Next Article in Journal
Cuproptosis and Diabetic Osteoporosis: Mechanisms and Therapeutic Prospects
Next Article in Special Issue
Mitochondria-Targeted Biophysical Priming of Autologous Biologics for Skin Regeneration and Wound Repair
Previous Article in Journal
Hereditary Endometrial Cancer: Lynch Syndrome, Mismatch Repair Deficiency, and Emerging Genetic Predispositions—A Comprehensive Review with Clinical and Laboratory Guidelines
Previous Article in Special Issue
Clinical Insights into Mesenchymal Stem Cell Applications for Spinal Cord Injury
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
IJMS
Volume 27
Issue 3
10.3390/ijms27031303
ijms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Cardiosphere-Derived Cells from Not Dilated and Dilated Human Myocardium Exhibit Enhanced Metabolic Potential Compared with Conventional Cardiac Mesenchymal Stem/Stromal Cells

by
Daiva Bironaite
* and
Rokas Mikšiūnas
Department of Regenerative Medicine, State Research Institute Centre for Innovative Medicine, LT08406 Vilnius, Lithuania
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2026, 27(3), 1303; https://doi.org/10.3390/ijms27031303
Submission received: 14 November 2025 / Revised: 16 January 2026 / Accepted: 23 January 2026 / Published: 28 January 2026
(This article belongs to the Special Issue Mesenchymal Stem/Stromal Cells: Mechanisms and Applications in Tissue Regeneration)
Browse Figures
Versions Notes

Abstract

Dilated cardiomyopathy (DCM) is a major contributor to heart failure and cardiac transplantation. This study investigated the metabolic potential of human myocardium-derived mesenchymal stem/stromal cells (hmMSCs) and subsequently cardiac sphere-derived cells (SDCs) obtained from healthy (non-dilated) and pathological (dilated) myocardial tissues. hmMSCs were isolated using the explant outgrowth method and expanded in monolayer culture. Small round cells loosely attached on hmMSCs were harvested and cultivated as cardiac spheroids for 1–3 days, subsequently obtaining SDCs. The cell morphology, proliferation rate, mitochondrial activity, and intracellular calcium levels were analyzed using flow cytometry, Seahorse metabolic assays, and spectrophotometry, while expression of cell progenitor and cardiac commitment genes were analyzed by quantitative PCR. Both healthy and pathological SDCs demonstrated significantly enhanced mitochondrial function—reflected by increased maximal respiration, ATP production, and coupling efficiency—along with reduced steady-state intracellular calcium levels compared with hmMSCs. SDCs from both healthy and dilated myocardium showed marked upregulation of several cardiac progenitor and lineage-commitment genes relative to hmMSCs. SDCs derived from both healthy and dilated myocardiums possess a more favorable metabolic, progenitor and cardiac commitment profile than conventional hmMSCs. hmMSCs and SDCs from dilated myocardium retain residual metabolic potential, which may be further enhanced under 3D culture conditions.

1. Introduction

Heart failure (HF) represents a major global health concern, characterized by increasing prevalence, escalating healthcare costs, and a rising incidence worldwide. Each year, over one million hospitalizations occur in the United States and Europe due to common cardiac diseases [1]. Dilated cardiomyopathy (DCM), a predominant cause of HF, is associated with high morbidity and mortality rates, making it the leading indication for cardiac transplantation [2]. Traditional pharmacological interventions primarily slow disease progression, leaving heart transplantation as the only curative option for advanced HF. Therefore, there is a pressing need to search for novel therapeutic approaches capable of preventing heart failure at the stage of dilated myopathy and improving its regeneration.
The heart was long considered a terminally differentiated organ with minimal regenerative capacity [3]. However, the discovery of cardiac-resident stem-like cells expressing c-KIT and other progenitor markers revealed an intrinsic, though limited, regenerative potential [4,5]. Because c-KIT+ and Sca1+ populations comprise less than 1% of total cardiac cells, human myocardium-derived MSCs (hmMSCs) have emerged as a promising alternative for cardiac repair promoting regeneration primarily through paracrine signaling, stimulating endogenous repair, modulating immune responses, reducing fibrosis, and/or by limited direct differentiation to cardiomyocytes [4,5]. Although rare transdifferentiation events into cardiomyocytes or vascular cells have been reported [6], clinical translation of hmMSCs remains challenging. This highlights the need for novel strategies—such as genetic, epigenetic, biomaterial-based, or culture-driven approaches—to enhance the regenerative capacity of hmMSCs.
One promising approach to improving the cardiac regenerative potential can be application of cardiospheres and cardiac-sphere-derived cells (SDCs) [7]. These spheroid-derived cells are clonogenic, express stem, endothelial and mesenchymal progenitor cell markers, exhibit self-renewal capacity, and can differentiate into cardiomyocytes and vascular cells both in vitro and in vivo [7]. Over the past decades, cardiac-sphere-derived cells have emerged as one of the most promising allogeneic cell sources for cardiac therapy [8]. SDCs display a heterogeneous gene expression profile, including high levels of stemness-associated genes such as c-KIT and MDR1, alongside cardiac-specific genes such as Nkx2.5, while showing lower expression of mature cardiomyocyte markers [9]. Moreover, SDCs exert cardioprotective, antifibrotic, and immunomodulatory effects [10,11,12,13], but despite evidence of safety following allogeneic SDC injection in post-infarction rat hearts, the risk of immune rejection in humans remains, and cell efficacy may also be undermined by donor age and comorbidities such as myopathies and others [14,15]. Collectively, these findings highlight the importance of SDCs, while underscoring the need for deeper insight into their metabolic biology, especially in healthy versus pathological/dilated human myocardiums.
Efficient cellular metabolism is essential for proper cardiac function, ensuring a continuous supply of oxygen and nutrients. Recent studies have shown that glucose and lactate levels are elevated, while free fatty acid levels decline in hearts with end-stage dilated cardiomyopathy [16]. This finding supports the theory that failing hearts undergo a metabolic shift from oxidative phosphorylation to glycolysis, driven by factors such as impaired mitochondrial dynamics, altered transcriptional regulation of mitochondrial proteins, and other metabolic disturbances [17]. Furthermore, recent metabolomic and transcriptomic advances show that, beyond morphogens and signaling factors, key metabolic pathways—especially oxidative phosphorylation and glycolysis—are crucial for stem cell self-renewal and differentiation [18]. These insights underscore the need to explore novel models for targeted metabolic regulation of human myocardium-derived cells to enhance cardiac regeneration.
Therefore, the aim of this study was to investigate the morphology, proliferation, and metabolic activity, isolated from both healthy and dilated myocardium in comparison to regular hmMSCs. The cardiac commitment genes between human healthy and pathological cardiospheres and SDCs of different passages were also determined. To our knowledge, this is the first study to demonstrate that healthy and dilated myocardium-derived SDCs enhances mitochondrial activity and promotes a more cardiac commitment phenotype, particularly in dilated myocardium-derived SDCs compared with conventional 2D-cultured hmMSCs.

2. Results

2.1. Isolation and Size Evaluation of Healthy/Not Dilated and Pathological/Dilated Myocardium-Derived hmMSCs and SDCs

The hmMSCs were isolated from healthy and dilated human ventricular myocardium biopsies using the explant outgrowth method. Myocardial biopsies were minced into ~1 mm3 fragments, and within 1–3 weeks, hmMSCs migrated out of the tissue and adhered to the fibronectin-coated surface (Figure 1a). The cells were subsequently expanded and used for further experiments (Figure 2c). Our previous studies demonstrated that human heart-derived hmMSCs express typical mesenchymal stem cell surface markers CD90, CD73, and CD105; are negative for CD45, CD10, and CD13; and are capable of differentiating into adipogenic, osteogenic, and chondrogenic lineages [19,20]. In addition, hmMSCs exhibited a tendency toward cardiomyogenic differentiation [19,20].
During explant cultivation, bright, round cells that were loosely attached to the hmMSCs were collected and cultured under three-dimensional (3D) conditions on poly(2-hydroxyethyl methacrylate) (pHEMA)-coated plates using a specialized cardiosphere growth medium (Figure 1b). After three days of cultivation, the resulting cardiospheres were dissociated, and the sphere-derived cells were seeded for further analysis (Figure 1d). Both healthy and pathological SDCs were smaller in size (Figure 1d) compared with their corresponding hmMSCs (Figure 1c,d).
More detailed cell attachment measurements revealed that SDCs from both healthy and pathological samples were approximately twofold smaller in size compared with their corresponding hmMSCs (Figure 2a,b). The cell attachment area also differed between SDCs and hmMSCs in both groups; specifically, the attachment area of healthy SDCs was about twofold smaller than that of healthy hmMSCs, whereas pathological SDCs exhibited an approximately fourfold smaller attachment area compared with pathological hmMSCs (Figure 2c,d). In addition, the attachment area between healthy and pathological hmMSCs differed significantly, as reported previously [20] (Figure 2d). These data suggest that both types of SDCs derived from healthy and DCM myocardium exhibited a reduction in size, which might be associated with a higher cardiac commitment and/or progenitor-like phenotype.

