± 349) compared to the control cells (Me = 6231 ± 184). The ROS production in response to 5 μM Dox (Me = 6995 ± 200) was not statistically significant compared to the controls (Figure 5E, ANOVA, post hoc Tukey).

3.6.2. Dox Do Not Alter the Glutathione Ratio in ADSCs

No GSH/GSSG ratio imbalance was evident in cells after 24 h Dox exposure at any concentration, as none of the values were statistically different from the control group (Figure 5G, ANOVA, post hoc Tukey).

3.6.3. Dox Changes the Expression of Oxidative Stress–Related Genes

Expression of the GPX4 gene was lower in cells treated with Dox only with 25 μM Dox. SOD1 and CAT1 showed higher expression in cells treated with 5 μM Dox and declined in cells treated with 25 μM Dox. SOD2 expression did not change in response to Dox treatment (Figure 5F, Kruskal–Wallis with multiple comparisons, medians and quartiles).

3.6.4. Dox Promotes Mitochondrial Superoxide

Results confirm that levels of mitochondrial superoxide increased primarily at 25 and 50 μM. We did not detect mitochondrial superoxide in control cells and cells treated with 5 μM (Figure 5H).

3.7. Effects of Dox on the SAPK/JNK Pathway Activity

3.7.1. Doxorubicin Changes the Expression of SAPK/JNK Pathway‐Associated Genes

The expression of MAP3K2 was similar in the control group and in cells treated with 5 μM Dox but was lower in cells treated with 25 μM and 50 μM Dox compared to the 5 μM group (ANOVA post hoc Tukey). The relative expression of MAP2K7 was lower in all the Dox‐treated groups compared to control cells. The level of JUN was lower in cells treated with 25 and 50 μM Dox compared to cells treated with 5 μM. No statistically significant changes were noted in MAP2K4 expression. However, the expression profile was similar to that of the other genes, with similar expression detected in control cells and in cells treated with 5 μM Dox, and lower expression observed in cells treated with 25 and 50 μM Dox. The expression of MAP3K1 did not change in any of the groups (Figure 6A).

FIGURE 6.

Effects of Dox on SAPK/JNK pathway (A) Expression of the MAP3K2, MAP2K4, MAP2K7, JUN, and MAP3K1 genes in ADSCs exposed to Dox (5, 25, and 50 μM) compared to control cells (0 μM). Means ± SD, *p < 0.05 versus control (0 μM), ^# p < 0.05 versus 5 μM, N = 6. (B) Comparison of the expression of JNK2, c‐JUN, p‐JNK in ADSCs after exposure to Dox versus control cells. Means ± SD; *p < 0.05 versus control (0 μM), ^# p < 0.05 versus 5 μM, ^p < 0.05 versus 25 μM, N = 4. (C) Representative Western blot results for the analyzed proteins.

3.7.2. Dox Changes the Expression of SAPK/JNK Pathway‐Associated Proteins

The expression of both the p46 and p54 isoforms of p‐JNK protein was higher in all the examined groups compared to the untreated controls (0 μM) (ANOVA post hoc Tukey). The expression of p‐JNK was higher in cells after incubation with 50 μM Dox than at lower concentrations (5 and 25 μM), suggesting a dose‐dependent response. JNK2 expression was significantly increased in all Dox‐treated groups compared with control cells, while c‐JUN expression was significantly reduced in cells treated with Dox at all concentrations (Figure 6B,C). No expression of the p‐c‐Jun protein was detected in any of the groups. These data confirmed the role of the SAPK/JNK pathway in response to Dox toxicity in the ADSC.

4. Discussion

Dox is a chemotherapeutic drug used as a therapy for many types of cancer. However, its use in cell‐based cancer therapy might be limited due to its negative effects on adult stem cells. At present, little is known regarding Dox's effects on ADSCs, despite the increasing popularity of adipose tissue as a source of cancer‐relevant therapeutic stem cells. The hypothesis proposes that exposure to Dox results in ADSCs damage, which is mediated through the activation of the SAPK/JNK signaling pathway. The aim of the present study was to determine whether Dox has potentially adverse effects on ADSCs intended for therapeutic use.

