ABSTRACT

   Effective treatment of ischemic disease requires the reconstruction of
   blood vessels through the delivery of angiogenic factors, such as
   chemicals, proteins, and cells. In particular, substantial efforts have
   focused on enhancing the therapeutic potential of mesenchymal stem
   cells (MSCs) for treating ischemic diseases. In this study, we
   investigated the use of electrical stimulation (ES) to potentiate the
   proangiogenic properties of human adipose‐derived MSCs. Electrically
   potentiated MSCs (epMSCs) were generated by applying optimized ES
   parameters (0.3 V, 100 Hz). EpMSCs exhibited significantly enhanced
   angiogenic potential, including upregulated expression of proangiogenic
   factors (e.g., vascular endothelial growth factor [VEGF]‐A and
   hepatocyte growth factor) and improved endothelial cell migration and
   tube formation in vitro. Transcriptomic and proteomic analyses revealed
   activation of key angiogenic pathways, particularly VEGFA–VEGFR2
   signaling, which plays a critical role in enhancing the functionality
   of epMSCs. In vivo studies using a murine hindlimb ischemia model
   demonstrated that epMSCs enhanced blood flow recovery, induced
   angiogenesis, and reduced muscle atrophy more effectively than
   unstimulated MSCs. Overall, these findings suggest that electrical
   potentiation of MSCs is a promising strategy for effectively enhancing
   their angiogenic capabilities for treating ischemic diseases.

   Keywords: mesenchymal stem cells, electrical stimulation, paracrine
   effects, angiogenesis, ischemia
     __________________________________________________________________

   This study explores the use of electrical stimulation (ES) to enhance
   the proangiogenic properties of mesenchymal stem cells (MSCs) for
   ischemic disease treatment. Electrically potentiated MSCs exhibited
   enhanced secretion of angiogenic factors, improved vascularization, and
   promoted blood flow recovery in a murine hindlimb ischemia model. These
   findings highlight ES as a promising strategy to advance MSC‐based
   therapies for ischemic tissue regeneration.

   graphic file with name MCO2-6-e70352-g003.jpg

1. Introduction

   Ischemic diseases, such as myocardial infarction, peripheral artery
   disease, and chronic wounds, result from inadequate blood circulation.
   Effective regeneration requires the formation of new blood vessels
   within ischemic tissues [[34]1]. To induce angiogenesis, various
   therapeutic substances have been transplanted, including cells (e.g.,
   endothelial cells) and angiogenic factors (e.g., vascular endothelial
   growth factor [VEGF]) [[35]2, [36]3, [37]4]. Unfortunately, several
   challenges remain unaddressed, including limited cell expansion, poor
   survival of transplanted cells, and insufficient angiogenic stimulation
   [[38]5]. Therefore, development of more effectiv and reliable
   therapeutic strategies is highly required.

   Recently, mesenchymal stem cell (MSC)‐based therapies have attracted
   significant attention for treating ischemia [[39]6]. MSCs possess
   unique capabilities of self‐renewal and differentiation into multiple
   lineages, and modulation of the microenvironment through secretion of
   various bioactive molecules [[40]7]. In particular, MSCs secrete growth
   factors, cytokines, and extracellular vesicles, which play important
   roles in tissue regeneration [[41]8, [42]9, [43]10, [44]11]. Their
   paracrine factors, such as VEGF, fibroblast growth factor (FGF), and
   angiopoietin‐1, can collectively induce a complex cascade in
   angiogenesis [[45]12]. However, the therapeutic efficacies of these
   treatments remain under pathological conditions.

   Accordingly, extensive studies have been performed to enhance MSC
   functionality by developing various approaches, including genetic
   modifications, stimulation (potentiation) with biochemical molecules,
   and the application of specialized culture conditions (e.g., 3D culture
   and hypoxia) [[46]13, [47]14, [48]15, [49]16, [50]17]. More recently,
   biophysical cues have recently emerged as promising tools for
   effectively potentiating MSCs. Biophysical cues typically include
   mechanical, electrical, magnetic, photonic, and thermal stimulations
   [[51]18]. Biophysical stimulation is cost‐effective, straightforward to
   implement, highly efficient, and reliable for modulating MSC functions
   compared with conventional biochemical stimulation [[52]19].
   Specifically, electrical stimulation (ES) with controllable parameters
   (e.g., voltage, frequency, and duration) can effectively influence MSC
   behaviors (e.g., proliferation, differentiation, and migration) by
   altering their transmembrane potential, receptor distribution,
   cytoskeleton organization, and ion influx [[53]18, [54]20].

   Previous studies demonstrated that ES can influence the differentiation
   of MSCs toward osteogenic and neurogenic lineages [[55]21, [56]22,
   [57]23, [58]24]. Additionally, ES has been reported to influence
   morphology and the expression of angiogenesis‐related markers in
   adipose‐derived MSCs [[59]25, [60]26]. However, to the best of our
   knowledge, comprehensive analyses of the effects of ES on MSCs’
   paracrine activities, specifically with respect to angiogenesis, for
   ischemic disease treatment have not been conducted.

   In this study, we aimed to determine whether ES, under appropriate
   conditions, could enhance the angiogenic potential of MSCs and their
   therapeutic efficacy in the treatment of ischemic diseases. We
   investigated the molecular and functional changes induced in
   electrically potentiated MSCs (epMSCs), including proangiogenic
   gene/protein expression and secretion of paracrine factors. In vitro
   studies were conducted to assess the ability of epMSCs to promote
   endothelial cell migration and tube formation. Additionally, in vivo
   studies were performed using a murine hindlimb ischemia (HLI) model to
   validate their efficacy in restoring blood flow, enhancing
   vascularization, and mitigating muscle atrophy (Figure [61]1).
   Complementary transcriptomic and proteomic analyses were employed to
   elucidate the molecular mechanisms underlying ES‐induced changes in MSC
   function, especially regarding angiogenesis.