2.2. Proliferation of Healthy and Pathological hmMSCs and SDCs

The proliferation of healthy and pathological hmMSCs and SDCs was measured using the Cell Counting Kit-8 (CCK-8) assay (Dojindo Molecular Technologies, Inc., Rockville, MD, USA) (Figure 3). In addition, cell proliferation evaluation by direct cell counting showed similar results (Supplementary Materials). Over six days of cultivation, pathological hmMSCs exhibited slightly lower proliferation rates compared with healthy hmMSCs, whereas both healthy and pathological SDCs proliferated significantly faster than their corresponding hmMSCs and showed exponential growth tendency. Overall, the proliferation rate of healthy and pathological SDCs increased approximately twofold compared with the respective hmMSCs.
These data also suggest that healthy hmMSCs and SDCs retained higher initial energetic resources required for proliferation than pathological cells. Both types of SDCs require an adaptation period to 2D growth conditions to achieve an increased proliferative rate.

2.3. Mitochondrial Membrane Potential (MMP) and iCa2+ of Healthy and Pathological hmMSCs and SDCs

Mitochondrial membrane potential (MMP) and intracellular calcium (iCa2+) levels in healthy and pathological hmMSCs and SDCs were evaluated by staining the cells with JC-1 dye and Cal-520, respectively, followed by flow cytometric analysis (Figure 4). Cells exhibiting elevated MMP showed increased accumulation of JC-1 aggregates (red fluorescence) and a higher red/green fluorescence ratio. In contrast, mitochondria with lower MMP displayed higher green fluorescence from monomeric JC-1 and a lower red/green ratio. Cells with higher intracellular calcium levels demonstrated stronger green fluorescence intensity.
The data showed that pathological hmMSCs had approximately threefold lower mitochondrial activity and a twofold higher steady-state level of iCa2+ compared with healthy hmMSCs (Figure 4). Interestingly, pathological SDCs exhibited a significant (~threefold) increase in mitochondrial activity and a reduction in iCa2+ levels, approaching those observed in healthy hmMSCs. These findings suggest that SDCs derived from DCM myocardium retain high energetic capacity and exhibit flexible iCa2+ signaling, potentially enhanced by 3D cultivation conditions. In contrast, healthy hmMSCs already displayed high mitochondrial activity, which increased only slightly in their corresponding SDCs. Similarly, the iCa2+ level in healthy hmMSCs cells was already balanced and did not require significant adjustment in SDCs.

2.4. Oxidative Phosphorylation and Glycolysis of Healthy and Pathological hmMSCs and SDCs

The promising impact of cardiospheres on mitochondrial activity prompted a more detailed analysis of oxidative phosphorylation and glycolytic parameters (Figure 5). Therefore, a comprehensive examination of mitochondrial function in both healthy and pathological hmMSCs and SDCs was conducted using the Seahorse XF Analyzer (Agilent Technologies, North Billerica, MA, USA). This system enables the assessment of mitochondrial respiration and glycolytic activity by measuring the oxygen consumption rate (OCR) and extracellular acidification rate (ECAR), respectively.
The Seahorse data confirmed previous observations: pathological hmMSCs exhibited significantly lower coupling efficiency (9.1 vs. 281.22 pM/min/mg protein) and ATP production (19.2 vs. 98.1 pM/min/mg protein), as well as a higher proton leak (98.14 vs. 19.24 pM/min/mg protein) compared with healthy hmMSCs (Figure 5). Interestingly, maximal respiration (177.68 vs. 119.39 pM/min/mg protein) and spare respiratory capacity (76.79 vs. 31.23 pM/min/mg protein) were higher in pathological hmMSCs than in healthy cells, suggesting that pathological cells retain mitochondrial energetic reserves that can potentially be activated for regenerative purposes.
Cultivation of both healthy and pathological cardiac progenitors in 3D cardiospheres, followed by assessment of their derived SDCs, revealed a significant increase in ATP production in both SDC types (from 68.91 to 115.08 pM/min/mg protein in healthy, and from 3.4 to 64.4 pM/min/mg protein in pathological cells). Coupling efficiency was also markedly upregulated only in pathological SDCs (from 9.1 to 366.97 pM/min/mg protein), while proton leak remained nearly unchanged between the two SDC types. Additionally, 3D cardiosphere cultivation enhanced basal respiration, maximal respiration, and spare respiratory capacity in both healthy and pathological SDCs.
These findings suggest that 3D cardiosphere cultivation enhances nearly all mitochondrial bioenergetic parameters, except for the proton leak, in SDCs compared with hmMSCs. Mitochondrial proton leak refers to protons re-entering the mitochondrial matrix independently of ATP synthase, which might be related to the intactness of mitochondrial inner membrane; this phenomenon may result from alterations in mitochondrial inner membrane lipid composition and/or structural disruptions. In such cases, proton leak reflects pathological uncoupling, not physiological regulation. The impact of 3D cardiosphere culture on glycolysis in healthy and pathological SDCs was also investigated (Figure 6a,b). The results confirmed that 2D-cultured pathological hmMSCs exhibited lower glycolytic activity compared with 2D-cultured healthy hmMSCs (Figure 6). Furthermore, 3D cardiosphere did not significantly improve glycolytic stress parameters in either healthy or pathological SDCs compared with their corresponding hmMSCs. These findings indicate that the reduced glycolytic activity observed in pathological hmMSCs and SDCs is associated with their slower proliferation rate compared with healthy cells (Figure 2). The improved mitochondrial activity accompanied by decreased glycolytic activity suggests a metabolic shift of SDCs from proliferation toward differentiation and regeneration.

2.5. Cardiac Commitment Gene Expression Profile of Healthy and Pathological Cardiospheres and SDCs Versus hmMSCs

The observed differences in cell morphology, proliferation, mitochondrial membrane potential, and metabolic activity between hmMSCs and SDCs prompted an investigation of the expression of cardiac commitment-related transcription factors (HOPX, Nkx2.5, MEF2C, GATA4), cell membrane (c-KIT, NOTCH1, CD90, TGFBR2), CDH2, GJA1, CACNA1C) and endoplasmic reticulum-associated genes (ITPR2, CALR) in cardiospheres and SDCs vs. hmMSCs (Figure 7).
Gene expression analysis revealed that healthy SDCs exhibited significantly higher expression of stemness-related genes such as c-KIT, NOTCH1, and THY1, whereas pathological SDCs showed increased expression of c-KIT and THY1 only (Figure 7). These findings suggest that SDCs from both healthy and pathological myocardiums retain and enhance their mesenchymal/stem-like phenotype, characteristic of progenitor cells. The 3D cardiosphere culture likely promotes a more primitive and regenerative phenotype, maintaining high expression of THY1 along with other stemness-associated markers (c-KIT, HOPX, and NOTCH1) (Figure 7).
Furthermore, the expression of cardiac commitment markers such as MEF2C and GATA4 was significantly upregulated in healthy SDCs, whereas NKX2-5 was the only commitment factor significantly increased in dilated myocardium-derived SDCs (Figure 7). In addition, the expression of intracellular calcium regulators such as CACNA1C (Cav1.2) and CALR was also higher in healthy SDCs than in pathological counterparts (Figure 7).
It is noteworthy that gene expression upregulation occurred earlier (at passage 1) in healthy SDCs, while in pathological SDCs, similar changes appeared later (at passage 2). These findings indicate that healthy myocardium-derived SDCs likely possess a more stable differentiation and regeneration program, supported by superior metabolic and mitochondrial functions. Nevertheless, SDCs derived from dilated myocardium still retain residual regenerative potential, which could potentially be enhanced through targeted modulation.

