Visualizing Age-Dependent Electrotaxis of Human Adipose-Derived Stem Cells Under Direct Current Electric Fields

Adipose-derived stem cells (ADSC) possess strong self-renewal capacity and significant regenerative potential. Under appropriate induction conditions, they can differentiate into osteoblasts, chondrocytes, and adipocytes, among others¹. Through paracrine signaling, ADSC can also support the generation of blood vessels and lymphatic vessels^2,3, inhibit scar formation⁴, and exhibit immune regulatory effects⁵. Therefore, ADSC is considered an ideal seed cell for cell-based therapies in regenerative medicine.

The function of ADSC is influenced by various donor characteristics, which may affect their potential efficacy in cell therapies, including donor age, body mass index, gender, and others⁶. ADSC derived from older donors exhibit more pronounced aging characteristics, with reduced adipogenic differentiation potential, osteogenic differentiation potential, and migratory capacity^7,8,9. Furthermore, the ability to promote keratinocyte proliferation and stimulate the body's self-repair mechanisms is diminished¹⁰.

Cell-directed migration is a key physiological mechanism in wound healing and tissue regeneration, with electric fields (EFs) playing an important role in guiding cell migration and promoting wound repair¹¹. For example, when the epithelial barrier is damaged, endogenous electric fields are immediately generated and persist until the wound heals and the epithelial barrier function is restored^12,13,14. As one of the signals guiding directed cell migration, electric signals have been shown to have a higher priority than other coexisting signals, such as contact inhibition, mechanical forces, and chemotactic factors^15,16. Several methods have been developed to promote wound healing and tissue regeneration, such as through drugs and external electric stimulation devices that enhance or mimic the endogenous electric fields in the human body^17,18. The ability of cells to migrate according to an electric field gradient is known as electrotaxis. In vitro studies have shown that many different types of cells are capable of directed migration in electric fields. Some cells migrate toward the anode, such as mammalian cranial neural crest cells, human astrocytes, and human breast cancer cells^19,20,21. Other cells migrate toward the cathode, such as human renal tubular epithelial cells, human dermal fibroblasts, and human neutrophils^22,23,24. ADSC can be guided to migrate toward the anode in direct current (DC) electric fields, and under prolonged culture conditions, exogenous pulsed electric current has been shown to effectively promote the directional migration of ADSC toward the anode while maintaining cell viability²⁵. To date, studies on the electrotaxis characteristics of human adipose-derived stem cells remain scarce, and the underlying mechanisms regulating their electrotactic behavior are still not fully understood.

We are the first to systematically investigate age-dependent electrotactic differences in hADSCs from young versus elderly female donors under gradient direct current electric fields (DCEF), and to further uncover the underlying mechanisms (involving sodium channel activity and PI3K-Akt signaling) using transcriptomic analysis. In this study, we examined the effects of DCEF on hADSCs. Using RNA sequencing and differential gene enrichment analysis, we compared gene expression profiles between two hADSC groups-derived from young and elderly female donors-following DCEF stimulation to identify age-related transcriptional differences. We hypothesized that donor aging impairs the electrotactic responsiveness of hADSCs, and that this age-related decline is linked to altered sodium channel activity and PI3K-Akt signaling.

Informed consent was obtained from all participants prior to tissue collection. This study was approved by the Ethics Committee of the First Affiliated Hospital of China Medical University (Approval No.: [2018]2018-110-2). Written informed consent was obtained from all participants prior to tissue collection

hADSC isolation and culture
Tissue collection: Human abdominal adipose tissue samples were obtained during routine elective liposuction procedures performed at the Department of Plastic Surgery of the First Affiliated Hospital of China Medical University. All donors were healthy female volunteers undergoing cosmetic abdominal liposuction, and no additional surgical interventions were performed for research purposes. Tissue specimens were collected by the operating surgeon under sterile conditions immediately after aspiration and transferred to the laboratory in sterile containers on ice within 30 min.

Sample size: A total of 6 donors were included in this study, consisting of 3 younger donors (age: 27.00 ± 4.58 years) and 3 older donors (age: 62.33 ± 4.04 years).

Tissue processing: Transfer adipose tissue fragments into a 50 mL conical tube, centrifuge at 1,800 x g for 3 min at 4 °C, and carefully aspirate the upper oil phase (to remove excess lipids). Add 0.2% collagenase Type IV (final volume: 5-10 mL per 1 g adipose tissue) to the tissue pellet, then place the tube in a 37 °C water bath or incubator for 45 min. Gently agitate the tube every 10-15 min during digestion to ensure uniform contact between collagenase and tissue. After digestion, terminate the reaction by adding low-glucose DMEM supplemented with 10% FBS and 1% penicillin/streptomycin (volume ratio of neutralization medium to collagenase solution = 1:1).

CAUTION: When handling collagenase, wear disposable gloves, a lab coat, and safety goggles to avoid skin and eye contact, prepare and use the reagent in a biological safety cabinet (BSC) to prevent aerosol contamination, and in case of spillage, wipe up immediately with 75% ethanol and dispose of contaminated materials as biological waste.

Cell isolation: Centrifuge the neutralized digestion mixture at 1,200 x g for 10 min at 4 °C to pellet the cells. Discard the supernatant, then resuspend the pellet in erythrocyte lysis buffer (155 mM NH₄Cl, 10 mM KHCO₃, 0.1 mM EDTA; pH 7.4; volume: 2-3 mL per pellet) and incubate at room temperature (22-25 °C) for 5 min to lyse residual red blood cells. Filter the cell suspension sequentially through 200 µm strainers to remove undigested tissue debris and obtain a single-cell suspension.

Cell counting: Before seeding, cell numbers were quantified using a hemocytometer. Briefly, hADSCs were detached with 0.25% trypsin-EDTA, resuspended in growth medium, and mixed 1:1 with 0.4% trypan blue solution. Viable cells (non-stained) were counted manually using a Neubauer hemocytometer under a light microscope, and total viable cell numbers were calculated as: cell number/mL = (average counted cells x dilution factor x 10⁴).