FIGURE 1.

   FIGURE 1
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   Schematic illustrations of electrically potentiated MSCs (epMSCs) and
   their improved angiogenic potentials.

2. Results

2.1. Investigation of the ES Conditions

   MSCs were electrically stimulated under various conditions to evaluate
   their viability and metabolic activity. In our previous study, we
   identified that high voltage (0.6 V) significantly reduced MSC
   viability and metabolic activity, whereas 0.3 V at both 1 and 100 Hz
   maintained levels comparable to unstimulated controls [[63]27]. Based
   on these results, we selected 0.3 V with either 1 Hz (low frequency;
   LF) or 100 Hz (high frequency; HF). In this study, MSCs were subjected
   to ES for 2 h using the laboratory‐established ES system (Figure
   [64]S1) and cultured for an additional 24 h (Figure [65]2A). Live/dead
   staining revealed minimal dead cells (<3%) in all groups
   (Figure [66]2B,C). The metabolic activity of the ES group was also not
   significantly different from that of unstimulated MSCs (control)
   (Figure [67]2D). Furthermore, ES did not alter MSC morphology or
   F‐actin organization (Figure [68]S2A). Similarly, no significant
   differences in cell area (Figure [69]S2B) or aspect ratio (Figure
   [70]S2C) were observed between unstimulated and stimulated MSCs. As
   results, the selected ES parameters (0.3 V, 1, and 100 Hz) were
   cytocompatible and suitable for MSC potentiation studies.

FIGURE 2.

   FIGURE 2
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   Electrical stimulation (ES) of hAD‐MSCs. (A) Schematic illustrations of
   in vitro experimental procedures and ES parameters. A constant voltage
   of 0.3 V with low frequency (LF, 1 Hz) and high frequency (HF, 100 Hz)
   was used for ES of hAD‐MSCs. (B) LIVE/DEAD staining images of the
   unstimulated MSCs (control) or MSCs stimulated with LF or HF. Green and
   red cells represent live and dead cells, respectively. Scale
   bars = 200 µm. (C) Cell viability of hAD‐MSCs (n = 4). Cell viability
   was expressed as the percentage of live cells among the total (live and
   dead) cells. (D) Relative metabolic activity of unstimulated or
   electrically stimulated MSCs. Relative metabolic activity of each group
   was demonstrated as absorbance at 450 nm normalized by that of the
   unstimulated control (n = 4).

2.2. Expression of Angiogenic Markers in epMSCs

   Potential alterations in the angiogenic potential of MSCs induced by ES
   were investigated by assessing the mRNA expression of key angiogenic
   factors (i.e., VEGF‐A, hepatocyte growth factor [HGF], basic FGF [bFGF]
   and insulin‐like growth factor 1 [IGF‐1]) [[72]28] (Figure [73]3A).
   Their gene expression levels were significantly higher in the HF group
   than those in the unstimulated and LF groups. Specifically, ES with HF
   led to 1.3‐, 3.9‐, 2.5‐, and 7.6‐fold increases in VEGF‐A, HGF, bFGF,
   and IGF‐1 expression, respectively, compared with unstimulated
   controls. In contrast, the LF group showed no significant changes in
   VEGF‐A, bFGF, and IGF‐1, and downregulated HGF expression. Altogether,
   HF (0.3 V, 100 Hz) was determined as the appropriate ES condition for
   subsequent studies. MSCs secrete a diverse range of therapeutically
   active molecules, including growth factors, cytokines, microRNAs, and
   other small molecules [[74]29]. Hence, we quantified the secretion of
   representative proangiogenic proteins (i.e., VEGF and HGF) in the
   conditioned media (CM). The epMSCs produced greater amounts of VEGF
   (1.5‐fold) and HGF (1.3‐fold) compared with unstimulated MSCs
   (Figure [75]3B). Time‐course profiling revealed that this increase was
   rapid within 12 h and sustained for up to 72 h (Figure [76]S3A,B).
   Staining of human growth factor arrays revealed that the production of
   various secretomes from MSCs were significantly altered by ES,
   including epidermal growth factor (EGF), FGF, HGF, platelet‐derived
   growth factor (PDGF), PDGF receptor, VEGF, IGF, and IGF‐binding protein
   (Figures [77]3C and [78]S4). VEGF‐A and HGF levels were significantly
   higher in epMSCs than MSCs, which was consistent with the quantitative
   real‐time PCR (qRT‐PCR) and enzyme‐linked immunosorbent assay (ELISA)
   results (Figure [79]3A,B). Altogether, our results indicate that ES
   significantly enhances the angiogenic potential of MSCs by upregulating
   key factors, such as VEGF and HGF.

FIGURE 3.

   FIGURE 3
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   Expression of typical angiogenic markers in epMSCs. (A) Relative mRNA
   expression of VEGF‐A, HGF, bFGF, and IGF‐1 genes. Expression of each
   gene was normalized to that of GAPDH (n = 4). (B) Quantification of
   VEGF‐A and HGF produced from MSCs and epMSCs (n = 4). (C) Fold change
   in the mean pixel density of each growth factor produced by MSCs and
   epMSCs (n = 4). CMs from MSCs and epMSCs were analyzed using a human
   growth factor antibody array to detect growth factor production in
   MSCs. Gene expression was normalized to GAPDH using the 2^−ΔΔCt method.
   An asterisk (*) denotes a statistically significant difference
   (p < 0.05).