We found that the cytotoxicity effects of Dox varied depending on its concentration, which was tested from 0.1 to 100 μM. In patients, the therapeutic concentration for Dox is 5–100 μM [22] short after administration, and the highest concentration of Dox observed in the plasma of patients following intravenous administration after 24 h and more ranges from 1 μM to 30 μM [13, 23, 24]. Effective doxorubicin (Dox) concentrations were determined based on cytotoxicity studies in cancer cell lines exhibiting variable drug sensitivity, with IC₅₀ values reported to range from 1 μM to 400 μM depending on cancer type [24]. In this study, Dox showed a significant dose‐dependent reduction in the viability of ADSCs, indicating its strong cytotoxic potential in this noncancerous cell population. We proved here that even low concentrations of Dox (1–5 μM) adversely affected ATP production and mitochondrial activity in the ADSCs at clinically relevant concentrations. Although the 5 μM concentration of doxorubicin is considered to be within the clinically relevant range, the higher concentrations were also used in this study, which represent conditions of acute experimental toxicity rather than typical systemic exposure. Local drug concentrations may differ substantially from circulating plasma levels depending on tissue distribution, drug accumulation, and the mode of drug delivery systems, for example, biomaterial‐based carriers, where local concentrations may exceed those achieved during systemic administration. Higher doses were included to characterize dose‐dependent cellular responses and to define the threshold at which changes are observed in aspects of stem cells. Future studies using clinically relevant exposure models, including prolonged treatment with lower doxorubicin concentrations, are warranted to further define the translational significance of these findings.

Dox also altered the ADSCs cell structure, evident as reductions in the lengths and regularity of the actin filaments, which are responsible for the movement and division of cells. Changes in the length and arrangement of actin fibers are indicators of cell dysfunction, and shorter fibers are associated with reduced cell adhesion [25]. In our study, the Dox‐treated ADSCs displayed aggregates of actin at the periphery of the cell as well as irregular protrusions at the cell edges. Actin aggregates have been shown to be associated with broken actin filaments, suggesting that the present observations indicated the disruption of actin filaments, which often occurs when actin polymerization is blocked [26]. These findings suggest that clinical doses of a chemotherapeutic agent might decrease the potential for cell survival in stem cell niches after cancer treatments. Disturbances of the cytoskeleton might also interfere with cell adhesion and cell division processes that are essential for stem cell cytophysiology [27]. At 5 μM, Dox induced cell cycle arrest at the G2/M phase. A similar outcome has also been reported by Kozhukharova et al. [28] following a 48‐h treatment of human MSCs originating from menstrual blood, bone marrow, and adipose tissue with 0.1 μM Dox. In the present study, higher concentrations of Dox (25 and 50 μM) decreased the number of cells in the G2/M phase, indicating the occurrence of severe DNA damage and disturbances in cell cycle checkpoints, what suggests the induction of replication stress and impaired progression through the cell cycle. These observations agree with data obtained from cancer cell studies for these concentrations [29]. However, all presented until now results proved that clinical used concentration of Dox (5 μM) inhibits proliferative potential of ADSCs.