Cell culture: Seed the filtered cells at a density of 5 x 10³ cells/cm² in growth medium (low-glucose DMEM + 10% FBS + 1% penicillin/streptomycin) and incubate at 37 °C with 5% CO₂. Replace the medium every 2-3 days to remove non-adherent cells and maintain optimal culture conditions. All hADSCs used for subsequent experiments -- including cell characterization (surface marker detection, trilineage differentiation assays), DCEF stimulation, migration and morphology analysis, senescence/proliferation assays, and RNA sequencing -- were at passage 3 to passage 5.

hADSC characterization
Expression of cell surface markers: Culture cells to 70% confluency, then dissociate adherent cells with 0.25% trypsin-EDTA (incubate at 37 °C for 3 min). Centrifuge the cell suspension at 300 x g for 5 min at 4 °C, discard the supernatant, and resuspend the cell pellet in 1x PBS (volume: 1-2 mL). Aliquot 1 x 10⁵ cells per sample (resuspended in 100 µL of 1x PBS) and add fluorescently conjugated antibodies at the specified dilutions: anti-CD44 (1:50), anti-CD90 (1:100), anti-CD29 (1:50), anti-CD73 (1:50), and anti-CD105 (1:20)^1,4,7. Incubate the antibody-cell mixture at 4 °C for 30 min in the dark to avoid fluorophore quenching. After incubation, centrifuge at 1,000 x g for 5 min at 4 °C and aspirate the supernatant completely. Resuspend the pellet in 1-2 mL of 1x PBS, centrifuge again at 1,000 x g for 3 min at 4 °C, discard the supernatant, and repeat this washing process 2x to remove unbound antibodies. Finally, resuspend the cell pellet in 200 µL of fresh 1x PBS, acquire fluorescence data using a flow cytometer, and perform quantitative analysis with FlowJo software (v10). The gating workflow followed standard MSC phenotyping procedures: FSC-SSC gate to exclude debris and select the main cell population based on size and granularity, followed by doublet exclusion using FSC-A vs. FSC-H and SSC-A vs. SSC-H plots to ensure single-cell events. Then, live cell gating was done to exclude trypan blue-positive or PI-positive dead cells. Fluorescence gating was done for single-stained controls, and unstained controls were used to set thresholds for CD29, CD44, CD73, CD90, CD105 (positive markers), and CD34, CD45 (negative markers). Representative gating plots were exported and analyzed to confirm the identity of hADSCs.

Differentiation potential
Adipogenic differentiation: Harvest P3-generation hADSCs at 85% confluency using 0.25% trypsin-EDTA, then seed them in 24-well plates at a density of 1 x 10⁵ cells per well (using complete DMEM medium) and incubate at 37 °C with 5% CO₂ until cells reach 85% confluency again. For induction, use the adipogenesis differentiation kit, reconstituting kit component A (Adipogenic inducer) and component B (Adipogenic maintainer) as per the manual, and use component A for the first 3 days before switching to component B for 1 day. Repeat this cycle for 3 weeks. For staining, aspirate the induction medium, gently wash cells with 3 mL of 1x PBS (avoid disrupting cell layers), fix cells with 4% paraformaldehyde (PFA) at room temperature for 20 min, then incubate with 0.3% Oil Red O solution (prepared in 60% isopropanol) at room temperature for 15 min. Wash cells 3x with 1x PBS to remove excess stain, then image lipid droplets using an inverted light microscope (20x).

Osteogenic differentiation: Seed hADSCs in 12-well plates at a density of 1 x 10⁵ cells per well, and when cells reach 85% confluency, use the osteogenesis differentiation kit for induction -- reconstitute kit component A (Osteogenic inducer) and component B (Osteogenic supplement) in low-glucose DMEM (final concentrations: 10% FBS, 1% penicillin/streptomycin, 0.1 µM dexamethasone, 10 mM β-glycerophosphate, 0.05 mM ascorbic acid-2-phosphate) and refresh the medium every 3 days for 3 weeks. For staining, fix cells with 4% PFA at room temperature for 15 min, wash 3x with 1x PBS, then incubate with 2% Alizarin Red S solution (pH 4.2, prepared in deionized water) at room temperature for 30 min. Wash cells 3x with 1x PBS to remove unbound stain, then image calcium-deposited nodules under a phase-contrast microscope (20x) and quantify mineralization by measuring the area of Alizarin Red S-positive nodules using ImageJ software.

Chondrogenic differentiation: Centrifuge 250,000 hADSCs at 500 x g for 5 min at 4 °C in a 15 mL conical tube to form a cell pellet, add 500 µL of complete DMEM medium to the tube, and incubate overnight at 37 °C with 5% CO₂ to stabilize the pellet. The next day, use the chondrogenesis differentiation kit for induction -- reconstitute kit component A (Chondrogenic inducer) in CO₂-independent medium (final concentrations: 1% penicillin/streptomycin, 10 ng/mL TGF-β3, 50 µg/mL ascorbic acid-2-phosphate, 100 µg/mL sodium pyruvate) and gently refresh the medium every 3 days (avoid disturbing the pellet). After 3 weeks of induction, fix the cell pellet with 4% PFA at room temperature for 30 min, aspirate the fixative, incubate with 0.1% (w/v) Toluidine Blue O solution (prepared in 0.1 M sodium acetate buffer, pH 4.0) at room temperature for 15 min, aspirate the stain, rinse the pellet 3x with deionized water to remove excess dye, then image under a bright-field microscope (20x) and quantify sulfated proteoglycan accumulation by measuring the mean optical density (OD) of Toluidine Blue O staining using ImageJ software.

CAUTION: When handling 4% PFA (a toxic and corrosive reagent), work exclusively in a fume hood while wearing disposable gloves, a lab coat, and safety goggles to avoid inhalation of vapor or contact with skin and eyes -- if contact occurs, rinse immediately with running water for 15 min and seek medical attention. For hazardous waste disposal, collect used PFA and PFA-contaminated materials (e.g., pipette tips, plates) in a dedicated chemical waste container labeled PFA Waste and dispose of using institutional hazardous waste protocols, ensuring not to mix with biological waste.