2.3. In Vitro HUVEC Culture for Evaluation of Angiogenic Functions of epMSCs

   To evaluate the proangiogenic effects of epMSCs, in vitro scratch
   closure and tube formation assays were performed by culturing HUVECs
   with CM from MSCs or epMSCs. The migration of HUVECs was quantitatively
   assessed using the scratch closure assay at 4‐h intervals
   (Figure [81]4A). HUVECs cultured with CM mixtures of MSCs or epMSCs
   showed significantly faster scratch closure after 4 h of incubation
   compared with the control; scratch closure in the epMSC CM group was
   superior to that in the MSC CM group. For instance, the epMSC CM group
   exhibited a 49.7 ± 3.4% reduction in scratch at 12 h compared with the
   initial scratch, whereas the control and MSC CM groups demonstrated
   scratch reductions of 23.8 ± 3.2 and 33.3 ± 1.1%, respectively
   (Figure [82]4B). Moreover, culture with the CM of MSCs (MSC CM and
   epMSC CM) significantly increased the tube formation of HUVECs compared
   with that with the serum‐free control group (Figure [83]4C). The epMSC
   CM group induced significantly higher numbers of branching points and
   closed loops compared with the other groups (Figure [84]4D,E). Overall,
   the results indicated that epMSCs appeared to secrete more
   proangiogenic molecules that could enhance the migration and
   tubulogenesis of HUVECs.

FIGURE 4.

   FIGURE 4
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   Functional assays of the angiogenic potentials of epMSCs with in vitro
   HUVEC culture with conditioned medium (CM). HUVECs were cultured with
   1:1 mixed serum‐free endothelial cell medium (ECM) with MEM‐α, MSC CM,
   and epMSC CM, respectively. (A) Representative optical micrographs of
   HUVEC scratches in individual groups at 0, 4, 8, and 12 h. Scale
   bar = 200 µm. (B) The corresponding scratch lengths (n = 4). (C)
   Representative optical micrographs of HUVECs tube formation in
   individual groups at 4 h. Scale bar = 200 µm. (D) The corresponding
   total branching points (n = 4) and (E) total number of loops of tubes
   at 4 h (n = 4). An asterisk (*) denotes a statistically significant
   difference (p < 0.05).

2.4. In Vivo Plug Assay

   In vivo angiogenic activity of the epMSCs was examined using a
   Matrigel‐based plug assay. Matrigel mixed with PBS (control), MSCs, or
   epMSCs was subcutaneously injected into mice, and the plugs were
   harvested on day 7 (Figure [86]S5). Plugs from the control group were
   transparent with no visible blood vessels, whereas those from the MSC
   group sparsely formed blood vessels. In contrast, the plugs from the
   epMSC group exhibited a reddish color with noticeable blood vessels
   (Figure [87]S5A). Hematoxylin and eosin (H&E) staining and von
   Willebrand factor (vWF) immunostaining revealed that the plugs of the
   control, MSC, and epMSC groups showed no vessels, a few blood vessels,
   and dense blood vessels, respectively. The hemoglobin contents were
   significantly higher in the MSC groups, especially in the epMSC group
   (Figure [88]S5B). Additionally, the vWF‐positive area in the epMSC
   group was 2.1‐fold larger than that in the MSC group (Figure [89]S5C).
   Macroscopic observations and histological analyses consistently
   demonstrated that epMSCs significantly promote blood vessel formation
   in vivo.

2.5. Transcriptomic and Proteomic Analyses of epMSCs

   Comprehensive transcriptomic and proteomic analyses were performed to
   elucidate the molecular mechanisms underlying the enhanced angiogenic
   potential of epMSCs. Quantitative RNA sequencing revealed differential
   expressions of 23,587 genes between MSCs and epMSCs (Figure [90]5A).
   Notably, 99 genes in the epMSC group exhibited 1.5‐fold or higher
   changes (p < 0.05) in expression compared with those in the MSCs group,
   in which 35 genes were upregulated and 64 genes were downregulated
   (Figure [91]5B). Gene ontology (GO) analysis identified 10 major
   ontologies involving 48 genes affected by ES. These genes were linked
   to diverse biological processes, such as cell differentiation,
   migration, angiogenesis, apoptosis, cell cycle, and various signal
   transduction pathways (Figure [92]5C). In particular, eight genes were
   associated with angiogenesis (Figure [93]5D). Among them, three genes
   (PTGS2, NR4A1, and ACKR3) inhibit angiogenesis, and five genes (NOX1,
   KLF5, NDNF, MEOX2, and EGR3) promote angiogenesis [[94]30, [95]31,
   [96]32, [97]33, [98]34, [99]35, [100]36]. Although NOX1 and EGR3
   expressions decreased, overall gene expression profile suggested a
   proangiogenetic tendency. The interactions among proteins corresponding
   to differentially expressed genes (DEGs) are demonstrated in Figure
   [101]S6. Analysis of functional enrichment and clustering revealed that
   blood vessel development (GO:0001568), with proteins highlighted in
   red, was the most prominently enriched among the identified biological
   processes within the network. These results suggest that ES induces MSC
   gene expression associated with angiogenesis.

FIGURE 5.

   FIGURE 5
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   Quantitative RNA sequencing analysis. (A) Volcano plot of
   differentially expressed genes (DEGs). The red and blue dots indicate
   the upregulated and downregulated genes in the epMSCs compared with
   those in the MSCs, respectively. (B) Hierarchical clustering analysis
   of DEGs between the MSCs and epSCs with a color scale indicating
   upregulation (red) and downregulation (blue). (n = 3 per group). (C)
   Ten major gene ontology (GO) analysis of the genes of which expressions
   were significantly influenced by ES. (D) Gene expression profile
   associated with GO terms related to angiogenesis. Genes depicted in
   blue and red indicate upregulation and downregulation of angiogenesis,
   respectively.