An important observation from this study is the dual ability of Dox to act as a pro‐senescence and a pro‐apoptosis factor on ADSCs. Much of the published research has shown that senescent cells exhibit resistance to apoptosis [30, 31]. Programmed cell death's function is to maintain internal homeostasis by eliminating damaged cells, also after drug treatment [15]. Consequently, resistance to apoptosis and promotion of premature senescence can have unfavorable impacts, such as promotion of oxidative stress and inflammation, which can impede cell recovery used for regenerative therapy. We proved that Dox has pro‐apoptotic effects in the ADSCs, as evidenced by increased cleavage of caspase‐3 and caspase‐9, upregulation of pro‐apoptotic BAX, and downregulation of anti‐apoptotic BCL‐2, indicating activation of the intrinsic apoptotic pathway. Mitochondrial damage likely promotes cytochrome c release, leading to caspase‐9 activation and subsequent activation of caspase‐3 [32, 33]. This process is tightly regulated by the BCL‐2 family, with BAX playing a key pro‐apoptotic role. Notably, apoptosis exhibited both dose‐ and time‐dependent dynamics. We observed increased phosphatidylserine externalization after prolonged (> 24 h) exposure to low, clinically relevant Dox concentrations (5 μM), suggesting that sustained treatment at this dose enhances apoptotic processes. These findings indicate that during anticancer therapy, when Dox persists in plasma, ADSCs may be susceptible to damage. Similarly, treatment of ADSCs with 5 μM Dox resulted in increased senescence‐associated β‐galactosidase activity, suggesting the induction of premature cellular senescence. Interestingly, low concentrations of Dox did not induce ROS and mitochondrial superoxide production comparable to higher doses, and ADSCs maintained a normal glutathione ratio, suggesting preserved oxidative homeostasis. Despite increased ROS generation at 25 and 50 μM, the glutathione redox balance remained largely stable, indicating that compensatory antioxidant mechanisms may still operate under these conditions. Higher Dox concentrations might reduce gene expression, what is consistent with reports in many studies about inhibition of antioxidant enzymes [34, 35], like in Dox‐induced cardiotoxicity [36]. Notably, this effect appeared to involve only selected genes (CAT and GPX4). This effect may be related to the different activity of ADSCs, especially paracrine potential [37]. Their secretome consists of antioxidant proteins that may partially counteract Dox‐induced oxidative damage [38]. These findings indicate that Dox‐induced oxidative stress became evident primarily at higher concentrations, while the cellular antioxidant system was still able to maintain overall redox homeostasis under the experimental conditions through 24 h treatment at clinical doses.

We hypothesized that the observed cellular changes may be associated with the SAPK/JNK pathway, which is known to regulate stress responses, apoptosis, and chemoresistance [17, 18]. Analysis of pathway‐related gene and protein expression supported its role as a mediator of the effects observed in this study. JNK kinases, also known as stress‐activated protein kinases (SAPKs), belong to the MAPK family and are involved in regulating cell proliferation and death. In humans, three isoforms (JNK1–3) have been identified. Activation of this pathway involves MAP2K enzymes (MKK4 and MKK7), which phosphorylate JNK proteins, and MAP3K kinases (MEKK1 and MEKK2), which are activated under stress conditions, including ROS exposure. Phosphorylated JNK subsequently activates c‐JUN, leading to AP‐1 activation and regulation of apoptosis and stress responses [17].

In our study, MAP3K, MAP2K4, and JUN expression remained unchanged in the untreated control cells and in cells treated with 5 μM Dox, but its expression was significantly reduced at the higher Dox concentrations (25 μM and 50 μM). In contrast, MAP2K7 expression was consistently reduced across all treatment groups, suggesting that Dox suppresses upstream regulators of JNK signaling, even at low doses. SAPK/JNK pathway proteins analysis revealed that even though pathway gene expression was reduced, JNK2 and phosphorylated JNK (p‐JNK) levels were increased, indicating pathway activation, alongside decreased c‐JUN expression at 24 h. These findings suggest that, at this time point, Dox predominantly changes JNK signaling at the protein level and this was consistent with the observed apoptosis, senescence, and mitochondrial defects in the cells. The expression of proteins observed here, especially the phosphorylated form of JNK, indicated that the JNK pathway is activated. Decreased c‐JUN expression and the lack of phospho‐c‐JUN expression could be a consequence of protein degradation or inhibitory feedback following JNK pathway activation. Some studies suggest that persistent JNK pathway activity in epithelial cells induces senescence and cell cycle arrest while also increasing ROS production [30]. Other research has shown that inhibiting the JNK pathway reduces cell proliferation and causes a loss of the pluripotent phenotype in induced pluripotent stem cells [19]. Overall, the current evidence indicates an important role of this pathway in stem cell activity.

Taken together, our results demonstrate that Dox, applied at clinically relevant concentrations, exerts broad cytotoxic effects on human ADSCs and affects key aspects of their viability, metabolic activity, cytoskeletal integrity, cell cycle regulation, and fate, including senescence and apoptosis. These deleterious effects are closely associated with changes in the SAPK/JNK signaling pathway, suggesting that stress‐activated protein kinase signaling plays a central role in mediating Dox‐induced stem cell dysfunction. These findings suggest that future research should focus on preventing stem cell quality loss and elucidating the molecular mechanisms underlying Dox effects on ADSCs. Strategies such as genetic modification of MSCs and targeted modulation of the JNK pathway may enhance the efficacy of stem cell‐based therapies [39]. Clinically, these findings raise concerns about the regenerative potential of ADSCs from patients previously treated with Dox. They emphasize the need for improved approaches—such as optimized cell preparation, preconditioning, and targeted therapies—to maintain stem cell function. Overall, integrating stem cell safety into cancer treatment and regenerative medicine is essential.