Electrotaxis chamber assembly: Drill two holes (0.75 cm in diameter) at opposite ends of the centerline on a 10 cm Petri dish lid, positioned 1 cm from the dish edge (to accommodate agar-salt bridges). Apply a thin layer of vacuum grease (≤ 2 cm in length, ≤ 1 cm in width) on the bottom of the Petri dish along the centerline, 4 cm from each edge (Figure 1A). Using sterile fine-tipped forceps, place a sterile 1 cm x 2 cm glass coverslip on each grease patch, and press gently to ensure a tight seal (avoid coverslip breakage; Figure 1B-C). Apply another thin layer of vacuum grease on top of each attached coverslip, then place a second sterile 1 cm x 2 cm glass coverslip directly on the grease to form a double-coverslip stack. Press gently to ensure a tight seal between the two coverslips (prevent medium leakage; Figure 1D). Along the diagonal line connecting the top-left corner of one coverslip stack to the nearest dish edge, and the bottom-right corner of the opposite stack to its nearest edge, apply vacuum grease to form a continuous barrier wall. The wall should be tightly adhered to the dish bottom (no gaps) and measure approximately 2 cm in height and 0.5 cm in width (Figure 1E). Sterilize the assembled chamber under UV light for 20 min (to eliminate microbial contamination), then store at room temperature under sterile conditions until use (Figure 1F).

DCEF stimulation: Place sterile glass slides (for cell seeding) in a 10 cm Petri dish and incubate overnight at 37 °C with 5% CO₂ to pre-condition the slides (promote cell adhesion). Prepare Steinberg's solution by diluting 10 mL of 10x stock solution in 90 mL of ddH₂O (sterile; final concentrations: 58 mM NaCl, 0.67 mM KCl, 0.44 mM Ca(NO₃)₂4H₂O, 0.83 mM MgSO₄7H₂O, 10 mM HEPES; pH 7.4). Add 0.3 g agar to 20 mL of the prepared Steinberg's solution, microwave for 20 s until the agar is fully dissolved and the solution is clear, then immediately fill custom glass salt bridges with the hot agar-Steinberg's mixture and allow it to solidify at room temperature. After solidification, sterilize the salt bridges under UV light for 15 min, and sterilize two beakers (each containing 30 mL of Steinberg's solution) under UV light for 15 min. Seed P3-generation hADSCs on the pre-conditioned sterile slides at a density of 2 x 10⁴ cells/cm², incubate at 37 °C with 5% CO₂ for 24 h to allow cell adhesion, then transfer the cell-seeded slides to the runway of the assembled electrotaxis chamber and secure them with sterile side slides (to prevent movement). Seal the runway by placing a 3 cm x 2 cm glass coverslip over the vacuum grease, pressing gently to ensure a tight seal, and applying additional vacuum grease along the edges of the top coverslip to form a barrier (prevent medium evaporation). Fill the chamber's reservoirs with CO₂-independent medium (supplemented with 10% FBS and 1% penicillin/streptomycin; volume: 5-8 mL per reservoir), insert the sterilized agar-salt bridges through the holes in the Petri dish lid (with one end immersed in the chamber's medium reservoir and the other end immersed in the Steinberg's solution-filled beakers), and connect the beakers to a power supply through Ag/AgCl electrodes (to deliver a stable direct current). Apply a DC electric field (DCEF) of 2 V/cm for 4 h, monitor the voltage every 15 min using a digital multimeter (adjust if necessary to maintain constant field strength), and capture time-lapse images of cell migration every 5 min at 4 random fields using the software (5x objective, bright-field mode).

NOTE: When using electrical equipment, ensure the power supply and electrodes are dry and placed on a non-conductive surface to avoid electric shock. Wear insulated gloves when connecting or disconnecting electrodes, turn off the power before adjusting the setup, and in case of medium leakage onto electrical components, turn off the power immediately and wipe up with absorbent paper soaked in 75% ethanol. In this study, by minimizing the thickness of the electrophoresis chamber and maintaining it at approximately 1 mm, along with conducting multi-point voltage measurements at both ends of the chamber, the variation in field strength was controlled within 10%. Under these conditions, it is considered that the electric field distribution within the culture area is essentially uniform²⁶.

Migration and morphology analysis
Live imaging: For hADSCs under DCEF stimulation, import the time-lapse image sequences (5-min intervals) into ImageJ software, manually track 100 cells across 4 independent fields from the start (t=0) to the end of stimulation (t=4 h) using the Manual Tracking plugin, and calculate the mean migration velocity (µm/min) and directionality (D/T ratio, where D = straight-line distance from start to end, T = total path length) using the Chemotaxis Tool plugin (ImageJ).

Tracking: Calculate the following migration parameters to quantify electrotactic behavior: Directedness (Σcosθi/n, where θi is the angle between the cell's displacement vector and the direction of the electric field (EF), and n is the total number of tracked cells, ranging from -1 to 1 with values closer to 1 indicating stronger anode-directed migration), Accumulated Distance (total path length traveled by each cell during the stimulation period, in µm), Track Speed (accumulated distance divided by stimulation time, in µm/h), and Euclidean Distance (straight-line start-to-end displacement of each cell, in µm, reflecting net migration).

Morphometry: After DCEF stimulation, fix cells with 4% PFA at room temperature for 20 min, wash 3x with 1x PBS, stain the cytoskeleton with Phalloidin-TRITC (1:500 dilution in 1x PBS; volume: 200 µL per well for 24-well plates) at room temperature for 30 min (to label F-actin), and counterstain nuclei with DAPI (1 µg/mL in 1x PBS; volume: 100 µL per well) for 5 min. Cells were imaged using a fluorescence microscope at 20x magnification (with representative high-resolution images captured at 40x). Phalloidin-TRITC was visualized using an excitation wavelength of 540-555 nm and an emission wavelength of 565-580 nm, while DAPI was imaged using an excitation wavelength of 358-405 nm and an emission wavelength of 420-480 nm. Then measure the following morphological parameters in ImageJ: Long Axis (maximum Feret diameter, the longest distance between any two points on the cell perimeter), Vertical Length (width of the cell perpendicular to the long axis), and Verticality (sinα, where α is the angle between the cell's long axis and the direction of the EF, with higher values indicating greater alignment with the EF).