3. Discussion

In recent decades, several clinical trials have investigated cell therapy using intracoronary infusion of autologous SDCs in post-MI patients [14], allogeneic SDCs in patients with progressive heart failure [21] or intracoronary administration of allogeneic SDCs in patients with post-MI left ventricular dysfunction [22]. Although SDC treatment was safe, it did not significantly reduce scar size compared with placebo at 6 months. However, reductions in left ventricular volumes and NT-proBNP levels indicated some disease-modifying bioactivity of SDCs. Despite the apparent therapeutic benefits of SDCs in cardiac regeneration, several questions remain regarding their biology—particularly when comparing cells derived from healthy versus dilated myocardium. One key question is whether it is possible to enhance the energetic and cardiac commitment status of autologous human dilated myocardium-derived SDCs compared with hmMSCs, and if so, which parameters change the most and how.
Human myocardium-derived stem/stromal cells (hmMSCs) can be isolated from biopsies of both healthy and dilated ventricular myocardium. In our previous studies, we demonstrated that hmMSCs express typical mesenchymal stem cell surface markers and are capable of differentiating into osteogenic, adipogenic, chondrogenic, and cardiomyogenic lineages [19,20]. Additionally, we investigated the epigenetic acetylation levels of healthy and dilated hmMSCs and showed that histone deacetylase inhibitors can enhance their regenerative potential [19,20]. In the present study, we compared healthy and dilated myocardium-derived hmMSCs, cardiospheres, and cardiosphere-derived cells (SDCs) to assess their morphological characteristics, proliferation, energetic/metabolic profiles, and cardiac commitment-related gene reprogramming.
Other studies have shown that three-dimensional (3D) cell cultures provide a more physiologically relevant model than traditional two-dimensional (2D) systems, enabling natural cell-environment interactions and more accurate predictions of biological behavior and drug responses [23,24,25]. It has also been reported that the transition from 2D to 3D culture is accompanied by metabolic reprogramming, characterized by increased mitochondrial respiration and glycolytic activity [26]. This study demonstrates that in pathological SDCs, the decreased mitochondrial membrane potential (MMP) and elevated intracellular Ca2+ levels in pathological hmMSCs were restored nearly to those of healthy hmMSCs. SDCs also exhibited higher oxidative phosphorylation and lower glycolytic activity compared with hmMSCs of both types. Detailed metabolic analysis revealed that while pathological hmMSCs showed reduced mitochondrial coupling and ATP production—both improved in SDCs—the increased proton leak observed in pathological hmMSCs persisted in SDCs. Interestingly, pathological hmMSCs exhibited higher spare respiratory capacity and maximal respiration than healthy hmMSCs, suggesting a retained energetic potential in DCM-derived hmMSCs. The enhancement of oxidative phosphorylation (OXPHOS) as an alternative energy production pathway to glycolysis in 3D cultures has been proposed to reflect increased energy demands and necessary metabolic adaptation to the 3D microenvironment [27,28].
Morphological and proliferation analyses revealed new tendencies related to the glycolytic activity of hmMSCs and SDCs. In this study, both healthy and pathological SDCs were smaller in size and proliferated faster than hmMSCs, while exhibiting significantly lower glycolytic activity. According to the generally accepted theory, adult stem cells display metabolic flexibility regulated by their adhesion status—adherent cells primarily utilize glycolysis, whereas suspended cells depend on oxidative phosphorylation (OXPHOS) to meet their ATP demands [29]. Our data suggest that SDCs rely more heavily on mitochondrial OXPHOS than on glycolysis to support their energy requirements. Enhanced mitochondrial function, biogenesis, or metabolic efficiency in SDCs may enable greater ATP production per glucose molecule, thereby sustaining higher proliferation despite reduced glycolytic flux. It is possible that the metabolic transition from OXPHOS to glycolysis in SDCs requires longer incubation under 2D culture conditions. Alternatively, the observed lower glycolytic rate may reflect a more mature and energetically balanced phenotype, consistent with the partial cardiac commitment of SDCs compared with undifferentiated hmMSCs [30].
Metabolic changes in SDCs are often accompanied by modulation of transcription factors and other genes related to cardiac progenitor identity and lineage commitment. In this study, the most pronounced expression of stemness-related genes—c-KIT, NOTCH1, and THY1 (CD90)—was observed in healthy SDCs at passage one, whereas in pathological SDCs, increased expression of c-KIT and THY1 was detected only at passage two. These data indicate that healthy SDCs more readily shift toward a primitive, progenitor-like state than pathological SDCs. Activation of the c-KIT receptor has been shown to promote cell survival and proliferation in both human and murine stem cells and cardiomyocytes [31]. Moreover, c-KIT+ cardiac progenitors are recognized as primitive stem cells with multilineage differentiation potential [32]. Clinically, the Phase I SCIPIO trial demonstrated the safety of autologous c-KIT+ cardiac cell administration in patients with ischemic heart failure [33]. However, other studies reported that endogenous c-KIT+ cells generate cardiomyocytes at a functionally insignificant level [34] and that transplanted c-KIT+ cells in infarcted hearts do not significantly differentiate into mature cardiomyocytes [33]. These findings suggest that activation of c-KIT alone may be insufficient to induce effective cardiac regeneration and likely requires the cooperative activation of other stemness-related genes such as NOTCH1, CD90, or additional regulatory factors.
For many years, Thy-1/CD90 was used primarily as a marker to define mesenchymal stem cells (MSCs). However, recent studies have shown that Thy-1 also possesses signaling functions and plays a critical role in determining MSC fate by promoting osteogenic differentiation, while inhibiting adipogenic differentiation [35]. It has also been reported that THY1 expression in fibrotic cardiac tissue is not associated with fibroblast differentiation, but may serve as a marker of fibroblast proliferation [36]. In addition, activation of the Notch signaling pathway has been observed in CD90+ liver cancer cells, where it promotes cell invasion, migration, and the expression of stemness-related genes [37,38]. However, the potential cooperative role of the Notch pathway with other stemness-associated genes such as c-KIT and CD90 in cardiac regeneration remains to be elucidated.
Finally, the expression of cardiomyogenic commitment genes MEF2C and GATA4 was significantly upregulated in healthy SDCs, whereas in pathological SDCs, only the early cardiomyogenic differentiation gene NKX2.5 was elevated. The transcription factors MEF2C and GATA4 are key regulators of cardiomyocyte maturation and structural gene expression. A recent study demonstrated that transfection of GATA4 and MEF2C, individually or in combination, induced cardiac differentiation in human umbilical cord-derived MSCs, leading to increased expression of cardiac genes such as NKX2.5, myosin heavy chain (MYH6, MYH7), connexin-43 (GJA1), and cardiac proteins including GATA4, NKX2.5, cardiac troponin T, and connexin-43 [39]. In this study, the higher expression of MEF2C and GATA4 in healthy SDCs suggests that these cells are further along the cardiomyogenic differentiation pathway, reflecting a more mature or committed cardiac phenotype. In contrast, NKX2.5—an early cardiac progenitor marker and one of the first transcription factors expressed during cardiac lineage specification [40]—was significantly upregulated in pathological SDCs, indicating that these cells may be arrested at an earlier developmental stage compared with healthy SDCs, maintaining a more primitive or progenitor-like phenotype rather than progressing toward full cardiomyogenic differentiation. DCM-derived SDCs may therefore require stronger stimuli than 3D cultivation alone to enhance cardiac commitment. The chronic stress, fibrosis, and/or metabolic impairments in dilated myocardium may further delay or inhibit normal differentiation, maintaining cells in a compensatory, progenitor-activated state. This area warrants further detailed investigation.

4. Materials and Methods

4.1. Isolation of hmMSC and Growing in 2D Conditions

Human healthy ventricular biopsies were obtained from 65–75-year-old men patients without mitral valve disease, non-dilated left ventricle, and preserved its function. Human dilated myocardium biopsies were obtained from 65–75-year-old men with mitral valve disorders and dilated left ventricles showing reduced ejection fraction (<45–50%). Healthy biopsies were obtained from inside the heart using a thin catheter; meanwhile, dilated myocardium biopsies were obtained from leftover material after mitral valve change operation. Cell isolation was performed according to previously described protocols with minor modifications [20]. Briefly, myocardial specimens were minced into fragments smaller than 1 mm3, thoroughly washed with phosphate-buffered saline (PBS) containing 2% antibiotics, and partially digested with trypsin. The tissue fragments were then cultured as explants on fibronectin-coated (2 mg/mL) six-well plates in Iscove’s Modified Dulbecco’s Medium (IMDM) supplemented with 20% fetal bovine serum (FBS), 100 U/mL penicillin G, and 100 U/mL streptomycin (Thermo Fisher Scientific, Waltham, MA, USA). Within approximately 1–3 weeks, a layer of stromal-like cells emerged from the explants. Once near confluence, cardiac outgrowth cells were detached using trypsin and transferred to 75 cm2 flasks coated with 0.2% gelatin for further expansion. The remaining tissue fragments were re-plated onto fibronectin-coated dishes to allow continued outgrowth formation. Cardiac outgrowth cells could be harvested up to four times from the same explants. Cell counting for the experiments was performed with Fast-Read 102® plastic chamber (Biosigma, Cona, Italy).