Unfortunately, the study has some limitations that must be considered. Study demonstrates an association between Dox‐induced cytotoxicity and alterations in SAPK/JNK signaling. However, functional validation experiments were not conducted, and a causal role of the SAPK/JNK pathway cannot be concluded. Future studies as pharmacological JNK inhibition or siRNA‐mediated knockdown are required to determine the contribution of this pathway to the cellular response of ADSCs to doxorubicin. Moreover, this study was performed using commercially available primary ADSCs derived from a single healthy donor. Although the use of a single‐donor cell population allowed the establishment of a controlled experimental model with reduced donor‐dependent variability, it also limits the generalizability of the findings. ADSCs exhibit donor‐dependent differences in proliferation, differentiation potential, and responses to stress. Therefore, future studies should validate these observations using cells obtained from multiple donors with diverse biological and clinical characteristics.

5. Conclusions

Dox exerts multifaceted cytotoxic effects on ADSCs at clinically relevant doses. Alterations in expression of the SAPK/JNK pathway can be detected at a Dox concentration of 5 μM and higher, corresponding to the physiological levels of Dox routinely used in cancer therapy. At this concentration, the earliest signs of apoptosis, decreased cell viability, reduced mitochondrial activity, and cytoskeletal disorganization are evident. Collectively, the results presented here highlight the need to broaden clinical and laboratory protocols concerning the use of ADSCs.

Author Contributions

A.S.‐S. conceived and designed the experiments; A.S.‐S., K.P.‐M., A.B. and E.B. performed the experiments; A.S.‐S. and B.S. analyzed the data, A.S.‐S., B.S. and P.C. wrote the manuscript; A.S.‐S. and P.C. contributed reagents/materials/analysis tools.

Funding

This study was supported by The National Science Centre (NCN) grant number 2023/07/X/NZ3/00379 and Ministry of Science and Higher Education, Student Science Clubs Create Innovation grant number SKN/SP/601468/2024 and by Medical University of Silesia in Katowice (Poland) grants number BNW‐2‐062/K/5/I, BNW‐2‐013/K/4/I and BNW‐2‐061/K/3/I.

Ethics Statement

The study protocol was approved by the Bioethics Committee of the Medical University of Silesia—decision no. PCB/CBN/0052/KB/211/22.

Consent

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Expression of mesenchymal stem cell markers—CD73, CD90, CD105 for confirmation MSC phenotype before conducted experiments. UC (red)—unstained control, IC (green)‐isotype control, antibodies (pink). FACS histogram representative of CD73+, CD90+ and CD105+ cells. Results are present as percentage of positive cells (CD73 = 95.98%, CD90 = 98.24%, and CD105 = 99.62%).

Figure S2: Multipotency assessment of ADSCs. Cytochemical staining of calcium deposits (Alizarin Red S), acid mucopolysaccharides (Toluidine Blue), and intracellular lipid droplets (Oil Red O), confirming osteogenic, chondrogenic, and adipogenic differentiation, respectively. Scale bars represent 20 μm.

Figure S3: Quantitative analysis ADSCs size after Dox‐treatment expressed in pixels (μm). Staining with Alexa Fluor 555 coupled Phalloidin (red). Cell nuclei were visualized with DAPI (blue). Magnification 200×. Results presented in main manuscript.

Figure S4: Full, uncropped Western blot membranes used for densitometric quantification.

Figure S5: Melt curve analysis confirming the specificity of the primer pairs used for RT‐qPCR. Representative melting curves for all primer pairs demonstrated a single peak, indicating the amplification of a single specific PCR product without detectable nonspecific products or primer‐dimer formation. These results confirm the specificity of the primers used for gene expression analysis.

Table S1: Specific primers used in RT‐qPCR.

Table S2: Specific antibodies used for western blot and flow cytometry analysis.

Data Availability Statement

Any raw data of presented results is made available upon justified request addressed to the corresponding author.