Senescence and proliferation assays
SA-β-Gal staining: Use a senescence detection kit to identify senescent cells (blue-stained cells) under a bright-field microscope (20x objective). Seed P3-generation hADSCs in 6-well plates at a density of 2 x 10⁵ cells per well, culture until 80% confluency, aspirate the medium, gently wash cells 3x with 1x PBS, add 1 mL of Fixative Solution (provided in the kit) per well and incubate at room temperature for 15 min, wash cells 3x with 1x PBS to remove residual fixative after fixation, add 0.5 mL of SA-β-gal Staining Working Solution (prepared by mixing 50 µL of SA-β-gal substrate with 450 µL of staining buffer per the kit manual) per well, seal the plate with transparent film (to prevent evaporation), incubate overnight (16-18 h) at 37 °C (avoid CO₂ exposure, as it alters pH and inhibits enzyme activity), capture bright-field images of ≥10 random fields per well (10x objective) the next day, count the number of blue-stained (SA-β-gal⁺) cells and total cells, and calculate the percentage of senescent cells as (Number of SA-β-gal⁺ cells / Total number of cells) x 100.

Ki67 immunostaining
After SA-β-gal staining, aspirate the staining solution, wash cells 2x with 1x PBS, permeabilize cell membranes with 0.1% Triton X-100 (prepared in 1x PBS; 1 mL per well) at room temperature for 10 min, and block non-specific antibody binding with 5% BSA (in 1x PBS; 1 mL per well) at room temperature for 1 h. Cells were then incubated with Anti-Ki67 primary antibody (1:200 dilution in 1% BSA/PBS; 500 µL per well) at 4 °C overnight. The next day, cells were washed 3x with 1x PBS (5 min each) remove unbound primary antibody and incubated with Alexa Fluor 488 488-conjugated secondary antibody (1:500 dilution in 1% BSA/PBS; 500 µL per well) at room temperature for 1 h in the dark. Cells were washed again 3x with 1x PBS and counterstained with DAPI (1 µg/mL in 1x PBS; 200 µL per well) for 5 min. Fluorescence images were captured using a fluorescence microscope at 20x magnification (with representative high-resolution images acquired at 40x). Alexa Fluor 488-labeled Ki67⁺ cells were visualized using an excitation wavelength of 485-495 nm and an emission wavelength of 515-545 nm, while DAPI-stained nuclei were imaged using an excitation wavelength of 358-405 nm and an emission wavelength of 420-480 nm. The percentage of proliferative cells was calculated as:(Number of Ki67⁺ cells / Number of DAPI⁺ nuclei) x 100.

NOTE: When handling primary and secondary antibodies, work in a BSC to prevent contamination, aliquot antibodies into single-use volumes to avoid repeated freeze-thaw cycles, and collect antibody-contaminated materials (e.g., pipette tips, plates) in a dedicated autoclavable Biological Waste bag for sterilization by autoclaving before disposal.

Age-dependent electrotaxis analysis of hADSCs: Use a DC pulse power supply to generate DCEF of three intensities: 0 mV/mm (control), 100 mV/mm, and 200 mV/mm, employ a live-cell workstation to observe and record cell migration in real time, and for each EF intensity, quantitatively compare the anodal migration capacity of hADSCs from young and elderly female donors by measuring Accumulated Distance, Euclidean Distance, Track Speed, and Directedness, with three independent biological replicates per group to ensure result reliability. Electrotactic migration parameters were quantified using the Manual Tracking and Chemotaxis Tool plugins in ImageJ (NIH). Individual cells were manually tracked for the entire 4-h stimulation period, and the following parameters were computed according to standard methods: Accumulated Distance: the total path length traveled by each cell during the recording period. Euclidean Distance: the straight-line distance between the cell's starting and ending positions. Track Speed: accumulated distance divided by total tracking time (µm/h). Directedness: calculated as the mean cosine of the angle (θ) between each displacement vector and the electric-field vector (values range from -1 to 1; values closer to 1 indicate strong anodal migration).

RNA sequencing
RNA extraction and preparation: Sterilize an ice-filled container under UV light for 15 min (to maintain RNA stability) and perform all subsequent steps on ice. Transfer hADSC-seeded glass slides (1 cm x 2 cm) from the electrotaxis chamber to a sterile Petri dish, wash cells 3x with 3 mL of ice-cold 1x PBS (add PBS gently, agitate slightly, and aspirate completely to remove residual medium), then add 1 mL of TRIzol reagent to each slide and incubate at room temperature for 20 min to fully lyse cells (ensure the lysate covers the entire cell layer). Transfer the lysate to an RNase-free 1.5-mL centrifuge tube, add 0.2 mL of chloroform, vortex vigorously for 15 s to induce phase separation, incubate the tube on ice for 15 min, then centrifuge at 12,000 x g for 15 min at 4 °C. This separates the lysate into three layers: upper aqueous phase containing RNA, middle protein layer, and lower organic phase. Carefully transfer the upper aqueous phase (≈ 0.5 mL) to a new RNase-free tube (avoid touching the middle layer), add 0.5 mL of isopropanol, vortex gently to precipitate RNA, incubate on ice for 10 min, then centrifuge at 12,000 x g for 10 min at 4 °C; RNA will form a white pellet at the bottom of the tube. Discard the supernatant, repeat the isopropanol precipitation once to improve RNA purity, aspirate the supernatant completely, air-dry the RNA pellet in a BSC at room temperature for 5-10 min (do not over-dry, as this reduces solubility), resuspend the RNA pellet in 30 µL of DEPC-treated water, incubate at 55 °C for 10 min to aid dissolution, quantify RNA purity by measuring the A260/A280 ratio (target: 1.8-2.0) using a spectrophotometer, assess RNA integrity using a RNA Nano Chip; target: RNA Integrity Number [RIN] > 8.0, and store qualified RNA samples at -80 °C until sequencing.