4.2. Formations of Cardiospheres

During explant outgrowth phase, small round loosely attached cells were harvested with 0.05% of trypsin and seeded on the poly-HEMA poly(2-hydroxyethyl methacrylate)-coated 96 well plates for the formation of cardiospheres. Collected cells were grown in special cardiosphere medium for 3 days 35% IMDM/65% DMEM-Ham’s F-12 (GIBCO), 3.5% Embryonic stem cell FBS, 2% B27, 100 U/mL penicillin G, 100 U/mL streptomycin (Thermo Fisher Scientific, Waltham, MA, USA), 0.1 mmol/L 2-mercaptoethanol, 10 ng/mL EGF and 20 ng/mL bFGF (Sigma Aldrich, St. Louis, MO, USA), 10 ng/mL Cardiotrophin-1 (Biochrom, Cambridge, UK), 1 unit/mL thrombin (Merck KGaA, Damstadt, Germany), to form 3D cardiospheres [29]. After this, cardiospheres were dissociated with 0.25% of trypsin-EDTA to obtain cardiosphere-derived cells (SDCs) that were seeded on the 25 cm2 flask coated with the 0.2% gelatine for the further growth in IMDM supplemented with 10% FBS and 100 U/mL penicillin G, 100 U/mL streptomycin. In this study 1st to 3rd passage of SDCs were used for the further experiments.

4.3. Proliferation Measurements

A total of 5000 cells were calculated with Fast-Read® 102 (Biosigma, Cona, Italy) and seeded in 24-well plate. The proliferation of ventricle-derived hmMSCs and SDCs was monitored continuously over a period of six days using the CCK-8 kit, as recommended by the manufacturer (Dojindo, Kumamoto, Japan). Briefly, 12.5 µL of CCK-8 solution was introduced to 500 µL of fresh growth medium, followed by a 3 h incubation at 37 °C followed by absorbance measurement at 450 nm using the SpectraMax i3 spectrophotometer (Molecular Devices, San Jose, CA, USA).

4.4. Evaluation of Intracellular Calcium Measurement Using Flow Cytometry

For flow cytometry analysis, cells were seeded into six-well plates at a density of 50,000 cells per well. After reaching the desired confluence, cells were detached using trypsin, transferred to flow cytometry tubes, and centrifuged at 600× g for 5 min. The cell pellets were stained with the 3 μM of calcium-specific fluorescent dye Cal-520 (Santa Cruz Biotechnology, Dallas, TX, USA) in IMDM with 1% PEST and incubated at 37 °C with 5% CO2 for 30 min. Following incubation, cells were washed twice with 1% BSA PBS solution and centrifuged at 600× g for 5 min. The supernatant was discarded, and cells were resuspended in 300 μL of PBS containing 1% BSA. Each sample was analyzed in triplicate, and fluorescence intensity was measured using a FACSAria III flow cytometer (BD Biosciences, San Jose, CA, USA).

4.5. Evaluation of Mitochondria Potential Using Flow Cytometry

The confluent cells monolayer was harvested with trypsin; cells were stained with 2 µM of JC1 in IMDM with 1% PEST at 37 °C with 5% CO2 for 30 min. After staining, cells were washed twice and resuspended in 300 μL of 1% BSA PBS solution. Flow cytometry was performed with BD FACSAria II instrument (BD Biosciences, San Jose, CA, USA); data were analyzed with BD FACSDiva software 6.1 (BD Biosciences, San Jose, CA, USA).

4.6. Metabolic Measurements with Seahorse

Seahorse XFp Analyzer (Agilent Technologies, Santa Clara, CA, USA) was used to evaluate the oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) of hmMSCs and SDCs. The cells were seeded at a density of 20,000 cells per well on Seahorse XFp culture plates a day before the measurements. Seahorse medium (Agilent Technologies, Santa Clara, CA, USA) was prepared following the manufacturer’s recommendations. For OCR measurements, the medium contained final concentrations of 10 mM glucose, 2 mM glutamax, and 1 mM sodium pyruvate. Mitochondrial parameters were assessed by sequentially introducing 10 µM oligomycin, 20 µM carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP), and 5 µM of rotenone (Agilent Technologies, Santa Clara, CA, USA).
In ECAR measurements, 1 mM of glutamine was added to the Seahorse medium. Glycolytic stress was evaluated by sequentially adding 100 mM glucose, 50 µM oligomycin, and 500 mM 2-deoxyglucose (DG) (Agilent Technologies, Santa Clara, CA, USA). First passage hmMSCs and SDCs were used for this experiment. After the experiment, all samples were lysed in lysis buffer (50 mM Tris HCl, pH = 6.8, 10% glycerol, 1% SDS), followed by centrifugation at 20,000× g and + 4 °C for 15 min. The protein concentration was determined using the Pierce™ Modified Lowry Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA). Energetic parameters were normalized by protein concentration and expressed as pM/min/mg/mL protein for OCR measurements and mpH/min/mg/mL protein for ECAR measurements.

4.7. Real-Time RT-PCR

RNA was purified approximately from 2 × 105 hmMSCs, SDCs or cardiospheres using GeneJetTM RNA purification kit (Thermo Fisher Scientific, Waltham, MA, USA). A total of 500 ng of RNA was incubated with DnaseI at 37 °C for 30 min, then reaction was stopped by adding 50 mM EDTA and heating at 65 °C for 5 min. (Thermo Fisher Scientific, Waltham, MA, USA). Reverse transcription was performed with a High Capacity cDNA reverse transcription kit (Thermo Fisher Scientific, Waltham, MA, USA). Real-time PCR was performed in triplicate using the 2× Maxima Probe qPCR Master Mix (Thermo Fisher Scientific, Waltham, MA, USA) on a AriaMx Real-Time PCR Machine (Agilent Technologies, Santa Clara, CA, USA) with an annealing temperature of 60 °C.
Expression of transcription factors (HOPX, GATA4, NKX2-5, MEF2C), cell membrane (cKIT, NOTCH1, THY1, TGFBRII, CDH2, GJA1, CACNA1C), and endoplasmic reticulum-related (ITPR2 and CALR) genes was determined (Thermo Fisher Scientific, Waltham, MA, USA) (Table 1). HPR1, ACTB, GAPDH, and B2M housekeeping genes were evaluated for expression stability across different cell types, patients, and culturing conditions using equal amounts of RNA for cDNA synthesis. Among these genes, B2M showed lowest coefficient of variation and NormFinder stability value; therefore, it was selected for gene expression normalization.

4.8. Statistical Analysis

The statistical analysis was performed using the Excel software (Microsoft Corporation, Redmont, WA, USA) and Graphpad Prism 6.01 (GraphPad Software, San Diego, CA, USA). The data are expressed as means ± standard deviation (mean ± SD) based on not less than three replicates from two to three healthy and dilated human myocardium-derived hmMSCs and SDCs. Significance of the data was determined using Student’s t-test, and significance levels were indicated at * p ≤ 0.05 or ** p ≤ 0.01 levels.

5. Conclusions

Our findings demonstrate that SDCs derived from both healthy and dilated atrial myocardium possess a more favorable metabolic, progenitor and cardiac commitment profile than conventional hmMSCs.
Although some mitochondrial features—such as proton leak—remained suboptimal, the overall enhancement in ATP production, coupling efficiency, and respiratory capacity underscores the metabolic advantages of both types of SDCs over conventional hmMSCs. The higher metabolic activity of cardiac SDCs over hmMSCs may also be related to the stronger expression of some stemness- and cardiomyogenesis-associated genes.
Collectively, while DCM-derived hmMSCs exhibited signs of impaired mitochondrial function, lower proliferation and altered calcium homeostasis, the SDC phenotype partially restored several key bioenergetic parameters, highlighting the residual metabolic potential of pathological cells, which can be targeted stimulated. Healthy SDCs possess a more mature or committed cardiac phenotype, whereas dilated myocardium-derived SDCs may require supplementary external stimuli, in conjunction with or beyond cardiosphere culture, to further achieve enhanced cardiac commitment.
A limitation of this study relates to the intrinsic differences in cellular homogeneity between SDCs and hmMSCs and the potential impact this could have on data interpretation. hmMSCs are known to represent a heterogeneous population with some variability in cell surface marker expression, differentiation state, and/or metabolic activity [54], whereas SDCs appear more phenotypically uniform under the conditions used here. The widely used definition of typical MSC surface markers may not be universally applicable, as markers expression is highly dependent on the tissue of origin [55,56].
This disparity in population homogeneity may partly account for the observed differences in both metabolomic and transcriptomic profiles, as bulk analyses inevitably reflect averaged signals that can be disproportionately influenced by dominant subpopulations. However, SDCs displayed highly proliferative, secretory and immunomodulatory properties that can also be found in activated or inflammatory cell types [57]. Moreover, sc-RNAseq data from human right atrial biopsies in comparison with SDCs uncovered transcriptomic similarities between SDCs and cardiac fibroblasts, also named as hmMSCs, but not with cardiac progenitor cells derived from human-induced pluripotent stem cells [57]. While our study provides valuable comparative insights, future work incorporating single-cell-level analyses in more refined phenotypic and functional stratification will be necessary to disentangle heterogeneity-driven cardiac regenerative effects from genuine biological differences between SDCs and hmMSCs.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/ijms27031303/s1.