   Proteomics analysis was performed to quantitatively assess the proteins
   secreted by MSCs and epMSCs. A total of 435 proteins were identified,
   including 418 proteins identified in both groups. Moreover, 14 and 8
   proteins were identified exclusively in the epMSC and MSC groups,
   respectively (Figure [103]6A). With a fold‐change (>1.5), 42 proteins
   showed high abundance, and 49 proteins showed low abundance in the
   epMSC group (Figure [104]6B). Proteins with high or low abundance in
   epMSCs were further categorized based on GO (Figure [105]6C). GO
   analysis of the differentially expressed proteins indicated the
   upregulation of proteins involved in several important biological
   processes, including positive regulation of cell proliferation,
   extracellular matrix organization, angiogenesis, ERK1/2 signaling, and
   protein kinase B (Akt) signaling, and cell surface receptor signaling.
   Pathway enrichment analysis further revealed significant connections
   between the epMSC secretome and key proangiogenic pathways [[106]37].
   The top 10 enriched pathways included the complement system, complement
   and coagulation cascades, and the VEGFA–VEGFR2 signaling pathway
   (Figure [107]6D). Moreover, several proangiogenic (e.g., PLAUR)
   proteins and immune‐modulating proteins (e.g., CCL2 (MCP‐1) and Annexin
   A1 (ANXA1)) were identified in the epMSC secretome. Collectively, these
   findings demonstrate that ES enhances the angiogenic potential of MSCs
   by influencing multiple interconnected pathways for sustained vascular
   growth.

FIGURE 6.

   FIGURE 6
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   Proteomics analysis of MSCs and epMSCs. (A) Venn diagram illustrating
   the distribution of proteins identified the CM of MSCs and epMSCs. (B)
   Hierarchical clustering analysis of the differentially expressed
   proteins between MSC CM and epMSC CM with a color scale indicating
   downregulation (blue) and upregulation (red). (C) GO analysis
   highlighting the top 10 biological processes significantly altered by
   ES with. (D) Functional enrichment analysis of differentially expressed
   proteins with the top 10 significantly enriched GO terms related to
   signaling pathways.

2.6. Assessment of Therapeutic Efficacy of epMSCs in a HLI Model

   The therapeutic efficacy of epMSCs was examined using a mouse HLI
   model. The experimental groups included PBS (control), unstimulated
   MSCs, and epMSCs. Each sample was injected intramuscularly into
   ischemic limbs (Figure [109]S7). The extent of limb salvage was
   monitored in the feet of HLI‐induced mice and quantified on day 14
   (Figures [110]7A,B and [111]S8). By day 3, blackened paws were observed
   in all experimental groups, which progressed to toe loss by day 7 in
   the control and MSC groups. By day 14, foot and toe necrosis were
   prevalent in the control and MSC groups. In contrast, the epMSC group
   exhibited minimal necrosis and no limb loss. Laser Doppler perfusion
   imaging was performed to monitor blood flow over a 2‐week period after
   the operation (Figure [112]7C). On day 3, blood perfusion was minimal
   in all groups (Figure [113]7D). However, on days 7 and 14, blood
   perfusion ratios gradually recovered in all groups. The epMSC group
   consistently exhibited the highest blood perfusion ratio among all
   groups. On day 14, blood flow ratios were 0.21 ± 0.03 (control),
   0.36 ± 0.04 (MSC), and 0.67 ± 0.11 (epMSC) (Figure [114]7E). These
   results suggest that epMSCs effectively promote angiogenesis,
   preventing limb necrosis and recovering blood flow in HLI.

FIGURE 7.

   FIGURE 7
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   Assessment of the therapeutic activity of epMSCs in a mouse hindlimb
   ischemic model. (A) Representative photographs of ischemic hindlimbs on
   days 3, 7, and 14 after surgery and treatment. (B) Limb assessment on
   day 14. (C) Laser Doppler perfusion imaging of the ischemic hindlimbs
   on days 3, 7, and 14. (D) Blood perfusion ratio of the ischemic limb in
   each group (n = 6). An asterisk (*) and a crosshatch (#) denote
   statistically significant difference (p < 0.05) when compared with the
   control and MSC groups, respectively. (E) Blood perfusion ratios of
   each group on day 14 (n = 6). An asterisk (*) denotes a statistically
   significant difference (p < 0.05).

   Ischemic injury typically causes the apoptosis of skeletal muscle
   fibers and structural damage [[116]38]. In our study, because MSCs can
   enhance blood circulation and facilitate improved limb salvage by
   secreting various angiogenic growth factors and immunomodulatory
   substances in the ischemic limb, we expected that transplantation of
   MSCs or epMSCs would alleviate muscle atrophy [[117]32]. Notable
   infiltration of inflammatory cells and structural disorganization were
   observed in gastrocnemius tissues from the control group. Conversely,
   the MSC group displayed lower cell infiltration and moderate
   vascularization. The epMSC group exhibited structural integrity, blood
   vessels, and centrally located nuclei within the muscle fibers
   (Figure [118]8A), which are widely recognized indicators of appropriate
   muscle regeneration [[119]39].

FIGURE 8.

   FIGURE 8
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   Histological analysis of the gastrocnemius and quadriceps muscles
   harvested on day 14 after surgery and treatment. (A) Representative
   images of H&E, Masson's trichrome (MT), and vWF staining of
   gastrocnemius in each group. Red and white arrows indicate blood
   vessels. (B) Fibrosis (n = 5) and (C) vWF‐positive areas (n = 5).
   vWF‐positive area was normalized to the DAPI‐positive area in each
   group. (D) Representative images of H&E, MT, vWF, and HNA staining of
   quadriceps in each group. (E) vWF‐positive (n = 5) and (F) HNA‐positive
   areas (n = 5). vWF‐positive or HNA‐positive areas were normalized to
   the DAPI‐positive areas in each group. Scale bars = 100 µm. An asterisk
   (*) denotes a statistically significant difference (p < 0.05).