CAUTION: TRIzol and chloroform are toxic, volatile, and carcinogenic, so handle them exclusively in a fume hood while wearing nitrile gloves, a lab coat, and safety goggles -- avoid inhalation of vapors, wipe with a paper towel and rinse with soap and water for 10 min if spilled on skin, and do not mix it with bleach or other oxidizing agents (as this may produce toxic gases). For hazardous waste disposal related to RNA extraction, collect mixtures, isopropanol supernatants, and contaminated tubes in a sealed, chemically resistant container labeled RNA Waste (do not mix with aqueous waste or biological waste) and dispose of following institutional hazardous waste protocols, ensuring not to pour down drains.

Data preprocessing and quality control: Prepare sequencing libraries using the VAHTS mRNA-seq Library Prep Kit following the manufacturer's protocol -- enrich mRNA using oligo(dT) magnetic beads, fragment mRNA into 200-300 bp fragments using divalent cations, synthesize first-strand cDNA using random hexamers followed by second-strand cDNA, perform end repair, add A-tails, ligate sequencing adapters, amplify libraries by PCR, and purify using beads. Sequence the libraries on a sequencing platform (150 bp paired end reads), generating approximately 50 million raw reads per sample, then use a software to trim low-quality reads (remove reads with >50% bases having Phred quality score <20) and adapter sequences, retaining approximately 48 million clean reads per sample (Q30 > 90%) for downstream analysis after trimming.

Bioinformatics analysis: Align the clean reads to the human reference genome (GRCh38/hg38) using Hisat2 v2.2.1 software and save alignment results as BAM files, use htseq-count v0.13.5 software to count the number of reads mapped to each gene (based on Ensembl gene annotations), normalize gene count data using the estimateSizeFactors function from the DESeq (2012) R package to eliminate batch effects and differences in sequencing depth, and perform differential expression analysis using the nbinomTest function (DESeq package), selecting differentially expressed genes (DEGs) using thresholds of fold change (FC) > 2 and adjusted p-value (padj) < 0.05. Conduct Gene Ontology (GO) enrichment analysis (biological process, molecular function, cellular component) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis on DEGs using clusterProfiler v4.0 R package, focusing on enrichments related to cell migration (e.g., cell adhesion, cell migration), ion transport (e.g., sodium ion transmembrane transport), and signal pathways (e.g., PI3K-Akt signaling pathway). Perform unsupervised hierarchical clustering on DEGs using pheatmap v1.0.12 R package and visualize gene expression patterns across samples with a heatmap (row-scaled z-scores).

Statistical analysis: All experimental data are presented as Mean ± Standard Error of the Mean (Mean ± SEM), with at least 3 independent biological replicates per group (n ≥ 3). Use Student's t-test to compare differences between two groups (e.g., young vs. elderly donors) and one-way Analysis of Variance (ANOVA) followed by Tukey's post-hoc test to compare differences among three or more groups (e.g., different EF intensities), performing statistical analyses using SPSS 23.0 software and defining statistical significance as follows: *p < 0.05, **p < 0.01, ***p < 0.001.

Application of electric fields guides the directional migration of hADSCs and enhances their migratory capacity
The extracted hADSCs exhibited typical mesenchymal stem cell surface markers (e.g., CD29, CD44, CD90, CD73, CD105) and possessed trilineage differentiation potential (adipogenic, osteogenic, chondrogenic; Figure 2A), meeting the standard criteria for hADSC identification. We applied direct current electric fields (DCEFs) at intensities of 0 mV/mm, 100 mV/mm, and 200 mV/mm to investigate the electrotactic response of hADSCs. In the control group without electric stimulation (0 mV/mm), hADSCs migrated in random directions. However, under stimulation with an exogenous DCEF at 200 mV/mm, the cells migrated directionally toward the anode (Figure 2B). Furthermore, the number of hADSCs migrating toward the anode increased with higher electric field intensities (Figure 2C-D). By quantifying the angle between the cell displacement vector and the electric field vector, we observed a significant enhancement in anode-directed migration with increasing field strength (Figure 2E). Notably, although statistical tests did not fully confirm significant differences between all voltage groups (some groups p > 0.05), the trend in the data suggests that increased voltage may correlate with enhanced cell migration -- this observational trend requires further validation in future studies via expanded sample sizes or correlation analysis.

Application of electric fields induces morphological changes in hADSCs
To evaluate the morphological effects of electric field stimulation on hADSCs, we quantified cell morphology using three parameters (long axis length, vertical length, and verticality) following exogenous DCEF treatment (Figure 3A). After 4 h of DCEF stimulation, hADSCs exhibited an elongated morphology, with a significant increase in long axis length (Figure 3B-C). Additionally, both vertical length (width perpendicular to the long axis) and verticality (alignment of the cell's long axis with the electric field, quantified as sinα) were significantly increased with higher field intensity (Figure 3D-E), indicating that DCEF modulates hADSC morphology in an intensity-dependent manner.

Electrotactic behavior of hADSCs is associated with donor age
A previous study has reported that highly passaged murine ADSCs show reduced anode-directed migration under electric field stimulation. To further investigate whether donor age affects electrotaxis in hADSCs, we compared cells derived from young (27.00 ± 4.58 years) and aged (62.33 ± 4.04 years) female donors. We first confirmed that hADSCs from aged donors exhibited increased senescence (higher SA-β-gal+ cell ratio) and reduced proliferative activity (lower Ki67+ cell ratio) compared to those from younger donors (Figure 4A). Under DCEF stimulation, the number of aged hADSCs migrating toward the anode was significantly lower than that of young hADSCs (Figure 4B, Supplementary Figure 1, and Supplementary Figure 2). Moreover, aged hADSCs showed reduced accumulated distance (total path length), Euclidean distance (net straight-line displacement), track speed, and directedness (alignment with the electric field) compared to young hADSCs (Figure 4C-F), confirming that donor age negatively correlates with hADSC electrotactic capacity.