Author Contributions

Conceptualization, D.B.; methodology, R.M.; software, R.M.; validation, D.B., R.M.; formal analysis, D.B.; investigation, R.M.; resources, D.B.; data curation, R.M.; writing—original draft preparation, D.B.; writing—review and editing, R.M.; visualization, R.M.; supervision, D.B.; project administration, D.B.; funding acquisition, D.B. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Lithuanian Research Council, grant number S-MIP-17-13 and S-PD-24-41.

Institutional Review Board Statement

The study was conducted in accordance with the Declaration of Helsinki, and approved by the Vilnius Regional Biomedical Research Ethics Committee (License No. 158200-14-741-257, Approval Date 10 June 2014).

Informed Consent Statement

Informed consent was obtained from all patients to obtain biopsy for this study.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

Acknowledgments

We acknowledge the cardiac surgeons K. Ručinskas and V. Janušauskas for patients’ selection and obtaining biosamples. The authors take full responsibility for the content of this publication.

Conflicts of Interest

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

Abbreviations

The following abbreviations are used in this manuscript:
SDCsSphere-derived cells
hmMSCsHuman cardiac myocardium-derived mesenchymal stem/stromal cells
DCMDilated cardiomyopathy
ATPAdenosine triphosphate
HFHeart failure
OCROxygen consumption rate
ECARExtracellular acidification rate
MMPMitochondrial membrane potential
KITKIT Proto-Oncogene, Receptor Tyrosine Kinase
NKX2-5NK2 Homeobox 5
MDR1 ATP Binding Cassette Subfamily B Member 1
HOPXHOP Homeobox
NOTCH1Notch Receptor 1
TGFBR2Transforming Growth Factor Beta Receptor 2
CDH2Cadherin 2
GJA1Gap Junction Protein Alpha 1
CACNA1CCalcium Voltage-Gated Channel Subunit Alpha1 C
ITPR2Inositol 1,4,5-Trisphosphate Receptor Type 2
CALRCalreticulin