   Fibrosis was significantly reduced in the MSC and especially epMSC
   groups. For example, the fibrosis areas of the control, MSC, and epMSC
   groups were 33.8 ± 5.9, 23.5 ± 2.8, and 11.9 ± 2.6, respectively
   (Figure [121]8B). In addition, Immunostaining for vWF confirmed
   increased vascularization in epMSC‐treated muscles (Figure [122]8C).
   The vWF‐positive area in the epMSC group was 6.8‐ and 3.8‐fold larger
   than that in the control and MSC groups, respectively. Similarly, the
   quadriceps muscles of the MSC‐treated group displayed improved
   structural integrity, reduced cell infiltration, and increased blood
   vessel formation (Figure [123]8D). Interestingly, fat deposition,
   typically caused by metabolic and musculoskeletal damages [[124]40],
   was observed in the control and MSC groups, but markedly reduced in the
   epMSC group. Additionally, larger areas of vWF‐positive cells were
   found in the MSC (25.0 ± 3.2%) and epMSC (38.3 ± 7.1%) groups than in
   control group (13.0 ± 5.1%) (Figure [125]8E). Human nuclear antigen
   (HNA) immunostaining revealed the enhanced viability and residence of
   the transplanted MSCs (Figure [126]8F). HNA‐positive areas were
   8.8 ± 2.4 and 12.9 ± 2.8% for the MSC and epMSC groups, respectively.
   Costaining for HNA and vWF revealed limited colocalization (Figure
   [127]S9). The percentages of vWF‐positive area within the HNA‐positive
   population were 12.2 ± 2.2 and 13.2 ± 2.5% in the MSC and epMSC groups,
   respectively, in which the difference was not significant. These
   results suggest that most transplanted cells contributed to
   vascularization through paracrine mechanisms rather than direct
   endothelial differentiation.

3. Discussion

   Although ES can modulate cell behavior, inappropriate ES parameter can
   generate harmful by‐products, including hydrogen peroxide, hydroxyl
   ions, and other intermediates, potentially leading to cellular damage
   [[128]41]. However, our in vitro studies confirmed the optimized ES
   conditions (0.3 V, 100 Hz for 2 h) did not impair cell viability,
   morphology, or metabolic activity.

   The major finding of this study is that ES significantly increases the
   expression and secretion of key proangiogenic factors, including
   VEGF‐A, HGF, bFGF, and IGF‐1. Among these, VEGF‐A and HGF are
   well‐established regulators of vascular growth and remodeling. The
   upregulation of these angiogenic genes and proteins indicates that ES
   rapidly activates intracellular signaling pathways and sustains the
   production of angiogenic factors. Notably, the enhancement was
   frequency dependent. HF stimulation induced greater gene and protein
   expression, which may be attributed to more efficient intracellular
   penetration and transcriptional activation [[129]42].

   Functional assays further confirmed that CM from epMSCs significantly
   enhanced migration and tubulogenesis of HUVECs. These in vitro results
   are in a good agreement with the in vivo Matrigel plug results, where
   epMSCs significantly increased hemoglobin content and vessel density.
   In the HLI model, epMSC treatment improved blood perfusion, reduced
   tissue necrosis, and attenuated muscle fibrosis. The observed
   histological improvements indicate the proangiogenic and
   tissue‐protective effects of epMSCs. Moreover, limited colocalization
   of HNA and vWF in transplanted tissues suggests that epMSC‐mediated
   therapeutic effects are predominantly paracrine rather than through
   direct differentiation.

   To elucidate the molecular mechanisms underlying these enhanced
   effects, transcriptomic and proteomic analyses were performed. RNA
   sequencing identified differentially expressed genes associated with
   angiogenesis, apoptosis, and immune responses. Especially,
   proangiogenic genes such as KLF5 and NDNF were upregulated, while
   angiogenesis‐inhibitory genes were downregulated. Proteomic analysis
   supported these findings, highlighting increased expression of proteins
   related to the VEGFA–VEGFR2, PI3K/Akt, and ERK1/2 pathways, which play
   important roles in endothelial cell functions [[130]43, [131]44]. In
   addition, secretome profiling identified the upregulation of PLAUR,
   CCL2, and ANXA1, which are known to mediate extracellular matrix
   remodeling, immune modulation, and endothelial stabilization,
   respectively. These overall trends in proangiogenic transcriptional and
   proteomic profiles can explain the enhanced angiogenic potential of
   epMSCs observed both in vitro and in vivo.

   Importantly, ES significantly enhanced the regenerative potential of
   MSCs by promoting their angiogenic capacity and reducing tissue damage
   in various ischemic diseases, such as peripheral artery disease or
   myocardial infarction. This approach offers a cost‐effective, scalable
   strategy that can be integrated into current MSC manufacturing
   processes. The ES parameters used in this study (low‐voltage: 0.3 V;
   high‐frequency: 100 Hz) are safe and clinically feasible. While we
   demonstrated therapeutic efficacy 14 d postsurgery and treatment in
   this study, the long‐term studies will be necessary to assess the
   sustainability of these improvements. For clinical translation of
   epMSC, several issues should be addressed, including the establishment
   standardized ES protocols and production conditions, long‐term safety
   and efficacy, and validation in large animal models.

4. Conclusion

   This study demonstrated that ES significantly enhances the angiogenic
   potential of MSCs, presenting a promising strategy for MSC‐based
   therapies for ischemic tissue regeneration. EpMSCs exhibited
   upregulation of key angiogenic factors, such as VEGF‐A and HGF, leading
   to enhanced endothelial cell migration and tube formation in vitro.
   Comprehensive transcriptomic and proteomic analyses further revealed
   that ES substantially activates angiogenic signaling pathways,
   including the VEGFA–VEGFR2 signaling. In vivo studies revealed that
   epMSC transplantation substantially promoted angiogenesis, enhanced
   blood flow recovery, and reduced muscle atrophy in ischemic tissues
   compared with unstimulated MSCs. We demonstrated the therapeutic
   efficacy of epMSCs and the mechanisms underlying their enhanced
   angiogenic capabilities. Altogether, this study establishes ES as a
   powerful approach to enhance the regenerative properties of MSCs,
   thereby advancing their therapeutic potential for treating ischemic
   diseases. Future investigations are necessary to optimize the ES
   parameters and elucidate the underlying mechanisms to fully harness the
   benefits of epMSCs in clinical applications.