Aging may impair Na+ channel activity and thereby attenuate electrotactic migration capacity
To explore the potential molecular mechanisms underlying the age-related decline in electrotaxis, we performed transcriptomic analysis of hADSCs derived from aged and young female donors following 200 mV/mm DCEF stimulation. Significant differences in gene expression were observed between the two groups (Figure 5A), with 747 genes upregulated and 624 genes downregulated in aged hADSCs compared to young hADSCs (Figure 5B). Among these DEGs, key genes involved in sodium transport and channel function showed notable expression changes: the voltage-gated sodium channel gene SCN5A (log2 FC = -0.87, padj < 0.05) and the Na+ /H+ exchanger gene SLC9A1 (encoding NHE1, log2 FC = -1.13, padj < 0.05) were downregulated in aged hADSCs, while the monoatomic cation channel gene TRPM7 (log2 FC = 0.62, padj < 0.05) was upregulated. GO enrichment analysis of DEGs revealed that sodium ion transmembrane transport was significantly enriched in the biological process category, while monoatomic cation channel activity and voltage-gated sodium channel activity were enriched in the molecular function category; voltage-gated sodium channel complex was also enriched in the cellular component category-these results suggest that altered sodium channel activity could be involved in the impaired electrotaxis of aged hADSCs, though a direct causal role remains unconfirmed (Figure 5C). Gene Set Enrichment Analysis (GSEA) further supported this associative observation, showing upregulation of monoatomic cation channel activity but downregulation of sodium channel activity in aged hADSCs, indicating that overall sodium channel function might be altered with aging, rather than definitively compromised (Figure 5D). To determine whether the observed transcriptomic differences were attributable to intrinsic age-related variation or a diminished electric-field responsiveness, we analyzed an independent RNA-seq dataset (GSE269853) comprising unstimulated hADSCs from young and aged female donors. Gene Set Variation Analysis (GSVA) of key pathways-including PI3K-Akt signaling and sodium channel activity-revealed no statistically significant differences between the two age groups (Figure 5F-G). Both boxplot and heatmap analyses showed only subtle downward trends in these pathways in aged hADSCs, while core cellular functions such as ion transport and cytoskeletal organization remained largely comparable. These findings suggest that the basal expression levels of EF-responsive genes are largely maintained with aging, supporting the interpretation that the reduced transcriptional activation observed in aged hADSCs under electric field stimulation primarily reflects attenuated responsiveness rather than inherent transcriptomic disparity. Consistent with our previous research (which demonstrated that inhibiting NHE1 weakens PI3K-Akt pathway activation and electric field-induced directed migration through disrupted Na+ /H+ transport15), KEGG pathway analysis of DEGs also enriched the PI3K-Akt signaling pathway -- with core pathway genes PI3KCA (log2 FC = -0.92, padj < 0.05) and AKT1 (log2FC = -0.75, padj < 0.05) downregulated in aged hADSCs (Figure 5E). This observation implies a potential associative link between sodium ion dynamics and PI3K signaling in hADSC electrotaxis regulation, rather than a confirmed mechanistic connection.

Collectively, the results of this study demonstrate four key observations that support the study's objective of investigating age-dependent electrotaxis in hADSCs and its underlying mechanisms: (1) Exogenous DCEF induces anode-directed migration of hADSCs in an intensity-dependent manner, with higher field strengths enhancing directional migration; (2) DCEF also modulates hADSC morphology, promoting cell elongation and alignment with the electric field in an intensity-dependent fashion; (3) Donor age is a critical regulator of hADSC electrotaxis -- aged hADSCs exhibit increased senescence, reduced proliferation, and significantly impaired anode-directed migration under DCEF compared to young hADSCs; (4) Transcriptomic analysis identifies compromised sodium channel activity and dysregulated PI3K-Akt signaling as potential molecular drivers of age-related electrotaxis decline, linking ion transport and signal transduction to functional deficits in aged hADSCs. Together, these findings confirm the existence of age-dependent electrotaxis in hADSCs and reveal key molecular mechanisms, providing a foundation for optimizing hADSC-based therapies in regenerative medicine by accounting for donor age and targeting the sodium channel-PI3K-Akt signaling pathway.

Data availability:
All raw sequencing data generated in this study have been deposited in the NCBI Gene Expression Omnibus (GEO) under the accession number PRJNA1372472. Processed data and analysis files are available from the corresponding author upon reasonable request. All other data supporting the findings of this study are included in the article and its Supplementary Information files.

Figure 1: Fabrication and application of the electrotaxis chamber. (A) Assembling 1 cm x 2 cm glass coverslips onto the Petri dish bottom along its centerline using vacuum grease, spaced 1 cm apart. (B) Building vacuum grease barrier walls. (C) Placing the hADSCs-seeded coverslip, covering it with a 3 cm x 2 cm coverslip, and applying vacuum grease walls to both sides of the upper coverslip while maintaining an open channel beneath. (D) Adding prepared CO2 -independent medium and securing the drilled Petri dish lid. (E) Top view of the assembled chamber showing the coverslips. (F) Schematic representation of the applied electric field. Please click here to view a larger version of this figure.

Figure 2: Electric field stimulation directs hADSC migration and enhances their migratory capacity. (A) Flow cytometric characterization of hADSCs showing positive expression of CD29, CD44, CD90, and CD105, and negative expression of CD34 and CD45 (upper panel). Representative images of adipogenic (Oil Red O), osteogenic (Alizarin Red S), and chondrogenic (Toluidine Blue O) differentiation are shown (lower panel). Scale bar = 100 µm. (B) Representative live-cell images showing migration trajectories of individual hADSCs (colored lines) before (0 h) and after 4 h of stimulation with 200 mV/mm DCEF. Arrows indicate the final cell positions. Scale bar = 100 µm. (C) Representative migration trajectories of hADSCs under increasing DCEF intensities (0, 100, and 200 mV/mm). Red and black lines represent anodal and cathodal migration, respectively. (D) Quantification of the proportion of anodally and cathodally migrating cells under different DCEF intensities. (E-H) Quantitative analyses of directional migration parameters, including (E) directionality, (F) accumulated distance, (G) Euclidean distance, and (H) track speed under varying DCEF intensities. Data are shown as mean ± SEM (n = 3 donors; 100 cells per donor). *p < 0.05, **p < 0.01, ***p < 0.001; ns, not significant. Please click here to view a larger version of this figure.