References

  1. Lazenby, R.A.; White, R.J. Advances and Perspectives in Chemical Imaging in Cellular Environments Using Electrochemical Methods. Chemosensors 2018, 6, 24. [Google Scholar] [CrossRef]
  2. Lakdawala, N.K.; Winterfield, J.R.; Funke, B.H. Dilated Cardiomyopathy. Circ. Arrhythmia Electrophysiol. 2013, 6, 228–237. [Google Scholar] [CrossRef]
  3. Lushaj, E.B.; Anstadt, E.; Haworth, R.; Roenneburg, D.; Kim, J.; Hematti, P.; Kohmoto, T. Mesenchymal Stromal Cells Are Present in the Heart and Promote Growth of Adult Stem Cells in Vitro. Cytotherapy 2011, 13, 400–406. [Google Scholar] [CrossRef]
  4. Beltrami, A.P.; Barlucchi, L.; Torella, D.; Baker, M.; Limana, F.; Chimenti, S.; Kasahara, H.; Rota, M.; Musso, E.; Urbanek, K.; et al. Adult Cardiac Stem Cells Are Multipotent and Support Myocardial Regeneration. Cell 2003, 114, 763–776. [Google Scholar] [CrossRef] [PubMed]
  5. Matsuura, K.; Nagai, T.; Nishigaki, N.; Oyama, T.; Nishi, J.; Wada, H.; Sano, M.; Toko, H.; Akazawa, H.; Sato, T.; et al. Adult Cardiac Sca-1-Positive Cells Differentiate into Beating Cardiomyocytes. J. Biol. Chem. 2004, 279, 11384–11391. [Google Scholar] [CrossRef] [PubMed]
  6. Yoon, J.; Shim, W.J.; Ro, Y.M.; Lim, D.-S. Transdifferentiation of Mesenchymal Stem Cells into Cardiomyocytes by Direct Cell-to-Cell Contact with Neonatal Cardiomyocyte but Not Adult Cardiomyocytes. Ann. Hematol. 2005, 84, 715–721. [Google Scholar] [CrossRef]
  7. Messina, E.; De Angelis, L.; Frati, G.; Morrone, S.; Chimenti, S.; Fiordaliso, F.; Salio, M.; Battaglia, M.; Latronico, M.V.G.; Coletta, M.; et al. Isolation and Expansion of Adult Cardiac Stem Cells From Human and Murine Heart. Circ. Res. 2004, 95, 911–921. [Google Scholar] [CrossRef]
  8. Emani, S.M.; Del Nido, P.J. Cell-Based Therapy With Cardiosphere-Derived Cardiocytes: A New Hope for Pediatric Patients With Single Ventricle Congenital Heart Disease? Circ. Res. 2018, 122, 916–917. [Google Scholar] [CrossRef]
  9. Davis, D.R.; Zhang, Y.; Smith, R.R.; Cheng, K.; Terrovitis, J.; Malliaras, K.; Li, T.-S.; White, A.; Makkar, R.; Marbán, E. Validation of the Cardiosphere Method to Culture Cardiac Progenitor Cells from Myocardial Tissue. PLoS ONE 2009, 4, e7195. [Google Scholar] [CrossRef]
  10. Boukouaci, W.; Lauden, L.; Siewiera, J.; Dam, N.; Hocine, H.-R.; Khaznadar, Z.; Tamouza, R.; Borlado, L.R.; Charron, D.; Jabrane-Ferrat, N.; et al. Natural Killer Cell Crosstalk with Allogeneic Human Cardiac-Derived Stem/Progenitor Cells Controls Persistence. Cardiovasc. Res. 2014, 104, 290–302. [Google Scholar] [CrossRef] [PubMed]
  11. De Couto, G.; Liu, W.; Tseliou, E.; Sun, B.; Makkar, N.; Kanazawa, H.; Arditi, M.; Marbán, E. Macrophages Mediate Cardioprotective Cellular Postconditioning in Acute Myocardial Infarction. J. Clin. Investig. 2015, 125, 3147–3162. [Google Scholar] [CrossRef]
  12. Kanazawa, H.; Tseliou, E.; Malliaras, K.; Yee, K.; Dawkins, J.F.; De Couto, G.; Smith, R.R.; Kreke, M.; Seinfeld, J.; Middleton, R.C.; et al. Cellular Postconditioning: Allogeneic Cardiosphere-Derived Cells Reduce Infarct Size and Attenuate Microvascular Obstruction When Administered After Reperfusion in Pigs With Acute Myocardial Infarction. Circ. Heart Fail. 2015, 8, 322–332. [Google Scholar] [CrossRef]
  13. Lauden, L.; Boukouaci, W.; Borlado, L.R.; López, I.P.; Sepúlveda, P.; Tamouza, R.; Charron, D.; Al-Daccak, R. Allogenicity of Human Cardiac Stem/Progenitor Cells Orchestrated by Programmed Death Ligand 1. Circ. Res. 2013, 112, 451–464. [Google Scholar] [CrossRef]
  14. Malliaras, K.; Li, T.-S.; Luthringer, D.; Terrovitis, J.; Cheng, K.; Chakravarty, T.; Galang, G.; Zhang, Y.; Schoenhoff, F.; Van Eyk, J.; et al. Safety and Efficacy of Allogeneic Cell Therapy in Infarcted Rats Transplanted With Mismatched Cardiosphere-Derived Cells. Circulation 2012, 125, 100–112. [Google Scholar] [CrossRef] [PubMed]
  15. Dimmeler, S.; Leri, A. Aging and Disease as Modifiers of Efficacy of Cell Therapy. Circ. Res. 2008, 102, 1319–1330. [Google Scholar] [CrossRef]
  16. Flam, E.; Jang, C.; Murashige, D.; Yang, Y.; Morley, M.P.; Jung, S.; Kantner, D.S.; Pepper, H.; Bedi, K.C.; Brandimarto, J.; et al. Integrated Landscape of Cardiac Metabolism in End-Stage Human Nonischemic Dilated Cardiomyopathy. Nat. Cardiovasc. Res. 2022, 9, 817–829. [Google Scholar] [CrossRef] [PubMed]
  17. Lopaschuk, G.D.; Karwi, Q.G.; Tian, R.; Wende, A.R.; Abel, E.D. Cardiac Energy Metabolism in Heart Failure. Circ. Res. 2021, 128, 1487–1513. [Google Scholar] [CrossRef] [PubMed]
  18. Shyh-Chang, N.; Ng, H.H. The metabolic programming of stem cells. Genes. Dev. 2017, 31, 336–346. [Google Scholar] [CrossRef]
  19. Miksiunas, R.; Aldonyte, R.; Vailionyte, A.; Jelinskas, T.; Eimont, R.; Stankeviciene, G.; Cepla, V.; Valiokas, R.; Rucinskas, K.; Janusauskas, V.; et al. Cardiomyogenic Differentiation Potential of Human Dilated Myocardium-Derived Mesenchymal Stem/Stromal Cells: The Impact of HDAC Inhibitor SAHA and Biomimetic Matrices. Int. J. Mol. Sci. 2021, 22, 12702. [Google Scholar] [CrossRef]
  20. Miksiunas, R.; Rucinskas, K.; Janusauskas, V.; Labeit, S.; Bironaite, D. Histone Deacetylase Inhibitor Suberoylanilide Hydroxamic Acid Improves Energetic Status and Cardiomyogenic Differentiation of Human Dilated Myocardium-Derived Primary Mesenchymal Cells. Int. J. Mol. Sci. 2020, 21, 4845. [Google Scholar] [CrossRef]
  21. Chakravarty, T.; Henry, T.D.; Kittleson, M.; Lima, J.; Siegel, R.J.; Slipczuk, L.; Pogoda, J.M.; Smith, R.R.; Malliaras, K.; Marbán, L.; et al. Allogeneic Cardiosphere-Derived Cells for the Treatment of Heart Failure with Reduced Ejection Fraction: The Dilated cardiomYopathy iNtervention with Allogeneic MyocardIally-Regenerative Cells (DYNAMIC) Trial. EuroIntervention 2020, 16, e293–e300. [Google Scholar] [CrossRef]
  22. Makkar, R.R.; Kereiakes, D.J.; Aguirre, F.; Kowalchuk, G.; Chakravarty, T.; Malliaras, K.; Francis, G.S.; Povsic, T.J.; Schatz, R.; Traverse, J.H.; et al. Intracoronary ALLogeneic Heart STem Cells to Achieve Myocardial Regeneration (ALLSTAR): A Randomized, Placebo-Controlled, Double-Blinded Trial. Eur. Heart J. 2020, 41, 3451–3458. [Google Scholar] [CrossRef]
  23. Antoni, D.; Burckel, H.; Josset, E.; Noel, G. Three-Dimensional Cell Culture: A Breakthrough in Vivo. Int. J. Mol. Sci. 2015, 16, 5517–5527. [Google Scholar] [CrossRef]
  24. Fang, Y.; Eglen, R.M. Three-Dimensional Cell Cultures in Drug Discovery and Development. SLAS Discov. 2017, 22, 456–472, Erratum in SLAS Discov. 202, 2, 1. https://doi.org/10.1177/2472555218787198. [Google Scholar] [CrossRef] [PubMed]
  25. Wanigasekara, J.; Carroll, L.J.; Cullen, P.J.; Tiwari, B.; Curtin, J.F. Three-Dimensional (3D) in Vitro Cell Culture Protocols to Enhance Glioblastoma Research. PLoS ONE 2023, 18, e0276248. [Google Scholar] [CrossRef]
  26. Rybkowska, P.; Radoszkiewicz, K.; Kawalec, M.; Dymkowska, D.; Zabłocka, B.; Zabłocki, K.; Sarnowska, A. The Metabolic Changes between Monolayer (2D) and Three-Dimensional (3D) Culture Conditions in Human Mesenchymal Stem/Stromal Cells Derived from Adipose Tissue. Cells 2023, 12, 178. [Google Scholar] [CrossRef]
  27. Du Plessis, S.; Agarwal, A.; Mohanty, G.; Van Der Linde, M. Oxidative Phosphorylation versus Glycolysis: What Fuel Do Spermatozoa Use? Asian J. Androl. 2015, 17, 230. [Google Scholar] [CrossRef]
  28. Nam, A.; Jain, S.; Wu, C.; Campos, A.; Shepard, R.M.; Yu, Z.; Reddy, J.P.; Von Schalscha, T.; Weis, S.M.; Onaitis, M.; et al. Integrin Avβ3 Upregulation in Response to Nutrient Stress Promotes Lung Cancer Cell Metabolic Plasticity. Cancer Res. 2024, 84, 1630–1642. [Google Scholar] [CrossRef] [PubMed]
  29. Afzal, J.; Chan, A.; Karakas, M.F.; Woldemichael, K.; Vakrou, S.; Guan, Y.; Rathmell, J.; Wahl, R.; Pomper, M.; Foster, D.B.; et al. Cardiosphere-Derived Cells Demonstrate Metabolic Flexibility That Is Influenced by Adhesion Status. JACC Basic. Transl. Sci. 2017, 2, 543–560. [Google Scholar] [CrossRef] [PubMed]
  30. Zumbaugh, M.D.; Johnson, S.E.; Shi, T.H.; Gerrard, D.E. Molecular and Biochemical Regulation of Skeletal Muscle Metabolism. J. Anim. Sci. 2022, 100, skac035. [Google Scholar] [CrossRef]
  31. Gude, N.A.; Firouzi, F.; Broughton, K.M.; Ilves, K.; Nguyen, K.P.; Payne, C.R.; Sacchi, V.; Monsanto, M.M.; Casillas, A.R.; Khalafalla, F.G.; et al. Cardiac C-Kit Biology Revealed by Inducible Transgenesis. Circ. Res. 2018, 123, 57–72. [Google Scholar] [CrossRef]