5. Materials and Methods

5.1. Cell Culture

   Human adipose‐derived MSCs (hAD‐MSCs) (PromoCell, Heidelberg, Germany)
   were maintained in MEM‐α (Gibco, Grand Island, NY, USA) supplemented
   with 10% fetal bovine serum (FBS; Gibco) and 1% antibiotic–antimycotic
   (100×; Gibco) in an incubator with 5% CO[2] at 37°C used at passage
   number 6. HUVECs (Gibco) were cultured in endothelial culture medium
   (ECM; SceinCell, CA, USA) supplemented with 5% FBS, 1%
   penicillin–streptomycin, and 1% endothelial cell growth supplement in
   an incubator with 5% CO[2] at 37°C and used at passage number 6.

5.2. Electrical Stimulation

   hAD‐MSCs were seeded onto sterile stainless‐steel plates
   (3.8 × 2.8 × 0.7 cm) at a density of 2 × 10^4 cells/cm^2 and incubated
   for 24 h at 37°C with 5% CO[2]. The cells were randomly divided into
   control (unstimulated) and two ES groups. A two‐electrode system was
   employed, in which a platinum mesh (5 × 10 mm) and the cell‐seeded
   stainless‐steel plate served as the counter and working electrodes,
   respectively, using a multichannel potentiostat (VersaSTAT3; Princeton
   Applied Research, Oakridge, TN, USA). Stimulation was applied at 0.3 V,
   5 ms pulse duration, and frequencies of 1 Hz (LF) or 100 Hz (HF) for
   2 h, according to previous studies with some modifications [[132]45,
   [133]46, [134]47, [135]48].

5.3. In Vitro Cytocompatibility Test

   One day after ES, the viability and metabolic activity of MSCs were
   assessed. Cell viability measured using the LIVE/DEAD
   viability/cytotoxicity kit (Invitrogen, Carlsbad, CA, USA) according to
   the manufacturer's protocol. Fluorescence images were acquired using a
   fluorescence microscope (DMI3000 B; Leica). Cell viability was
   expressed as the percentage of live (green) cells among total cells.
   The metabolic activity was measured using WST‐1 assay (DoGen, Seoul,
   Republic of Korea). After 1 h of incubation in WST‐1 solution (1:10 in
   the media) at 37°C, the absorbance of the supernatant at 450 nm was
   measured using a microplate reader (Varioskan LUX; Thermo Fisher
   Scientific) and were normalized to the unstimulated group.

5.4. Cellular Morphology

   Cells were fixed in 3.7% w/v paraformaldehyde at 25°C for 20 min,
   permeabilized/blocked (0.1% v/v Triton X‐100, 4% w/v BSA, and 2% v/v
   goat serum in Dulbecco's phosphate buffered saline [DPBS]; Gibco) at
   25°C for 1 h, and stained with Alexa Fluor 488‐conjugate phalloidin
   solution (1:200; Thermo Scientific) at 25°C for 30 min in the dark.
   Nuclei were stained with DAPI (1:1000 in DPBS) at 25°C for 3 min.
   Fluorescence images were acquired using a fluorescence microscope. Cell
   area and aspect ratio were measured using ImageJ software by tracing
   cell boundaries. For each group, measurements were obtained from 20 to
   40 cells selected from five randomly chosen fields per replicate
   (n = 4).

5.5. Quantitative RT‐PCR Analysis

   After 24‐h culture in the growth medium, total RNA was extracted from
   unstimulated and stimulated cells using TRIzol (Thermo Scientific). RNA
   concentration and purity were assessed by UV–visible spectroscopy
   (BioDrop Duo; BioDrop, UK). Complementary DNA was synthesized using a
   High‐Capacity cDNA Reverse Transcription Kit (Applied Biosystems).
   qRT‐PCR was performed using Power SYBR Green PCR Master Mix and a
   StepOnePlus Real‐Time PCR system under standard cycling conditions:
   95°C for 10 min, followed by 40 cycles of 95°C for 15 s and 60°C for
   1 min. Expression levels of VEGF, HGF, bFGF, and IGF‐1 were normalized
   to glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) using the 2^−ΔΔCt
   method [[136]49]. Primer sequences are listed in Table [137]S1. All
   reactions were performed in triplicate, and results were presented as
   fold changes relative to the unstimulated control.

5.6. Collection of CM

   Two hours after ES, cells were washed twice with DPBS and incubated in
   serum‐free MEM‐α for additional 72 h. CM were collected, centrifuged at
   2000 rpm for 30 min at 4°C to remove debris, filtered using a 0.2 µm
   filter, and stored at −80°C for further analysis.

5.7. Enzyme‐Linked Immunosorbent Assay

   Human VEGF‐A and human HGF in the CM were quantified using ELISA kits
   (Elabscience, Houston, TX, USA) according to the manufacturer's
   protocols. The absorbance at 450 nm was measured using a microplate
   reader.

5.8. Growth Factor Antibody Array

   Growth factors in the CM were analyzed using a Human Growth Factor
   Antibody Array Kit (RayBiotech, GA, USA) according to the
   manufacturer's protocols. Antibody‐printed membranes were incubated in
   blocking buffer at 25°C for 30 min, incubated overnight with CM at 4°C,
   then sequentially incubated with biotinylated antibody cocktail for 2 h
   and HRP–streptavidin solution at 25°C for 2 h. Detection buffer was
   applied onto the membranes [[138]50]. Signals were detected using a
   chemiluminescence imaging system (Gel doc; Bio‐Rad, ChemiDoc XRS) with
   Image Lab Software. Signal intensities were normalized to those of
   positive controls.