Figure 3: Electric field stimulation induces morphological changes in hADSCs. (A) Schematic representation of morphological quantification parameters. The cellular morphology was characterized by the major (long) axis (red), the perpendicular (vertical) length (green), and the angle α, defined between the electric field (EF) vector and the cellular long axis. (B) Representative images showing morphological changes in hADSCs before (0 h) and after 4 h of stimulation with 200 mV/mm DCEF. The dashed white lines outline the cell contour, indicating elongation along both the long (red) and vertical (green) dimensions. Scale bar = 50 µm. (C-E) Quantitative analysis of morphological parameters under different DCEF intensities (0, 100, and 200 mV/mm): (C) cell long axis, (D) vertical length, and (E) verticality (sin α). Data are presented as mean ± SEM (n = 3 donors; ≥50 cells per condition). Statistical analysis was performed using one-way ANOVA with Tukey's post-hoc test (*p < 0.05, **p < 0.01, ***p < 0.001; ns, not significant). Please click here to view a larger version of this figure.

Figure 4: Age-dependent differences in migratory capacity of young versus elderly female-derived hADSCs under electric field stimulation. (A) Representative images showing senescence-associated β-galactosidase (SA-β-gal) staining and Ki67 immunofluorescence in hADSCs derived from young and elderly female donors. Quantification of SA-β-gal+ and Ki67+ cells indicate higher senescence and lower proliferative activity in aged hADSCs. Scale bars = 500 µm (SA-β-gal) and 100 µm (Ki67). (B) Quantitative comparison of the number of hADSCs migrating toward the anode under varying electric field strengths (0, 100, and 200 mV/mm). (C-F) Field strength-dependent migration parameters comparing young and aged hADSCs: (C) accumulated distance, (D) Euclidean distance, (E) track speed, and (F) directionality (cos θ). Data are presented as mean ± SEM (n = 3 donors per group; ≥ 80 cells analyzed per condition). Statistical significance was determined by two-way ANOVA followed by Tukey's post-hoc test (*p < 0.05, **p < 0.01, ***p < 0.001; ns = not significant). Please click here to view a larger version of this figure.

Figure 5: Differential gene expression analysis of hADSCs from aged and young female donors following 200 mV/mm DCEF stimulation. (A) Heatmap showing hierarchical clustering of differentially expressed genes (DEGs) between hADSCs from young and aged female donors after exposure to 200 mV/mm DCEF. Expression values are normalized and color-coded according to the Z-score. (B) Volcano plot of DEGs between young and aged hADSCs. Significantly upregulated genes (red) and downregulated genes (blue) were identified using the thresholds |log2FC| > 1 and p < 0.05. (C) Gene Ontology (GO) enrichment analysis of DEGs categorized into biological process (BP, left), molecular function (MF, middle), and cellular component (CC, right). The top enriched GO terms are ranked by significance (-log10(p) value) and gene count. (D) Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis highlighting the top 10 enriched signaling pathways, including calcium signaling, PI3K-Akt, and TGF-β pathways. Bar length indicates enrichment significance (-log10(p) value). (E) Gene set enrichment analysis (GSEA) of representative pathways showing enrichment in monoatomic anion/cation symporter activity (left) and sodium channel activity (right). NES: normalized enrichment score. (F) Boxplots showing GSVA enrichment scores of key pathways in unstimulated hADSCs derived from young (green) and aged (red) female donors. Pathways analyzed include cytoskeletal polarity, ion transport and potential, PI3K-Akt signaling, and sodium channel activity. No statistically significant differences were observed between the two age groups (ns, not significant; unpaired t-test). (G) GSVA enrichment heatmap illustrating sample-wise clustering of the same pathways. Both analyses revealed only mild downward trends in sodium channel activity and PI3K-Akt signaling in aged hADSCs, while overall pathway activity remained largely comparable across groups. Please click here to view a larger version of this figure.

Supplementary Figure 1: Migration trajectories of hADSCs from young female donors under varying DCEF intensities. Red and black trajectories indicate migration toward the anode and cathode, respectively. Axial scales are in micrometers (µm). Please click here to download this File.

Supplementary Figure 2: Migration trajectories of hADSCs from old female donors under varying DCEF intensities. Red and black trajectories indicate migration toward the anode and cathode, respectively. Axial scales are in micrometers (µm). Please click here to download this File.

Early in the tissue injury repair and regeneration process, electric fields emerge as one of the earliest physical cues established in the microenvironment. Their directional properties are spatiotemporally correlated with the recruitment of cells to the injury site²⁷. Endogenous electric fields originate from the transepithelial potential generated by polarized ion transport in epithelial tissues. When the epithelial barrier is disrupted or undergoes aging, the potential is secondarily altered, while undamaged or non-senescent regions maintain ion transport, resulting in a voltage gradient that generates a DCEF parallel to the epithelial surface²⁸. In this scenario, the center of injury acts as a cathode, serving as a key component of the electric fields. ADSC reside within a specific stem cell niche in adipose tissue²⁹. A growing body of evidence suggests that ADSC can be mobilized from their native niche and participate in skin repair and regeneration³⁰. In this study, we investigated whether ADSCs exhibit electrotaxis in response to an external DCEF, thereby assessing their responsiveness to endogenous EFs during skin regeneration. Our findings demonstrated that under DCEF stimulation, ADSC migrate directionally toward the anode, with their migration polarity positively correlated with EF strength. Given the critical role of ADSC in wound healing and tissue regeneration, several studies have explored their electrotactic behavior. For instance, murine ADSCs also migrate anodally within similar EF ranges³¹, consistent with our results. During the initial phase of wound healing, under the dominance of inflammatory signals, ADSC may preferentially respond to chemotactic cues and migrate toward the injury center³². However, in the subsequent repair phase, when EF signals intensify, the electrotactic response may suppress their retention at the cathodic wound core and redirect them to peripheral regions, contributing to angiogenesis and matrix remodeling. Notably, Hammerick³³ obtained contradictory results, suggesting that electrotactic heterogeneity may exist even among mesenchymal stem cells from the same species³⁴, highlighting the need for further cross-species validation of the effects of endogenous currents on ADSC.