  32. Gambini, E.; Pompilio, G.; Biondi, A.; Alamanni, F.; Capogrossi, M.C.; Agrifoglio, M.; Pesce, M. C-Kit+ Cardiac Progenitors Exhibit Mesenchymal Markers and Preferential Cardiovascular Commitment. Cardiovasc. Res. 2011, 89, 362–373. [Google Scholar] [CrossRef]
  33. Sanganalmath, S.K.; Bolli, R. Cell Therapy for Heart Failure: A Comprehensive Overview of Experimental and Clinical Studies, Current Challenges, and Future Directions. Circ. Res. 2013, 113, 810–834. [Google Scholar] [CrossRef] [PubMed]
  34. Van Berlo, J.H.; Kanisicak, O.; Maillet, M.; Vagnozzi, R.J.; Karch, J.; Lin, S.-C.J.; Middleton, R.C.; Marbán, E.; Molkentin, J.D. C-Kit+ Cells Minimally Contribute Cardiomyocytes to the Heart. Nature 2014, 509, 337–341. [Google Scholar] [CrossRef] [PubMed]
  35. Saalbach, A.; Anderegg, U. Thy-1: More than a Marker for Mesenchymal Stromal Cells. FASEB J. 2019, 33, 6689–6696. [Google Scholar] [CrossRef] [PubMed]
  36. Hudon-David, F.; Bouzeghrane, F.; Couture, P.; Thibault, G. Thy-1 Expression by Cardiac Fibroblasts: Lack of Association with Myofibroblast Contractile Markers. J. Mol. Cell. Cardiol. 2007, 42, 991–1000. [Google Scholar] [CrossRef]
  37. Luo, J.; Wang, P.; Wang, R.; Wang, J.; Liu, M.; Xiong, S.; Li, Y.; Cheng, B. The Notch Pathway Promotes the Cancer Stem Cell Characteristics of CD90+ Cells in Hepatocellular Carcinoma. Oncotarget 2016, 7, 9525–9537. [Google Scholar] [CrossRef]
  38. Yang, Z.F.; Ho, D.W.; Ng, M.N.; Lau, C.K.; Yu, W.C.; Ngai, P.; Chu, P.W.K.; Lam, C.T.; Poon, R.T.P.; Fan, S.T. Significance of CD90+ Cancer Stem Cells in Human Liver Cancer. Cancer Cell 2008, 13, 153–166. [Google Scholar] [CrossRef]
  39. Razzaq, S.S.; Khan, I.; Naeem, N.; Salim, A.; Begum, S.; Haneef, K. Overexpression of GATA Binding Protein 4 and Myocyte Enhancer Factor 2C Induces Differentiation of Mesenchymal Stem Cells into Cardiac-like Cells. World J. Stem Cells 2022, 14, 700–713. [Google Scholar] [CrossRef]
  40. Targoff, K.L.; Schell, T.; Yelon, D. Nkx Genes Regulate Heart Tube Extension and Exert Differential Effects on Ventricular and Atrial Cell Number. Dev. Biol. 2008, 322, 314–321. [Google Scholar] [CrossRef]
  41. Friedman, C.E.; Cheetham, S.W.; Negi, S.; Mills, R.J.; Ogawa, M.; Redd, M.A.; Chiu, H.S.; Shen, S.; Sun, Y.; Mizikovsky, D.; et al. HOPX-Associated Molecular Programs Control Cardiomyocyte Cell States Underpinning Cardiac Structure and Function. Dev. Cell 2024, 59, 91–107.e6. [Google Scholar] [CrossRef]
  42. Tong, Y.-F. Mutations of NKX2.5 and GATA4 Genes in the Development of Congenital Heart Disease. Gene 2016, 588, 86–94. [Google Scholar] [CrossRef]
  43. Barnes, R.M.; Harris, I.S.; Jaehnig, E.J.; Sauls, K.; Sinha, T.; Rojas, A.; Schachterle, W.; McCulley, D.J.; Norris, R.A.; Black, B.L. MEF2C Regulates Outflow Tract Alignment and Transcriptional Control of Tdgf1. Development 2016, 143, 774–779. [Google Scholar] [CrossRef]
  44. Heuvelmans, L.; Mostert, D.; Spanò, G.; Stoll, M.; De Windt, L.J. GATA4: Orchestrating Cardiac Development and Beyond. Cardiovasc. Res. 2025, 16, 2476–2483. [Google Scholar] [CrossRef]
  45. Marino, F.; Scalise, M.; Cianflone, E.; Mancuso, T.; Aquila, I.; Agosti, V.; Torella, M.; Paolino, D.; Mollace, V.; Nadal-Ginard, B.; et al. Role of C-Kit in Myocardial Regeneration and Aging. Front. Endocrinol. 2019, 10, 371. [Google Scholar] [CrossRef] [PubMed]
  46. Kachanova, O.; Lobov, A.; Malashicheva, A. The Role of the Notch Signaling Pathway in Recovery of Cardiac Function after Myocardial Infarction. Int. J. Mol. Sci. 2022, 23, 12509. [Google Scholar] [CrossRef] [PubMed]
  47. Gambini, E.; Rurali, E.; Barbagallo, V.; Pirola, S.; Scopece, A.; Biondi, A.; Bassetti, B.; Casaburo, M.; Eramo, L.; Marinelli, G.P.A.; et al. Angiogenic and Reparative Potency of a Human Cardiac CD90−Mesenchymal Subpopulation in Heart Ischemic Model. Transl. Res. 2025, 283, 22–35. [Google Scholar] [CrossRef] [PubMed]
  48. Langlois, D.; Hneino, M.; Bouazza, L.; Parlakian, A.; Sasaki, T.; Bricca, G.; Li, J.Y. Conditional Inactivation of TGF-β Type II Receptor in Smooth Muscle Cells and Epicardium Causes Lethal Aortic and Cardiac Defects. Transgenic Res. 2010, 19, 1069–1082. [Google Scholar] [CrossRef]
  49. Tsai, Y.-W.; Tseng, Y.-S.; Wu, Y.-S.; Song, W.-L.; You, M.-Y.; Hsu, Y.-C.; Chen, W.-P.; Huang, W.-H.; Chng, J.-C.; Lim, C.-L.; et al. N-Cadherin Promotes Cardiac Regeneration by Potentiating pro-Mitotic β-Catenin Signaling in Cardiomyocytes. Nat. Commun. 2025, 16, 896. [Google Scholar] [CrossRef]
  50. Michela, P.; Velia, V.; Aldo, P.; Ada, P. Role of Connexin 43 in Cardiovascular Diseases. Eur. J. Pharmacol. 2015, 768, 71–76. [Google Scholar] [CrossRef]
  51. Harvey, R.D.; Hell, J.W. CaV1.2 Signaling Complexes in the Heart. J. Mol. Cell. Cardiol. 2013, 58, 143–152. [Google Scholar] [CrossRef] [PubMed]
  52. Garcia, M.I.; Boehning, D. Cardiac Inositol 1,4,5-Trisphosphate Receptors. Biochim. Biophys. Acta BBA–Mol. Cell Res. 2017, 1864, 907–914. [Google Scholar] [CrossRef]
  53. Groenendyk, J.; Wang, W.-A.; Robinson, A.; Michalak, M. Calreticulin and the Heart. Cells 2022, 11, 1722. [Google Scholar] [CrossRef]
  54. Fonseca, L.N.; Bolívar-Moná, S.; Agudelo, T.; Beltrán, L.D.; Camargo, D.; Correa, N.; Del Castillo, M.A.; Fernández De Castro, S.; Fula, V.; García, G.; et al. Cell Surface Markers for Mesenchymal Stem Cells Related to the Skeletal System: A Scoping Review. Heliyon 2023, 9, e13464. [Google Scholar] [CrossRef] [PubMed]
  55. Dominici, M.; Le Blanc, K.; Mueller, I.; Slaper-Cortenbach, I.; Marini, F.C.; Krause, D.S.; Deans, R.J.; Keating, A.; Prockop, D.J.; Horwitz, E.M. Minimal Criteria for Defining Multipotent Mesenchymal Stromal Cells. The International Society for Cellular Therapy Position Statement. Cytotherapy 2006, 8, 315–317. [Google Scholar] [CrossRef]
  56. Cao, Y.; Boss, A.L.; Bolam, S.M.; Munro, J.T.; Crawford, H.; Dalbeth, N.; Poulsen, R.C.; Matthews, B.G. In Vitro Cell Surface Marker Expression on Mesenchymal Stem Cell Cultures Does Not Reflect Their Ex Vivo Phenotype. Stem Cell Rev. Rep. 2024, 20, 1656–1666. [Google Scholar] [CrossRef]
  57. Kogan, P.-S.; Wirth, F.; Tomar, A.; Darr, J.; Teperino, R.; Lahm, H.; Dreßen, M.; Puluca, N.; Zhang, Z.; Neb, I.; et al. Uncovering the Molecular Identity of Cardiosphere-Derived Cells (CDCs) by Single-Cell RNA Sequencing. Basic. Res. Cardiol. 2022, 117, 11. [Google Scholar] [CrossRef] [PubMed]
Ijms 27 01303 g001
Figure 1. The phenotype of healthy and pathological human myocardium-derived mesenchymal stromal cells (hmMSC) and cardiosphere-derived cells (SDCs). (a) hmMSCs migrate out of cardiac explant; (b) 3D cardiospheres on non-adherent surface; (c) Morphology of healthy and pathological human myocardium-derived hmMSC; (d) Morphology of healthy and pathological SDCs. The representative micrograph of healthy hmMSCs was taken from our previous publication [20].
Figure 1. The phenotype of healthy and pathological human myocardium-derived mesenchymal stromal cells (hmMSC) and cardiosphere-derived cells (SDCs). (a) hmMSCs migrate out of cardiac explant; (b) 3D cardiospheres on non-adherent surface; (c) Morphology of healthy and pathological human myocardium-derived hmMSC; (d) Morphology of healthy and pathological SDCs. The representative micrograph of healthy hmMSCs was taken from our previous publication [20].
Ijms 27 01303 g001
Ijms 27 01303 g002
Figure 2. Cell count and surface area of healthy (not dilated) and pathological (dilated) human myocardium-derived mesenchymal stromal cells (hmMSC) and sphere-derived cells. Cell size count of healthy (a) and pathological (b) hmMSCs; (c) Major axis length of healthy and pathological hmMSCs; (d) Attachment surface area of healthy and pathological hmMSCs. Data are shown as mean ± standard deviation (SD) and are significant at * p ≤ 0.05, *** p ≤ 0.001, where n = 50 from three patients for each group measured by ImageJ 1.54p program and calculated using the Graphpad Prism 6 and Excel programs.
Figure 2. Cell count and surface area of healthy (not dilated) and pathological (dilated) human myocardium-derived mesenchymal stromal cells (hmMSC) and sphere-derived cells. Cell size count of healthy (a) and pathological (b) hmMSCs; (c) Major axis length of healthy and pathological hmMSCs; (d) Attachment surface area of healthy and pathological hmMSCs. Data are shown as mean ± standard deviation (SD) and are significant at * p ≤ 0.05, *** p ≤ 0.001, where n = 50 from three patients for each group measured by ImageJ 1.54p program and calculated using the Graphpad Prism 6 and Excel programs.