5.9. Quantitative RNA Sequencing and Data Analysis

   Total RNA was extracted from 1 × 10^6 cells and subjected to
   Quantitative RNA sequencing (Quant‐Seq) analysis (Ebiogen, Seoul,
   Republic of Korea). Raw read counts were normalized using the Trimmed
   Mean of M‐values method and transformed to count‐per‐million. The
   results were analyzed using ExDEGA software (Ebiogen). DEGs
   (fold‐changes ≥ 1.5, p ≤ 0.05) were visualized as a volcano plot.
   Pathway enrichment analysis was performed using the DAVID
   ([139]http://david.abcc.ncifcrf.gov/) and QuickGO
   ([140]https://www.ebi.ac.uk/QuickGO/) databases. Ninety‐nine genes were
   represented as a heatmap using the MultiExperiment Viewer software (J.
   Craig Venter Institute, Rockville, MD, USA, version 4.9.0). Ten key GO
   biological processes were identified. Protein–protein interactions
   among DEGs were analyzed using the STRING v12.0, web tool
   ([141]http://string‐db.org/), with k‐means clustering (three clusters).

5.10. Proteomics Analysis

   CM were concentrated using a Vivaspin Turbo 15 RC (3 kDa cutoff;
   Satorius, Göttingen, Germany). Protein concentration was measured using
   the Bradford assay. Proteins (30 µg) were reduced with
   1,4‐dithiothreitol (Sigma–Aldrich, St. Louis, MO, USA), alkylated with
   iodoacetamide (Sigma–Aldrich), and digested overnight with trypsin
   according to the literature [[142]51]. Samples were desalted using C18
   spin columns (Thermo Scientific) and analyzed by liquid
   chromatography‐tandem mass spectrometry using an Ultimate 3000 RSLC
   system coupled with a Q‐Exactive Orbitrap Plus mass spectrometer
   (Thermo Scientific). Peptides were separated with a 0.1% formic acid
   gradient at 3 µL/min. Data were processed using Thermo Scientific
   Xcalibur software, and protein identification/quantification was
   performed using Proteome Discoverer (Thermo Fisher Scientific).
   Intensities were normalized using the total peptide amount method,
   log2‐transformed, and median‐centered. Pathway enrichment analysis was
   performed using WikiPathway gene sets in Enrichr‐KG
   ([143]https://maayanlab.cloud/enrichr‐kg/, adjusted p < 0.05).
   Protein–protein interactions were analyzed using STRING v12.0. All
   analyses were performed in triplicate.

5.11. Scratch Closure Assay

   HUVECs (5 × 10^4 cells/well) were seeded in 24‐well plates and cultured
   until 90% confluence. A scratch was made using a 200 µL pipette tip,
   and detached cells were removed by DPBS washing. CM from MSCs or epMSCs
   was mixed with serum‐free ECM and added to the cell plates. A 1:1
   mixture of serum‐free MEM‐α and serum‐free ECM was used as the control.
   Cells were incubated at 37°C with 5% CO[2]. Wound closure was
   periodically monitored via optical microscopy (DMI3000B; Leica) for up
   to 12 h.

5.12. Tube Formation Assay

   HUVECs were cultured in a serum‐free medium for 12 h and seeded onto
   Matrigel (Corning, New York City, NY, USA)‐coated 24‐well plates
   (1.2 × 10^5 cells/well). A 1:1 mix of serum‐free ECM with either
   serum‐free MEM‐α, MSC CM, or epMSC CM was added. Cells were incubated
   at 37°C with 5% CO[2] for 8 h. Optical micrographs were acquired 4 h
   after seeding. ImageJ software (National Institute of Health, Bethesda,
   MD, USA) was used to analyze the number of branching points and loops.

5.13. In Vivo Matrigel Plug Assay

   All animal experiments were approved by the Committee on Animal
   Research and Ethics of Gwangju Institute of Science and Technology
   (GIST, Republic of Korea (approval number: GIST‐2021‐086)).
   Six‐week‐old male BALB/c athymic nude mice were anesthetized using 2%
   isoflurane (Ankuk Inc., Republic of Korea). Matrigel mixed with either
   unsimulated MSCs or epMSCs (2 × 10^5 cells/mL, 500 µL) was
   subcutaneously injected into both sides of the backs using an 18 G
   needle (n = 4 per group). One week after injection, the plugs were
   retrieved and analyzed by H&E staining, immunostaining for vWF, and
   hemoglobin quantification according to the literature [[144]52].

5.14. HLI Model

   Eight‐week‐old male BALB/c athymic nude mice were anesthetized using 2%
   isoflurane. Ischemia was induced through double‐knot ligation of the
   femoral and saphenous artery using 8‐0 silk sutures. MSCs (1.5 × 10^6
   cells in 50 µL DPBS) were injected into the adductor muscle
   (approximately 3 mm depth) using a 27‐gauge needle. The incision was
   closed using 6‐0 silk suture. Experimental groups included control
   (PBS), MSCs, and epMSCs.

5.15. Laser Doppler Perfusion Imaging and Limb Salvage Measurement

   Hindlimb blood perfusion was assessed on days 0, 3, 7, and 14 after
   surgery using PeriScan PIM 3 (Perimed AB, Stockholm, Sweden). Mice were
   anesthetized via intraperitoneal injection of ketamine/xylazine mixture
   (4:1, 300 µL). Blood flow from the knee joint to the toe (2.5 × 3.5 cm)
   was measured from color‐coded digital images. On day 14, limb condition
   was graded as limb loss, foot necrosis, toe necrosis, and limb salvage
   (n = 6 per group).

5.16. Histological Analyses of Ischemic Limbs

   On day 14, gastrocnemius and quadriceps of ischemic limbs were
   harvested, fixed, dehydrated, and paraffinized [[145]53]. Tissue
   sections (5 µm) were stained with H&E and MT, imaged using a VS200
   Research Slide Scanner (OLYMPUS, Tokyo, Japan). Fibrosis was quantified
   from the MT‐stained images as the percentage of the fibrotic (blue)
   area relative to the injured area using ImageJ. For immunofluorescence
   staining, the sections were incubated in the blocking solution (3% goat
   serum in DPBS) at 25°C for 1 h, incubated in the primary antibody
   solution (anti‐vWF (Chemicon, Temecula, CA, USA)) or anti‐HNA (Abcam,
   Cambridge, UK); 1:100 in 0.05% Triton‐X 100 and 1.5% goat serum) at 4°C
   overnight, washed twice with DPBS, and incubated in Alexa Fluor 555‐ or
   Alexa Fluor 488‐conjugated secondary antibody solution (1:200 in DPBS;
   Invitrogen) at 37°C for 1 h. Autofluorescence was quenched with 0.1%
   Sudan black B, and nuclei were stained with DAPI. Fluorescence images
   were acquired using a Research Slide Scanner. The vWF‐ or HNA‐positive
   areas were measured using ImageJ and normalized to the corresponding
   DAPI‐positive areas.

5.17. Statistical Evaluation

   All tests were performed at least three times and results were reported
   as mean ± standard deviation unless otherwise noted. Data normality was
   assessed using the Shapiro–Wilk test. Group comparisons were performed
   using one‐way analysis of variance followed by Tukey's posthoc test for
   multiple comparisons in OriginPro software (OriginLab Corporation,
   Northampton, MA, USA). For transcriptomic and proteomic analyses, p
   values were adjusted for the false discovery rate method, in which a
   false discovery rate <0.05 was considered statistically significant.
   Additional statistical details are provided in each relevant
   subsection.

Author Contributions

   Jongdarm Yi: conceptualization, data curation, formal analysis,
   investigation, methodology, validation, visualization, writing—original
   draft. Seungjun Lee: data curation, formal analysis, investigation,
   methodology, validation, visualization, writing—review and editing, and
   writing—original draft. Chiseon Ryu: formal analysis, investigation,
   and validation. Gaeun Kim: methodology, investigation, and validation.
   Junghyun Kim: investigation and validation. Jae Young Lee: supervision,
   writing—review and editing, writing—original draft, conceptualization,
   and funding acquisition. All authors have read and approved the final
   manuscript.

Ethics Statement

   All experiments involving animals were approved by the Committee on
   Animal Research and Ethics of Gwangju Institute of Science and
   Technology (GIST, Republic of Korea (approval number: GIST‐2021‐086)).

Conflicts of Interest

   All authors declare no conflicts of interests.

Supporting information

   Supporting Table 1: List of primer sequences for real‐time qPCR.
   [146]MCO2-6-e70352-s004.docx^ (6.6MB, docx)

   Supporting Fig 1: (a) Schematic illustration and (b) real images of the
   lab‐established ES system. A SUS316L stainless steel plate and platinum
   mesh were used as a working electrode and a counter electrode,
   respectively.
   [147]MCO2-6-e70352-s003.tif^ (1,010.7KB, tif)

   Supporting Fig 2: (A) Fluorescence staining for F‐actin and nuclei of
   electrically non‐stimulated or stimulated MSCs on stainless steel
   plates. Scale bar = 200 µm. Quantitative analyses of (B) cell area and
   (C) aspect ratio between electrically stimulated and non‐stimulated
   MSCs. For each group, measurements were obtained from 20–40 cells
   selected from five randomly chosen fields per replicate (n = 4). An
   asterisk (*) denotes a statistically significant difference (p < 0.05).
   [148]MCO2-6-e70352-s005.tif^ (2.1MB, tif)

   Supporting Fig 3: (A) Quantification of VEGF and (B) HGF secretion in
   conditioned media from unstimulated MSCs and epMSCs at 12, 24, 48, and
   72 hours post‐stimulation, measured by ELISA (n=4). An asterisk (*)
   denotes a statistically significant difference (p < 0.05) compared to
   the corresponding MSC control at each time point.
   [149]MCO2-6-e70352-s007.tif^ (358.1KB, tif)

   Supporting Fig 4: Growth factor secretion profiles of the MSCs and
   epMSCs. The rectangles highlight the expression of EGF, PDGF family,
   IGFBP family, HGF, and VEGF‐A.
   [150]MCO2-6-e70352-s006.tif^ (1.1MB, tif)

   Supporting Fig 5: In vivo Matrigel plug assay. (A) Representative
   images of the gross morphology, H&E, and vWF immunofluorescence of the
   plugs retrieved at 7 days after transplantation. Arrows indicate
   vessel‐like structures. Scale bars are 5mm, 100µm, and 200µm for
   morphology, H&E, and vWF/DAPI immunofluorescence images, respectively.
   (B) Hemoglobin contents in the plugs of the individual groups (n=4).
   Hemoglobin contents were normalized by the weights of harvested plugs.
   (C) vWF‐positive area, normalized by the 4',6‐diamidino‐2‐phenylindole
   (DAPI)‐positive areas in each group (n=4). An asterisk (*) denotes a
   statistically significant difference (p < 0.05).
   [151]MCO2-6-e70352-s010.tif^ (1.4MB, tif)

   Supporting Fig 6: STRING analysis of the protein networks identified 84
   protein nodes among 99 DEGs. Ten proteins corresponding to the blood
   vessel development (GO:0001568) are highlighted in red.
   [152]MCO2-6-e70352-s002.tif^ (2.5MB, tif)

   Supporting Fig 7: Schematic representation of intramuscular MSC
   injection in the murine hindlimb ischemia (HLI) model. MSC were
   performed intramuscularly into the ischemic limb, specifically
   targeting the central region of the adductor muscle group. The
   injection was conducted approximately 3 mm deep into the muscle tissue
   to ensure accurate cell delivery and optimal engraftment.
   [153]MCO2-6-e70352-s008.tif^ (1.4MB, tif)

   Supporting Fig 8: Photographs of ischemic hindlimbs from all of the
   groups on Day 14 after transplantation (n=6).
   [154]MCO2-6-e70352-s001.tif^ (2MB, tif)

   Supporting Fig 9: Co‐localization analysis of HNA positive cells with
   endothelial marker (vWF) (n=5).
   [155]MCO2-6-e70352-s009.tif^ (1.2MB, tif)

Acknowledgments