We also observed that exposure to DCEF induced morphological changes in ADSC, including elongation of the cell long axis. Previous studies suggested that this is attributable to cytoskeletal rearrangement, where most actin filaments reorient parallel to the cell's long axis under EF stimulation^35,36. During skin regeneration, ADSC must traverse the dense collagen matrix in the dermis, and favorable morphological adaptation is believed to facilitate chemotaxis and migration³⁷. This implies that elongation of ADSC under endogenous EFs may enhance their dermal migratory capacity.

Electrotaxis is also critically regulated by the redistribution of membrane receptors. Electric fields can polarize membrane protein distribution, particularly for negatively charged proteins, which accumulate at the cathodal side due to the presence of positively charged hydration layers³⁸. This asymmetric redistribution leads to polarized activation of downstream signaling cascades²⁷. Epidermal growth factor receptor (EGFR) has been identified as a key factor in electrotaxis in various cell types, including corneal epithelial cells, keratinocytes, fibroblasts, breast cancer cells, and lung adenocarcinoma cells^{39,40,41,42,43}. In addition, other membrane receptors have also been implicated in electrotaxis. It has been shown that fibroblasts expressing high levels of integrins αMβ2, β1, α2, αIIbβ3, and α5 migrate cathodally under electric field stimulation, whereas those expressing integrins β3, α6, and α9 migrate anodally. Expression of integrins α4, αV, and α6β4 is associated with loss of directional migration⁴⁴. Early studies have also demonstrated that growth factors such as bFGF, VEGF, and TGF-β exhibit altered membrane distribution in response to electric fields⁴⁵. Asymmetric receptor activation typically implies directional bias in signal transduction⁴⁶. Our previous work showed that the PI3K pathway was involved in EF-guided epithelial cell migration¹⁵. Given that EF-induced changes in cells occur rapidly⁴⁷, ion channel asymmetry is also considered a key element in electrotaxis, with calcium, sodium, and potassium channels all implicated in different cell types⁴⁸. Although studies have been conducted on ADSC electrotaxis, species diversity remains limited. Hammerick³² found transient calcium influx in murine ADSCs under EF stimulation; however, calcium channel blockade did not significantly alter migration speed, whereas inhibition of PI3K or MAPK pathways reduced both migration velocity and directionality.

This study has notable limitations. The small donor sample size (3 young and 3 elderly females) restricts the generalizability of age-dependent electrotaxis findings, and the exclusive focus on female donors overlooks potential gender-related differences. This study only included female donor samples. The selection of this protocol was mainly to initially focus on age-related mechanisms in a single-gender context and reduce the interference of gender differences on the experimental results. However, this design also makes it difficult to directly generalize the research results to the male population, and the potential regulatory role of gender factors on the electromigration ability of hADSC cannot be ruled out. Future studies need to include male donor samples to verify the universality of the conclusion. Additionally, the RNA-seq analysis in this study only compared two groups: young hADSCs + 200 mV/mm EF and aged hADSCs + 200 mV/mm EF, without setting control groups of young and aged hADSCs under no EF stimulation. This experimental design makes it impossible to distinguish whether the observed differential gene expression (e.g., altered sodium channel and PI3K-Akt pathway-related genes) originates from inherent age-related differences in hADSCs (even without EF intervention) or from a specific reduction in the response of aged hADSCs to EF stimulation, thereby limiting the clarity of mechanistic inferences about age-dependent electrotaxis.

Alternative approaches could strengthen these findings. In vivo murine skin wound models with age-matched mice would connect in vitro observations to physiological wound healing. Electrophysiological techniques like patch-clamp recording could directly measure sodium channel activity in hADSCs under DCEF, while CRISPR-Cas9 knockout of key sodium channel or PI3K-Akt genes would confirm their causal role. 3D collagen hydrogel models could also better simulate dermal matrix stiffness, refining assessments of hADSC electrotaxis in more physiological contexts.

Future research should address these gaps. Expanding the donor cohort to include more samples, male donors, and individuals with diverse clinical backgrounds (e.g., obesity, diabetes) will improve the generalizability of the results. In vivo validation using aged animal models -- testing if modulating sodium channel or PI3K-Akt activity restores hADSC electrotaxis and wound healing -- is critical. Exploring electric field-assisted cell therapy (combining young hADSCs with DCEF) and investigating electrotaxis in other skin cells (e.g., fibroblasts) could inform therapeutic strategies, while single-cell RNA sequencing may reveal subpopulation-specific responses to electric fields in hADSCs.

Aging is associated with a decline in tissue repair and regeneration capacity, involving factors such as cellular senescence, stem cell exhaustion, inflammation, and mitochondrial dysfunction^49,50,51,52. Our data showed that aging reduces hADSC anodal migration directionality under DCEF (Figure 4), providing further evidence for age-related hADSC functional decline. Previous studies have linked reduced EGFR expression in aged rat epidermis to decreased PI3K signaling⁵³, and similar findings have been reported in human foreskin-derived skin progenitor cells, where PI3K pathway activity is critical for self-renewal and anti-aging functions⁵⁴. In the current study, our RNA-seq data indicated PI3K pathway downregulation in hADSCs from elderly female donors, alongside enriched differences in sodium channel-related genes (Figure 5) -- these observations of altered pathway transcription and weakened electrotaxis are correlated, though a direct causal link has not yet been established. It is important to note that aged hADSCs may also have other physiological and molecular defects (e.g., impaired mitochondrial function, altered overall metabolism, or epigenetic changes) that could contribute to reduced migratory capacity, and the current data cannot rule out the influence of these factors. While our early work demonstrated an association between NHE1-mediated Na+/H+ transport and PI3K phosphorylation in hADSC migration¹⁵, the current study only captured transcriptional changes -- we did not directly measure NHE1 protein expression, sodium channel activity, or PI3K protein phosphorylation, and transcriptional alterations do not necessarily reflect changes in protein function or activity. This limitation is partly tied to the lack of no EF control groups in the RNA-seq experiment; without comparing gene expression in young and aged hADSCs under non-stimulated conditions, we cannot fully disentangle inherent age-related transcriptional differences from EF-specific response changes, and thus the proposed connection between sodium channel/PI3K-Akt pathway changes and age-related electrotaxis decline remains to be validated with more direct evidence⁵⁵.