Ijms 27 01303 g002
Ijms 27 01303 g003
Figure 3. Proliferation rate of healthy and pathological myocardium-derived hmMSC and SDCs. Data are shown as mean ± standard deviation (SD) and are significant at * p ≤ 0.05 levels from not less than three repeats (n = 3) of three patients from each group as calculated using the Excel program. Data are significant comparing healthy and pathological hmMSCs and SDCs with corresponding hmMSCs.
Figure 3. Proliferation rate of healthy and pathological myocardium-derived hmMSC and SDCs. Data are shown as mean ± standard deviation (SD) and are significant at * p ≤ 0.05 levels from not less than three repeats (n = 3) of three patients from each group as calculated using the Excel program. Data are significant comparing healthy and pathological hmMSCs and SDCs with corresponding hmMSCs.
Ijms 27 01303 g003
Ijms 27 01303 g004
Figure 4. Mitochondrial membrane potential (MMP) and intracellular calcium (iCa2+) of healthy and pathological hmMSCs and SDCs. (a) Flow cytometric MMP measurement using 5,5′,6,6′-Tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC1) dye; (b) Flow cytometric iCa2+ measurement using Cal520 (Abcam, Cambridge, UK). Data are shown as mean ± standard deviation (SD). The ** p ≤ 0.01, n = 4–5 from five experiments and three patients from each group. Student t test was calculated by Graphpad Prism 6 program.
Figure 4. Mitochondrial membrane potential (MMP) and intracellular calcium (iCa2+) of healthy and pathological hmMSCs and SDCs. (a) Flow cytometric MMP measurement using 5,5′,6,6′-Tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC1) dye; (b) Flow cytometric iCa2+ measurement using Cal520 (Abcam, Cambridge, UK). Data are shown as mean ± standard deviation (SD). The ** p ≤ 0.01, n = 4–5 from five experiments and three patients from each group. Student t test was calculated by Graphpad Prism 6 program.
Ijms 27 01303 g004
Ijms 27 01303 g005
Figure 5. Mitochondrial parameters of healthy and pathological hmMSCs and SDCs. Changes of oxygen consumption rate (OCR) ((pM)/min/mg/mL)) protein in healthy (a) and pathological (b) hmMSCs and SDCs; (c) Calculated mitochondrial parameters in hmMSCs and SDCs of both cell types. Data are shown as mean ± standard deviation (SD). The * p ≤ 0.05, ** p ≤ 0.01, n = 4 from four experiments and two patients from each group. Student’s t-test was calculated by an Excel program.
Figure 5. Mitochondrial parameters of healthy and pathological hmMSCs and SDCs. Changes of oxygen consumption rate (OCR) ((pM)/min/mg/mL)) protein in healthy (a) and pathological (b) hmMSCs and SDCs; (c) Calculated mitochondrial parameters in hmMSCs and SDCs of both cell types. Data are shown as mean ± standard deviation (SD). The * p ≤ 0.05, ** p ≤ 0.01, n = 4 from four experiments and two patients from each group. Student’s t-test was calculated by an Excel program.
Ijms 27 01303 g005
Ijms 27 01303 g006
Figure 6. Glycolytic activity of healthy and pathological hmMSCs and SDCs. Changes in extracellular acidification rate (ECAR) ((mpH)/min/mg/mL)) of protein in healthy (a) and pathological (b) hmMSCs and SDCs; (c) Calculated glycolytic parameters in hmMSCs and SDCs. Data are shown as mean ± standard deviation (SD). Data are shown as mean ± standard deviation (SD). The * p ≤ 0.05, n = 4 from four experiments and two patients from each group. Student’s t-test was calculated by an Excel program.
Figure 6. Glycolytic activity of healthy and pathological hmMSCs and SDCs. Changes in extracellular acidification rate (ECAR) ((mpH)/min/mg/mL)) of protein in healthy (a) and pathological (b) hmMSCs and SDCs; (c) Calculated glycolytic parameters in hmMSCs and SDCs. Data are shown as mean ± standard deviation (SD). Data are shown as mean ± standard deviation (SD). The * p ≤ 0.05, n = 4 from four experiments and two patients from each group. Student’s t-test was calculated by an Excel program.
Ijms 27 01303 g006
Ijms 27 01303 g007
Figure 7. Expression of cardiac commitment genes in healthy (not dilated) and pathological (dilated), cardiospheres and sphere-derived cells (SDCs) vs. hmMSCs. (a) Relative gene expression in healthy SDCs and cardiospheres vs. hmMSCs; (b) Relative gene expression in pathological/dilated SDCs and cardiospheres vs. hmMSCs. The expression of genes is shown as (2−ΔCT × 100,000,000). Data are shown as mean ± standard deviation (SD). The * p ≤ 0.05, n = 3 from three experiments and two patients from each group. Student’s t-test was calculated by Excel program.
Figure 7. Expression of cardiac commitment genes in healthy (not dilated) and pathological (dilated), cardiospheres and sphere-derived cells (SDCs) vs. hmMSCs. (a) Relative gene expression in healthy SDCs and cardiospheres vs. hmMSCs; (b) Relative gene expression in pathological/dilated SDCs and cardiospheres vs. hmMSCs. The expression of genes is shown as (2−ΔCT × 100,000,000). Data are shown as mean ± standard deviation (SD). The * p ≤ 0.05, n = 3 from three experiments and two patients from each group. Student’s t-test was calculated by Excel program.
Ijms 27 01303 g007
Table 1. Analyzed gene function in heart development and disease.
Table 1. Analyzed gene function in heart development and disease.
Protein (Gene)Function SummaryTaqman
Assay Id		Key Reference (DOI)
Transcription Factors
HOPXRegulates cardiomyocyte gene networks via protein–protein interactions rather than direct DNA binding.Hs04188695_m1[41]
NKX2-5Essential for early heart development and cardiac conduction system formation.Hs00231763_m1[42]
MEF2CControls cardiomyocyte differentiation and heart morphogenesis, including outflow tract formation.Hs00231149_m1[43]
GATA4Zinc-finger transcription factor required for cardiac morphogenesis and cardiomyocyte survival.Hs00171403_m1[44]
Cell Membrane-Related Proteins
c-KIT (CD117)Tyrosine kinase receptor involved in stem/progenitor cell maintenance and cardiac repair.Hs00174029_m1[45]
NOTCH1Regulates heart development and cardiomyocyte/vascular differentiation.Hs01062014_m1[46]
THY1 (CD90)Cell surface glycoprotein involved in cardiac fibroblast regulation and cell adhesion.Hs00174816_m1[47]
TGFBR2TGF-β receptor critical for heart vessels and descending thoracic aorta developmentHs00234253_m1[48]
CDH2 (N-cadherin)Essential for cardiomyocyte adhesion and intercalated disk integrity.Hs00983056_m1[49]
GJA1 (Connexin43)Main gap junction protein for cardiomyocyte electrical coupling.Hs00748445_s1[50]
CACNA1C (CaV1.2)L-type Ca2+ channel subunit essential for cardiac excitation–contraction coupling.Hs00167681_m1[51]
Endoplasmic Reticulum-Related Proteins
ITPR2ER IP3 receptor regulating Ca2+ release in cardiac signaling and hypertrophy.Hs00181916_m1[52]
CALRER chaperone regulating calcium homeostasis and cardiac development.Hs00189032_m1[53]
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Bironaite, D.; Mikšiūnas, R. Cardiosphere-Derived Cells from Not Dilated and Dilated Human Myocardium Exhibit Enhanced Metabolic Potential Compared with Conventional Cardiac Mesenchymal Stem/Stromal Cells. Int. J. Mol. Sci. 2026, 27, 1303. https://doi.org/10.3390/ijms27031303

AMA Style

Bironaite D, Mikšiūnas R. Cardiosphere-Derived Cells from Not Dilated and Dilated Human Myocardium Exhibit Enhanced Metabolic Potential Compared with Conventional Cardiac Mesenchymal Stem/Stromal Cells. International Journal of Molecular Sciences. 2026; 27(3):1303. https://doi.org/10.3390/ijms27031303

Chicago/Turabian Style

Bironaite, Daiva, and Rokas Mikšiūnas. 2026. "Cardiosphere-Derived Cells from Not Dilated and Dilated Human Myocardium Exhibit Enhanced Metabolic Potential Compared with Conventional Cardiac Mesenchymal Stem/Stromal Cells" International Journal of Molecular Sciences 27, no. 3: 1303. https://doi.org/10.3390/ijms27031303

APA Style

Bironaite, D., & Mikšiūnas, R. (2026). Cardiosphere-Derived Cells from Not Dilated and Dilated Human Myocardium Exhibit Enhanced Metabolic Potential Compared with Conventional Cardiac Mesenchymal Stem/Stromal Cells. International Journal of Molecular Sciences, 27(3), 1303. https://doi.org/10.3390/ijms27031303

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop