Intermediate Conductance Calcium‐Activated Potassium Channel Activation in Stem Cell Antigen‐1 Positive Cells Contributed to Vascular Restenosis

Abstract

Background

Vascular restenosis, a common complication following vascular reconstruction, results in stenosis or blockage that impairs vascular remodeling. This process is driven by the activation of vascular stem cells. Growing evidence suggests that intermediate conductance calcium‐activated potassium (IK_Ca) channels play a crucial role in regulating the function of these cells. This study aims to explore how IK_Ca channels influence Sca‐1⁺ (stem cell antigen‐1 positive) stem cells in the context of vascular anastomotic restenosis.

Methods

To investigate the role and mechanism of the IK_Ca channel in Sca‐1⁺ cell activation and its involvement in vascular anastomosis restenosis, we assessed the impact of IK_Ca channel deficiency on the vascular restenosis‐promoting ability of Sca‐1⁺ cells using a mouse femoral artery anastomosis model. Mechanistic insights were gained through patch‐clamp electrophysiology, intracellular Ca²⁺ measurement, and molecular biology techniques.

Results

Genetic deletion of IK_Ca channels in IK_Ca ^−/− mice led to reduced neointimal formation and decreased proliferation of Sca‐1⁺ cells at the anastomotic site. In vitro studies confirmed the presence of functional IK_Ca channels in Sca‐1⁺ cells and demonstrated that IK_Ca and TRPC1 (transient receptor potential canonical 1) channels cooperate in regulating membrane potential and intracellular Ca²⁺ levels. Furthermore, our findings suggest that IK_Ca channel‐mediated modulation of ERK (extracellular signal‐regulated kinase) and p38 phosphorylation underpins a key signaling mechanism in this process.

Conclusions

This study clarifies the role of the IK_Ca‐TRPC1‐Ca²⁺ pathway in activating vascular Sca‐1⁺ cells and establishes the contribution of the IK_Ca channel to vascular restenosis development. Understanding how IK_Ca channels affect the function of vascular Sca‐1⁺ cells provides valuable insights into the complex mechanisms of vascular remodeling during vascular stenosis.

Nonstandard Abbreviations and Acronyms

IK_Ca channel

intermediate conductance calcium‐activated potassium channel

MCP‐1

monocyte chemoattractant protein‐1

Sca‐1⁺

stem cell antigen‐1 positive

SMMHC

smooth muscle myosin heavy chain

TRPC1 channel

transient receptor potential canonical channel

VSMCs

vascular smooth muscle cells

Research Perspective.

What Is New?

• The crucial role of intermediate conductance calcium‐activated potassium (IK_Ca) channels in Sca‐1⁺ (stem cell antigen‐1 positive) cells lies in maintaining intracellular Ca²⁺ and membrane potential through activation of the IK_Ca–TRPC1 (transient receptor potential canonical 1)–Ca²⁺ cascade, promoting vascular restenosis via ERK (extracellular signal‐regulated kinase)/p38 MAPK (mitogen‐activated protein kinase) signaling that regulates their migration and proliferation.

• The genetic deletion of IK_Ca channels in IK_Ca ^−/− mice led to significant reductions in neointimal formation and decreased proliferation of Sca‐1⁺ cells at the anastomotic site, highlighting the potential regulatory role of IK_Ca channels in stem cell biology and providing a therapeutic target for improving patient outcomes in vascular stenosis.

What Question Should Be Addressed Next?

• Future research will specifically focus on determining whether there are other Ca²⁺ influx pathways besides TRPC1 and systematically explore downstream signaling pathways, including those involving Notch, Wnt, and YAP/TAZ (yes‐associated protein/transcriptional coactivator with PDZ‐binding motif) signaling mechanisms.

Vascular restenosis is a frequent complication of vascular injury, including arterial reconstructive surgery and transplantation, and can result in a range of symptoms and even limb ischemia, thereby presenting a significant risk to life. Medications, interventional therapy, and in severe cases, surgery can be used to clear blockages and enhance blood flow. However, effective management of vascular restenosis remains a challenge. Further research and advancements in medical technology are crucial for enhancing the outcomes of individuals affected by this condition. ¹ , ² , ³ Vascular restenosis, such as that occurring at vascular anastomoses, is a complex phenomenon initiated by vascular injury. This injury triggers significant structural alterations within the blood vessel, encompassing changes in vascular cell behavior, extracellular matrix modifications, and vascular architecture remodeling. ⁴ , ⁵ Key cellular players in this process include vascular smooth muscle cells (VSMCs), endothelial cells, fibroblasts, and vascular stem cells. ⁶ Exploring the complex molecular mechanisms and delving into the role of stem cell‐derived cells in vascular remodeling is crucial to enhance our understanding of this process.

Stem cells are widely recognized for their unique ability to renew and differentiate into various cell types. In a study conducted in 2004, ⁷ Hu et al. identified a group of cells in the aortic adventitia of ApoE ^−/− (apolipoprotein E) mice that displayed stem cell characteristics yet did not originate from the bone marrow. These cells were identified by the presence of specific markers such as Sca‐1 (stem cell antigen‐1), c‐kit, CD34, and Flk1. Further research has demonstrated that Sca‐1‐labeled (Ly‐6A/E) cells are distributed throughout different layers of blood vessels, including the adventitia, media, and intima, and may play an important role in the remodeling and adaptation of blood vessels. Although these cells typically remain in their respective locations, they possess the capability to become activated, migrate toward areas of vessel damage, and differentiate into VSMCs. Additionally, it has been observed that Sca‐1⁺ cells are capable of moving from the adventitia to the intima, thereby contributing to the regeneration of new tissue in venous grafts. ⁸ , ⁹ , ¹⁰ , ¹¹

Bioelectrical signals are fundamental components of living organisms and are primarily controlled by ion channels located on the cell membrane. These essential proteins manage the flow of ions, thereby regulating membrane potential, controlling excitability, shaping action potentials, triggering muscle contraction, and modulating cellular processes via Ca²⁺ signaling. ¹² Researches demonstrated that multiple ion channels collaborate to maintain optimal vascular function, and impaired ion channel function is closely linked to the development of vascular dysfunction and disease. ¹³ , ¹⁴ Despite this, the roles and mechanisms of various ion channels in stem cell bioelectrical signaling remain uncertain because of the limited understanding of stem cell cellular physiology and biophysics. ¹⁵ K⁺ channels expressed widely across cell types, are crucial in regulating cell migration and proliferation post injury. They modulate membrane potential; alter pH, volume, and shape; influence Ca²⁺ dynamics; activate growth factor receptors and downstream signaling; and interact with migration‐related proteins. ¹⁶ Among them, calcium‐activated potassium channels (K_Ca channels) are particularly important. Activated by intracellular Ca²⁺ ([Ca²⁺]_i), they help regulate membrane potential in both excitable and nonexcitable cells and modulate calcium signaling. K_Ca activation induces membrane hyperpolarization, enhancing Ca²⁺ influx through calcium‐permeable channels like nonselective cation channels. This sustained Ca²⁺ entry generates stable bioelectrical signals that influence cellular processes and gene transcription. ¹⁷ K_Ca channels are classified into 3 subtypes based on conductance: large conductance, intermediate conductance (IK_Ca), and small conductance. The IK_Ca channel, encoded by KCNN4, is a transmembrane protein expressed in various tissues including blood vessels. It regulates processes such as migration, tissue invasion, metastasis, and cell cycle progression. ¹⁸ IK_Ca channels are associated with various vascular diseases and findings indicate that targeting IK_Ca channels could open new possibilities for developing innovative approaches to prevent restenosis after angioplasty. ¹⁹ , ²⁰

Previous research has established the critical role of IK_Ca channels in various physiological functions and their potential dysregulation in vascular diseases. ²¹ Although IK_Ca activation on nonexcitable cells has been reported to regulate Ca²⁺ elevation, ²² , ²³ the specific Ca²⁺ entry mechanism regulated by IK_Ca channels remains unclear. TRPC channels (transient receptor potential canonical channels) are key players in cellular Ca²⁺ homeostasis and regulation, critically modulating numerous fundamental physiological processes. They mediate Ca²⁺ influx and play pivotal roles in vascular tone, vascular permeability, mechanosensation, secretion, angiogenesis, cell proliferation, and cell death, being closely associated with the pathogenesis of various vascular diseases. Specifically, activation of the TRPC1 subtype has been shown to control Ca²⁺ entry pathways. ²⁴ TRPC1 can cooperatively mediate Ca²⁺ influx with IK_Ca, thereby influencing fundamental processes such as cell proliferation, migration, and differentiation. ²⁵ However, the role of IK_Ca channels on Sca‐1⁺ cell in vascular remodeling—particularly in vascular stenosis—remains unexplored. Furthermore, despite their identification as K⁺ channels on stem cells, the precise functions and downstream signaling of IK_Ca within these cells, including whether IK_Ca couples Ca²⁺ signaling through TRPC1 channels, are unknown. To address these questions, we conducted experiments to test this hypothesis and to further investigate the role of IK_Ca using our established mouse model.

METHODS

The authors declare that all supporting data are available within the article (and its online supporting files). The unprocessed raw data sets (images) generated and analyzed during the current study are available from the corresponding author upon reasonable request.

Animals and Study Approval

Specific pathogen free C57BL/6 mice (C57 mice), aged 6 to 8 weeks and weighing 20 to 22 g, were used in the experiments. C57 mice were obtained from Chengdu Dashuo Experimental Animal Co., Ltd. (license number: SCXK [Chuan] 2015‐030). Specific pathogen free transgenic mice (Sca‐1 CreER, Sca‐1 CreER/Rosa26‐LSL‐tdTomato (tandem dimer Tomato), and KCNN4‐flox/Sca‐1 CreER/Rosa‐LSL‐tdTomato mice), with an average age of 3 to 6 months and body weight of ∼30 g, were obtained from Shanghai Southern Model Biotechnology Co., Ltd. (License number: SCXK [Shanghai] 2019‐0002). These mice were housed and maintained in the Laboratory Animal Center of Southwest Medical University (license number: SYXK [Sichuan] 2023‐0065). For the preparation of transgenic animals, please refer to Figure S1A. The mice were housed in a room with controlled light and humidity conditions: temperature maintained at 21 °C±2 °C, humidity at 50%±10%, and a regulated light/dark cycle of 12 hours each (lights on from 8 pm to 8 am). All animal experiments strictly adhered to the guidelines for the welfare and use of experimental animals as prescribed by the regulation for experimental animals in China and approved by the Ethics Committee of Southwest Medical University (protocol 201 903‐190). To determine the sample size for the animal experiments, a pilot study was conducted using hematoxylin and eosin staining and immunofluorescence tracing. This preliminary investigation successfully confirmed the involvement of Sca‐1⁺ cells in vascular remodeling, thereby informing the sample size estimation. In accordance with ethical principles, we computed the effect size (Hedges’ g) based on existing data, while also incorporating sample size and significance level (α=0.05) to evaluate statistical power. The number of animals required was determined to ensure that the experiment could detect significant changes in primary outcomes with reliability, thereby upholding both scientific rigor and ethical standards. The work has been reported in accordance with the Animals in Research: Reporting In Vivo Experiments guidelines. ²⁶ All mice were anesthetized using a nose cone with 1.1% isoflurane at a flow rate of 1.0 L/min, delivered through a specialized evaporator. They underwent modeling surgery and were euthanized under deep anesthesia using 3.0% isoflurane at a flow rate of 1.5 L/min, by cardiac perfusion of phosphate‐buffered saline at the designated time to obtain tissue samples for additional analysis.

Preparation of Mouse Femoral Arterial Anastomosis Injury Model

The preparation of the mouse femoral artery anastomosis model and tamoxifen treatment strategy are outlined in Figure S1B. The mice were anesthetized using a dedicated isoflurane evaporator. After the mice were fully anesthetized, they were positioned supine and their leg skin was incised longitudinally. Using microforceps, the femoral artery was carefully separated from the femoral nerve and vein. To aid in the subsequent procedures, a small spacer was gently inserted beneath the isolated femoral artery. The proximal and distal ends of the artery were secured with 2 arterial clips (distance between the clamps was 1 cm) before transection using microscissors. Additional drops of heparin saline solution were applied at the surgical site until no visible blood obstruction was observed. To support the vessel, we insert a 1‐cm long 6‐0 surgical suture into the femoral artery. In the lumen just to provide support for the vessel during subsequent suturing and simulate the potential damage caused by a guidewire intervention. The vascular incisions were carefully connected using 12‐0 sutures (Ningbo Medical Suture Needle Co., Ltd), with 4 needles (Medical Device Registration Certificate No. 20192020463) evenly spaced apart. Heparin saline solution was added during the procedure if needed. Toward the end of the suturing process, the nylon thread inside the vessel was removed. Once suturing was complete, the small spacer beneath the vessel was removed, and the arterial clips at both ends were released to observe the restoration of blood flow. Only when the blood flow was fully restored and no bleeding occurred, the skin of the mice was stitched using 6‐0 sutures. Additionally, the skin at the suture site was disinfected with an appropriate amount of iodophor. Five weeks post surgery, the targeted section of the femoral artery was carefully severed and transferred into a centrifuge tube that was prefilled with OCT. The sample was then frozen in liquid nitrogen and stored at −80 °C until further analysis. Subsequently, the frozen sample was carefully sliced in sequence, spanning the entire length of the vessel segment exhibiting hyperplasia. Each slice was 10 μm thick. For further analysis, 3 to 5 sections from each animal were chosen from the front, center, and back portions of the hyperplasia segment, at intervals of 100 to 200 μm. The average data per mouse were calculated and then used to calculate the average per group. In the sham‐operated group, a sham procedure was performed where ∼1 cm of the femoral artery was isolated and exposed without being cut. The vessel remained in this state for ∼10 minutes—a duration equivalent to the time required for anastomosis in the model group. The surgery was otherwise identical in all other respects to that performed in the model group. We maintained strict control over both the experimental procedures and the sampling locations. In the model group, a standardized incision of approximately 1 cm was made at the femoral artery site, followed by placement of a 6‐0 nylon suture. All vascular incisions were consistently performed at this predefined location. Similarly, in the sham operation group, the surgical sites were carefully controlled to correspond to analogous anatomical regions. All the animal models of vascular anastomosis were performed by one individual, fluorescent staining and scanning were performed by another, and the data were analyzed by a different individual. All the experiments were blind.

Methods for Primary Isolation and Culture of Sca‐1⁺ Cells

Sca‐1⁺ cells were isolated in a similar manner to that previously described. ²⁷ Sca‐1⁺ cells were isolated from the adventitia of the thoracic aorta of C57Bl6/J mice. Primary cultured adventitia cells were screened using magnetic beads (Anti‐Sca‐1‐VioBright FITC and Anti‐FITC MicroBead) to obtain purified Sca‐1⁺ cells. These cells were then cultured in American Type Culture Collection DMEM supplemented with 10% FBS (Millipore, Burlington, MA, USA), 1% penicillin–streptomycin, and 0.02% leukemia inhibitory factor in a humidified incubator at 37 °C and 5% CO₂. To maintain purity, the cells were sorted using magnetic beads every 5 generations. All the stem cells used in the experiments were within 15 generations. In this study, isolated and cultured Sca‐1⁺ cells were treated with the differentiation factor TGF‐β1 (transforming growth factor alpha; 2 ng/mL) for 3 days. mRNA levels of SM22 (smooth muscle 22) and calponin were measured to confirm the differentiation potential of Sca‐1⁺ cells, thereby verifying their fundamental stem cell characteristic—the ability to differentiate into smooth muscle lineage cells.

Electrophysiology

Whole‐cell perforated patch clamp technology is usually used to record current unless otherwise specified using an EPC‐10 amplifier and Patchmaster software (Heka Elektronik, Lambrecht, Germany). For the total current recording, the bath solution consisted of (in mM): NaCl 137, KCl 5.9, MgCl₂ 1.2, glucose 14, HEPES 10, and CaCl₂ 1.8 (NaOH adjust pH 7.4); the pipette solution consisted of (in mM): D‐aspartic acid 110; KOH 110, KCl 30, NaCl 10, HEPES 10, MgCl₂ 1, EGTA 0.05 (KOH adjust pH 7.2), and 300 μg/mL nystatin. For K⁺ current recording, the bath solution and pipette solution consisted of (in mM): KMeSO₃140, KCl 2, MgCl₂ 2, HEPES 20, EGTA 1 (KOH adjusted to pH 7.2), and 300 μg/mL nystatin. The membrane potential was generally held at −30 mV for the total current recording and 0 mV for the K⁺ current recordings. Either a ramp pulse at 0.45 V/s from −100 to +80 mV or a set of depolarizing step pulses (400 ms duration and in 10 mV‐increments at 1‐second intervals) was applied. Currents were low‐pass filtered at 1 KHz, digitized at 10 KHz, and analyzed using the pCLAMP software (version 10.0; Molecular Devices). The series resistance and total cell capacitance were calculated from the uncompensated capacitive transients in response to 10 ms hyperpolarizing step pulses (5 mV). The series resistance in the perforated patch recordings was <30 MΩ.

In this study, one method to observe the effect of Ca²⁺ on the current was to use ATP to induce intracellular Ca²⁺ transients and to observe the current increase caused by instantaneous [Ca²⁺]_i increase. Another method to observe the effect of intracellular Ca²⁺ on the current is the use of calcium ionophores, ionomycin. Total currents or K⁺ currents were measured using the whole‐cell perforated patch configuration. Extracellular 100 μM ATP, 1 μM ionomycin, or 1‐[(2‐chlorophenyl) diphenylmethyl]‐1h‐pyrazole (TRAM34) was applied and the changes in current were measured.

The effect of intracellular Ca²⁺ on the current was also observed using the whole‐cell perforated patch‐clamp technique with Ca²⁺ in the pipette solution. Because Ca²⁺ in the pipette cannot enter the intracellular fluid after perforating the membrane with nastytin, the current at the cellular base Ca²⁺ concentration can be recorded at this time. The membrane was then ruptured by negative‐pressure suction to form a standard whole‐cell configuration. At this time, Ca²⁺ in the pipette solution was buffered into the cells. Under higher [Ca²⁺]_i (~500 nM), the change in membrane current was recorded, thus reflecting the increase in ion channel current caused by intracellular Ca²⁺. The higher [Ca²⁺]_i (~500 nM) pipette solution consisted of (in mM): KMeSO₃140, KCl 2, MgCl₂ 2, HEPES 20, EGTA 1, and CaCl₂ 0.81 (KOH adjusted pH 7.2)and 300 μg/mL nystatin.

In our experimental investigation of cell membrane potential, we used the patch‐clamp current‐clamp mode. First, we measured the resting membrane potential of the cells. Furthermore, we examined hyperpolarization of the membrane potential induced by activated K_Ca, which was triggered by an elevation in [Ca²⁺]_i induced by ionomycin. Changes in membrane potential were measured after blocking the IK_Ca channel with TRAM34 or apamin (a type of small conductance specific blocker). Membrane potential experiments were conducted using the total current‐recording solution.

Methods for Sca‐1⁺ Cell Migration Assays

Transwell migration assays were performed to evaluate the migration of Sca‐1⁺ cells. In the experiment, the upper chambers of the transwell plates were seeded with cells at a density of (2–3)×10⁵ cells/mL, whereas the lower chambers were filled with DMEM supplemented with 10% FBS. The cells were then incubated in a humidified 5% CO₂ atmosphere at 37 °C. Various treatments were applied, including MCP‐1 (monocyte chemoattractant protein‐1) or ATP, the IK_Ca channel antagonist TRAM34, TRPC1 channel blocker Pico154, and adenovirus short hairpin RNA (shRNA) interference targeting KCNN4. In the ATP‐containing group, a final concentration of 30 μM ATP was added to the upper chamber and 500 μL of complete medium was added to each well in the lower chamber. The MCP‐1 group was added to the lower chamber at a final concentration of 3 ng/mL MCP‐1. After 24 hours, migrated cells were stained and observed under a bright‐field microscope at 100× magnification.

Methods for Sca‐1⁺ Cell Proliferation Assays

The CCK‐8 assay was used to measure the viability of Sca‐1⁺ cells. Initially, cells were seeded at a density of 5×10³ cells/well in 96‐well plates. Each well contained 100 μL of complete medium, and the plates were placed in a 37 °C incubator with a 5% CO₂ atmosphere for overnight incubation. Following the adherent culture, the medium was replaced with serum‐free medium supplemented with 10 ng/mL PDGF‐BB (platelet‐derived growth factor‐BB), 10 μM ATP, TRAM34, orPico145. After a 24‐hour incubation period with the drugs, 10 μL of CCK8 reagent was added to each well and incubated for 3 hours. Subsequently, optical density at 450 nm was measured to determine the number of viable cells.

Intracellular Calcium Measurement

The intracellular Ca²⁺ transient (free Ca²⁺ concentration ([Ca²⁺]_i) or F340/F380) was measured in cultured mouse Sca‐1⁺ cells using a TILLvisION 4.0 imaging system (Till Photonics, Gräfelfing, Germany) at room temperature (22 °C–25 °C). ²⁷ Sca‐1⁺ cells were loaded with 5 μM fura‐2/AM for 30 minutes, and [Ca²⁺]_i was estimated using the fluorescence emission ratio at 510 nm from 340 nm (15 ms) and 380 nm (10 ms) excitation. The equation used to estimate [Ca²⁺]i was [Ca²⁺]_i=Kd×(Sf2/Sb2)×(R−Rmin)/(Rmax − R), where Kd is the dissociation constant for fura‐2/calcium (224 nM), R is the ratio of the fluorescence emission evoked by 340‐ and 380‐nm light excitation, and Sf2/Sb2 is the ratio of fluorescence emission evoked by 380‐nm excitation in Ca²⁺‐free Tyrode’s solution and saturating [Ca²⁺] solution. Tyrode’s solution used in this experiment contained the following components in mM concentrations: NaCl 127, KCl 5.9, CaCl₂ 2.4, MgCl₂ 1.2, glucose 12, and HEPES 10. The pH of the solution was adjusted to 7.4 with NaOH. Transient changes in [Ca²⁺]_i were induced by applying ATP (30 and 100 μM) or ionomycin (1 μM) in Tyrode’s solution with 2.4‐mM Ca²⁺.

Western Blotting

Total membrane and phosphorylated proteins were extracted from Sca‐1⁺ cells and quantified using the BCA method. The samples were separated by sodium dodecyl sulfate‐polyacrylamide gel electrophoresis (SDS‐PAGE) and transferred to nitrocellulose membranes. The membranes were incubated with the indicated primary antibodies overnight at 4 °C and then with a horseradish peroxidase‐conjugated secondary antibody (1:2000). The antibodies used included anti‐KCNN4 (1:200, Cat#:ab215990, Alomone), phospho‐p44/42 MAPK (p‐Erk1/2) (Thr202/Tyr204) (D13.14.4E) XP Rabbit mAb #4370 (1:1000, Cell Signaling Technology), p44/42 MAPK (Erk1/2) (137F5) rabbit mAb (1:1000, Cell Signaling Technology), p38 MAPK Antibody #9212, and Phospho‐p38 MAPK (Thr180/Tyr182) (D3F9) XP Rabbit mAb #4511 (Cell Signaling Technology 1:1000). The target protein bands were visualized using an enhanced chemiluminescence substrate and a gel imaging system. Expression levels were quantified relative to caveolin or glyceraldehyde‐3‐phosphate dehydrogenase, which served as gel loading controls.

Immunofluorescence

Sca‐1⁺ cells were gently washed with PBS and fixed with 4% PFA on sterile glass slides. They were then blocked with BSA and incubated overnight with an anti‐KCNN4 receptor (extracellular) antibody (1:200, Alomone, Jerusalem, Israel, #APR‐102). The following day, the cells were treated with a secondary antibody (donkey anti‐rabbit IgG (H+L) highly cross‐adsorbed or conjugated to Alexa Fluor 488, 1:500, Thermo Fisher). To prevent fluorescence quenching and stain nuclei, a HelixGen antifluorescence quenching agent containing DAPI was added dropwise. The images were captured using a confocal microscope (Zeiss‐LSM‐980, Germany).

Hematoxylin and Eosin Staining

Frozen tissue sections were prepared with a thickness of 10 μm using a cryostat. The cells were then equilibrated to room temperature and fixed with 4% neutral paraformaldehyde for 20 to 30 minutes. Subsequently, they were washed with running water for 30 seconds. The sections were stained with hematoxylin for 12 to 15 minutes and washed again with running water. To differentiate the sections, they were treated with 1% hydrochloric acid‐ethanol for 5 seconds and soaked in tap water for 10 minutes. Eosin stain was then added for 30 to 40 seconds, followed by another wash with running water. A gradient decoloration process was performed to further decolorize the sections. The sections were soaked in 75%, 85%, 95%, and absolute ethanol for 2 to 5 seconds each. The tissue sections were then made transparent by immersing in xylene for 1 to 2 seconds, dried, and sealed under coverslips with 1 to 2 drops of neutral resin. As a result of the staining process, the cytoplasm, collagen fibers, and muscle fibers were distinguished by different shades of red, whereas the nuclei were distinguished by blue staining.

Statistical Analysis

Statistical analyses were performed in R (v4.4.3), using the coin package for permutation tests (10 000 permutations) and sandwich/lmtest for heteroskedasticity‐consistent robust SE (HC3) robust inference in linear models. Observations were obtained from independent experimental units. Continuous variables are summarized as mean±SD. For 1‐factor experiments, 2‐group comparisons were analyzed using permutation t tests and comparisons involving >2 groups were analyzed using permutation 1‐way ANOVA (10 000 permutations). For 2‐factor experiments, outcomes were analyzed using linear models with HC3 robust inference. Models were initially fit including main effects and an interaction term; when the interaction term was not supported, results were summarized using an additive main‐effects model (excluding the interaction). When the interaction term was supported, conditional comparisons (simple effects) were obtained using estimated marginal means. Effect estimates with 95% CIs are reported alongside P values. For migration assays, measurements were baseline normalized within each biological replicate to the paired internal control condition (control set to 100%), and analyses were performed on the normalized values. The specific analysis methods employed for each group of data are described in the results section. Transgenic mice expressing KCNN4‐flox×Sca‐1‐CreERT×Rosa26‐tdTomato were used in this study to examine the impact of IK_Ca channels on neointima formation at vascular anastomosis. This effect served as the primary experimental outcome of the experiment. By analyzing the available data, the effect size (Hedges’ g) was determined and considered in conjunction with the sample size and significance level (α=0.05) to evaluate the statistical test’s power. The specific number of animals and tissue sections used for each experiment are detailed in the appropriate legend, and each experimental cohort consisted of an equal number of males and females.

RESULTS

IK_Ca on Sca‐1⁺ Cell Contribute to Regeneration and Remodeling Following Femoral Arterial Anastomosis

In a study on femoral artery anastomosis, C57BL/6 mice with Sca‐1⁺ cells expressing tdTomato were used to create a model. These mice, referred to as IK_Ca ^+/+ mice, were used as controls. Additionally, a femoral artery anastomosis model was established in KCNN4 knockout mice (Kcnn4‐flox×Sca‐1‐CreERT×tdTomato), which permitted the lineage tracing of Sca‐1⁺ cells. These mice were labeled as IK_Ca ^−/−mice. In addition to using the tail‐polymerase chain reaction ‐based identification technique to detect transgenic mice, Figure S1C provides evidence of the knockout of a specific IK_Ca protein in Sca‐1⁺ cells. Hematoxylin and eosin staining conducted 5 weeks post injury revealed noteworthy alterations such as intimal hyperplasia, thickening, and lesions (Figure 1A and 1B). Compared with the sham group, the intimal area dramatically increased from 3554±902 μm² (based on data obtained from 30 sections of hyperplasia segments from 10 control mice; 3 sections were selected from the front, center, and back portions of each animal) to 28 289±6882 μm² (based on data obtained from 27 sections of hyperplasia segments from 9 anastomosis group mice; 3 sections were selected from the front, center, and back portions of each animal) (P=0.0000). Additionally, the ratio of intimal area to medial area increased from 0.28±0.04 to 1.96±0.50 (P=0.0000). Furthermore, the proportion of Sca‐1⁺ cells, indicated by red tdTomato fluorescence, among the total cells around the anastomosis, demonstrated a significant increase (P=0.0000) from 9.50%±2.33% (based on 24 sections from 8 control mice) to 24.22%±6.49% (18 sections from 6 mouse anastomosis studies were examined). One section was extracted from the front, center, and back portions of the hyperplastic segment in each animal. A section of the anastomosis site revealed a notable increase in red‐tdTomato‐Sca‐1⁺ cells surrounding the adventitial tissue of blood vessels. These cells exhibited migration and proliferation into the intimal hyperplasia (Figure 1C).

Figure 1. The formation of neointima occurred following the vascular anastomosis.

A, Neointimal hyperplasia was observed following anastomosis injury in mice. B, Statistically significant intimal hyperplasia was found in the anastomosis group (P=0.0000, permutation t test, control: 30 sections/10 mice; anastomosis: 27 sections/9 mice). C, The proportion of Sca‐1⁺ cells (tdTomato) significantly increased after anastomosis (P=0.0000, permutation t test, control: 24 sections/8 mice; anastomosis: 18 sections/6 mice). α‐SMA indicates α‐smooth muscle actin; CON, control; Sca‐1⁺, stem cell antigen‐1 positive; and tdTomato, tandem dimer Tomato.

We found that the absence of IK_Ca contributed to a decrease in intimal hyperplasia after vascular anastomosis (Figure S2A). When compared with the control group (IK_Ca ^+/+ mice), the group of IK_Ca ^−/− mice displayed a significantly lower ratio of neointima area to media area (1.34±0.18 versus 1.96±0.50, P=0.0012. n=24 sections from 8 IK_Ca ^−/− mice and n=27 sections from 9 IK_Ca ^+/+ mice. Three sections were selected from the front, center, and back portions of the hyperplastic segment in each animal) (Figure S2A). The power of the statistical test and the estimated Hedges’ g effect size are shown in the legend of Figure S2. The proportion of Sca‐1⁺ cells labeled with tdTomato surrounding the neointima and perivascular tissues significantly decreased when the IK_Ca channel in Sca‐1⁺ cells was knocked out (Figure 2A; 16.8%±2.8% versus 24.2%±6.5%, P=0.0100. n=24 sections from 8 IK_Ca ^−/− mice and n=18 sections from 6 IK_Ca ^+/+ mice. Three sections were selected from the front, center, and back portions of the hyperplastic segment in each animal). Additionally, we analyzed the differentiation of Sca‐1⁺ cells into α‐SMA (α‐smooth muscle actin) and SMMHC (smooth muscle myosin heavy chain)‐labeled cells in the neointima. We calculated the proportion of α‐SMA‐labeled cells originating from Sca‐1⁺ cells and that of SMMHC‐labeled cells (mature SMCs) originating from Sca‐1⁺ cells. The percentage of mature SMCs labeled with SMMHC derived from Sca‐1⁺ cells was significantly lower in the IK_Ca ^−/− group than that in the IK_Ca ^+/+ group (70.00%±4.18% versus 52.83%±10.15%, P=0.0064). Conversely, there was no notable difference in the proportion of cells labeled with α‐SMA derived from Sca‐1⁺ cells between the IK_Ca ^+/+ group and IK_Ca ^−/− group (47.67%±14.99% versus 44.40%±5.32%, P=0.6691). Data were collected from 6 IK_Ca ^−/− animals and 5 IK_Ca ^+/+ animals (Figure 2B; Figure S2B). Our findings further support that the knockout of IK_Ca has no effect on the involvement of Sca‐1⁺ cells in endothelial repair. We observed that the number of Sca‐1⁺ cells differentiating into CD31⁺ cells during the endothelial repair process remained unaffected despite the IK_Ca knockout in Sca‐1⁺ cells (Figure S2C; sections from 6 IK_Ca ^−/− mice and 5 IK_Ca ^+/+ mice, P=0.6451).

Figure 2. IK_Ca knockout in Sca‐1⁺ cells inhibits vascular remodeling and SMC differentiation.

A, The graphic shows Sca‐1⁺ cells per total cells in IK_Ca ^+/+ and IK_Ca ^−/− with representative pictures (P=0.0100, permutation t test, n=24 sections from 8 IK_Ca ^−/− mice and n=18 sections from 6 IK_Ca ^+/+ mice). B, IK_Ca channel regulates SMC differentiation. tdTomato⁺/α‐SMA⁺ and tdTomato⁺/SMMHC⁺ cells were visualized (a, b). The ratios of tdTomato⁺/α‐SMA⁺ and tdTomato+/SMMHC+ cells were assessed. The ratio of tdTomato copositive cells was significantly lower within the SMMHC⁺ population (P=0.0064, permutation t test, from 15 sections from 6 IK_Ca ^−/− animals and 13 sections from 5 IK_Ca ^+/+ animals) (c). The images in (A) and (B) demonstrated tdTomato labeled Sca‐1+ cells in red, Hoechst33342‐stained nucleus in blue, α‐SMA (a smooth muscle marker) in green, and SMMHC (a mature SMC marker) in magenta. Statistical comparisons were based on number of mice examined. α‐SMA indicates α‐smooth muscle actin; CON, control; IK_Ca, intermediate conductance calcium‐activated potassium channel; Sca‐1⁺, stem cell antigen‐1 positive; SMC, smooth muscle cell; SMMHC, smooth muscle myosin heavy chain; and tdTomato, tandem dimer Tomato.

All in vivo experiments demonstrated that downregulation or impairment of IK_Ca channels in Sca‐1⁺ cells significantly reduced neointimal formation following vascular anastomosis. Thus, IK_Ca channels are critical for functional restoration postoperatively.

Expression of IK_Ca Channel on Sca‐1⁺ Cells

Adventitial cells harvested from thoracic aortas of C57 mice were cultured in flasks. Figure S3A shows the cell growth in culture. Immunomagnetic beads were used to screen cultured cells selectively, as illustrated in Figure S3B. To confirm the identity of the screened cells, immunofluorescence with a Sca‐1 antibody was performed; the results are shown in Figure S2B. Additionally, flow cytometry results, presented in Figure S3C and S3D, revealed that 94.72% of the cells were Sca‐1⁺ cells. Immunofluorescence experiments on cells were conducted to identify the fluorescence of Sca‐1 and IK_Ca channel markers in cultured cells, as shown in Figure S3E. After intraperitoneal injection of tamoxifen into Sca‐1 CreER/Rosa26‐LSL‐tdTomato, and KCNN4‐flox/Sca‐1 CreER/Rosa‐LSL‐tdToamto mice, adventitias of the aorta were extracted for vascular en face analysis. This analysis revealed the presence of IK_Ca channels (marked in green) in adventitia Sca‐1⁺ cells (tdTomato‐marked red cells) in their natural environment (Figure S1C). In experiments involving the differentiation of Sca‐1⁺ cells induced by TGFβ‐1, a significant increase in the mRNA levels of smooth muscle cell markers SM22α and calponin was observed after 3 days of stimulation. Additionally, polymerase chain reaction analysis showed an increase in IK_Ca channel mRNA and SMMHC protein levels following differentiation of Sca‐1⁺ cells into SMCs. However, no changes in IK_Ca channel protein were detected (Figure S4).

We determined the type of ion channel current in Sca‐1⁺ cells. The total current of the Sca‐1⁺ cells was recorded under physiological conditions, and 3 common types of currents were observed (see the typical recordings in Figure S5A‐a through S5A‐f). Statistical analysis of the currents from different cell generations revealed minimal changes in the total current (Figure S5B). Owing to the complex composition of the current system, further identification is not currently possible. To simplify the identification process, we implemented specialized methods to control the pipette and bath solutions, specifically targeting the investigation of the K⁺ current in Sca‐1⁺ cells. To achieve this, we replaced Cl⁻ in both extracellular and intracellular solutions with sulfonate ions (MeSO₃ ⁻). In addition, the K⁺ current of Sca‐1⁺ cells was recorded using 140 mM K⁺ concentration in both the inner and outer cell solutions. Two commonly observed types of K⁺ currents were recorded, as illustrated in Figure S5C‐a through S5C‐d. Notably, Sca‐1⁺ cells typically exhibited K⁺ currents with amplitudes <100 pA, as depicted in Figure S5C‐a and S5C‐b. It has been observed that there is a significant variation in the outward and inward currents among different cell generations. Currents in the form of Figure S5C‐c and S5C‐d were typically recorded in cells of higher generations, exhibiting distinct inward rectification characteristics. Statistical analysis was performed on the currents of cells from different generations, revealing substantial variability in K⁺ currents (Figure S5D).

Additionally, the Ca²⁺ activation properties of K⁺ currents in Sca‐1⁺ cells were investigated. First, an increase in ATP‐induced current was observed. Extracellular ATP signaling activates the release of intracellular Ca²⁺ stores and increases the intracellular free Ca²⁺ concentration ([Ca²⁺]_i) (Figure 3A). Statistical analysis showed that increased [Ca²⁺]_i induced by ATP (100 μM) caused an increase in K⁺ current of 791.9%±1301.7% (n=26 tested cells, at −80 mV) and 284.5%±373.3% (n=26 tested cells, at +80 mV). Current activation showed significant cell‐to‐cell differences. ATP‐induced increases in cellular K⁺ currents are not universal, and statistical analysis revealed that 30 of the 51 recorded cells were activated in response to ATP stimulation. Consequently, ATP resulted in an increase in current in ∼60% of the cells.

Figure 3. K_Ca current in Sca‐1⁺ cells.

A, ATP‐induced current enhancement. Typical recordings (A‐a) and summary data (A‐b) showing that intracellular free Ca²⁺ ([Ca²⁺]_i) and current (A‐c and A‐d) on Sca‐1+ cells were stimulated by extracellular ATP (recorded in Tyrode solution). Statistics in (A‐b) showed the significant increase of [Ca²⁺]_i induced by ATP (P=0.0000, permutation t test, n=22 tested cells). B, Modulation of K_Ca channel activity by intracellular Ca²⁺. Representative perforated recordings (B‐a, B‐c) and standard whole‐cell recordings (B‐b, B‐d) from 2 cells are shown. In each pair (B‐a/B‐b and B‐c/B‐d), the membrane was ruptured by suction to transition from perforated to whole‐cell mode. (B‐a) and (B‐b) were recorded from the same cell with a Ca²⁺‐free pipette solution ([Ca²⁺]_i≈0). (B‐c) and (B‐d) were from another cell with 500 nM [Ca²⁺]_i in the pipette. (B‐e) Statistical data showed that the percentage change of current compared with before cell membrane rupture at −60 mV (P=0.0006, permutation t test) and +60 mV membrane potential (P=0.0027, permutation t test), indicating that the increase of intracellular Ca²⁺ significantly increased the channel current. The “n” denotes the number of cells tested. C, Ionomycin‐induced Ca²⁺ influx enhanced K_Ca channel activity and hyperpolarized the membrane. A representative recording (C‐a) and summary data (C‐b) show that ionomycin significantly increased K_Ca current at +30 mV under symmetric 140 mM K⁺ conditions (P=0.0001, permutation t test, n=9). In a typical experiment (C‐c), the IK_Ca blocker TRAM34 (100 nM) partially reversed this hyperpolarization. Statistical analysis (C‐d) confirmed that ionomycin caused significant membrane hyperpolarization (P=0.0000, permutation t test, n=17). ATP indicates adenosine triphosphate; IK_Ca, intermediate conductance calcium‐activated potassium channel; K_Ca, calcium‐activated potassium channel; Sca‐1⁺, stem cell antigen‐1 positive; and TRAM34, 1‐[(2‐chlorophenyl) diphenylmethyl]‐1h‐pyrazole.

Furthermore, modulation of K⁺ current in Sca‐1⁺ cells was achieved using pipette solutions to regulate intracellular Ca²⁺. The Ca²⁺ within the pipette solution was meticulously buffered to guarantee the desired entry of Ca²⁺ into the cell, ensuring efficient activation of the K⁺ current. After perforating the membrane with nastytin, we recorded the current at the cellular base level of the Ca²⁺ concentration. Subsequently, the current was recorded at lower or higher Ca²⁺ concentrations after buffering the Ca²⁺ in the pipette solution into the cell. This was achieved by rupturing the cell through negative‐pressure suction to form a standard whole‐cell configuration. To eliminate the disruptive influence of sudden current fluctuations resulting from membrane rupture, we initially examined the current using a pipette solution containing a minimal concentration of Ca²⁺ (~0 Ca²⁺), supplemented with 1 mM EGTA (Figure 3B‐a and 3B‐b). Subsequently, alterations in the membrane current were measured at a concentration of 500 nM [Ca²⁺]_i in the pipette solution (Figure 3B‐c and 3B‐d). These findings revealed that elevated levels of intracellular Ca²⁺ led to an increase in ion channel current. Representative recordings are depicted in Figure 3B‐a through 3B‐d, and statistical data are presented in Figure 3B‐e.

In perforated patch clamp mode, the cells were treated with 1 μM ionomycin, a calcium ionophore, and K⁺ current recording was performed. The results showed that among the 29 cells tested, 20 (68.95% of the sample population) responded to ionomycin. The current induced by ionomycin was significantly increased (Figure 3C‐a). Analysis of the effects of ionomycin on K⁺ current demonstrated a significant increase in K⁺ current from 69.52±69.06 to 308.49±198.63 pA (n=9, P=0.0001), representing a remarkable increase of 700%±769% (at +30 mV membrane potential), as shown in Figure 3C‐b.

To investigate the effect of ionomycin on the activation of K_Ca channels by increasing [Ca²⁺]_i, we employed the patch‐clamp current‐clamp technique. Our findings revealed that the activation of K_Ca channels induced by an increase in [Ca²⁺]_i led to membrane hyperpolarization. Furthermore, we discovered that inhibiting IK_Ca channels in Sca‐1⁺ cells using TRAM34 inhibited membrane hyperpolarization induced by ionomycin. Typical recordings are illustrated in Figure 3C‐c, and statistical results of ionomycin‐induced membrane hyperpolarization are shown in Figure 3C‐d. Interestingly, it was found in our experiment that there was considerable variation in the response of cells to ionomycin. Significant differences were observed in the onset time, amplitude change of the induced current, and duration of action among various cell types (Figure S6A displaying 3 representative records). We also found that inhibition of cell membrane hyperpolarization by TRAM34 displayed significant heterogeneity among cells. Specifically, TRAM34 effectively reversed the hyperpolarization induced by ionomycin by an average of 68.87%±31.87% (n=6 tested cells, refer to Figure S6B‐a and S6B‐b representative records). Moreover, the combined application of apamin (small conductance blocker) and TRAM34 (IK_Ca blocker) almost completely reversed the hyperpolarization caused by ionomycin, as evidenced by the typical records in Figure S6B‐c. These findings strongly supported the presence of Ca²⁺ activated K⁺ channels in most Sca‐1⁺ cells. Activation of these channels results in significant hyperpolarization of the cell membrane.

Our study revealed that ATP can effectively initiate temporary changes in intracellular Ca²⁺ levels in a time‐dependent manner. ²⁷ Therefore, we also examined the time‐dependent nature of the alterations in K⁺ current induced by the ATP‐induced increase in [Ca²⁺]_i (typical recordings are depicted in Figure 4A). Consistent with the duration of ATP‐activated [Ca²⁺]_i, the current displayed transient activation characteristics, persisting for ∼3 to 5 minutes. Initially, we assessed the sensitivity of the cells to ATP to establish that the tested cells possessed Ca²⁺ sensitivity (Figure 4B‐a through 4B‐d). After the immediate effect of ATP subsided, we examined the effect of TRAM34 on the same cells to identify the sensitive current of TRAM34 (Figure 4B‐e through 4B‐h). This specific current is known as TRAM34 and the intracellular Ca²⁺‐sensitive current, also referred to as IK_Ca (Figure 4B‐g). Our analysis revealed that TRAM34 effectively reduced the current by 22.8%±16.3% (n=6 tested cells) at a membrane potential of −80 mV and by 24.9%±12.6% (n=6 tested cells) at a membrane potential of +80 mV. These findings highlight the intensity of the IK_Ca current observed in the resting state.

Figure 4. Characteristics of IK_Ca currents in Sca‐1⁺ cells.

A, The ATP‐induced K⁺ current exhibits marked time‐dependent activation. A transient current was consistently evoked, which persisted for ~3 to 5 min. This current was recorded in real‐time from a single cell using both ramp (A‐a to A‐d) and step‐pulse (A‐e to A‐h) stimulation protocols. B, IK_Ca current in vascular Sca‐1⁺ cells. In symmetrical 140 mM K⁺ solution, a small current (~50 pA) was recorded in Sca‐1⁺ stem cells (B‐a). This current was sensitive to 100 μM ATP (B‐b to B‐d), suggesting intracellular Ca²⁺ dependence. Application of 100 nM TRAM34 partially inhibited the current (B‐f). The TRAM34–sensitive component corresponds to the IK_Ca current (B‐g). Recordings in (B‐d) and (B‐h) used ramp pulses; others used step depolarization. All experiments were performed on the same cells with continuous administration, and cells were thoroughly washed after each treatment to remove reagents. ATP indicates adenosine triphosphate; IK_Ca, intermediate conductance calcium‐activated potassium channel; Sca‐1⁺, stem cell antigen‐1 positive; and TRAM34, 1‐ [(2‐chlorophenyl) diphenylmethyl]‐1h‐pyrazole.

IK_Ca Is Involved in the Migration of Sca‐1⁺ Cells Induced by ATP or MCP‐1

Previous studies by our group confirmed that ATP, as a signal substance, can promote the migration of Sca‐1⁺ cells. ²⁷ In this experiment, ATP was used to activate the migration of Sca‐1⁺ cells, and the effect of the IK_Ca channel inhibitor TRAM34 on the ATP‐induced migration of Sca‐1⁺ cells was further examined. As shown in Figure 5A‐a, 30 μM ATP promoted the migration of Sca‐1⁺ cells, which increased by 19.69%±3.83% (P=0.0000, n=9 independent biological replicates) compared with the control group. In the presence of ATP, blockade of IK_Ca channels by TRAM34 inhibited Sca‐1⁺ cell migration by 20.59%±6.21% (P=0.0038, n=9, interaction P=0.0205) (Figure 5A‐a).

Figure 5. Effect of IK_Ca channels on the migration and proliferation of Sca‐1⁺ cells.

A, Effect on migration. ATP increased Sca‐1⁺ cell migration, which was significantly inhibited by the IK_Ca channel blocker TRAM34 (a). MCP‐1 also enhanced migration, and TRAM34 similarly suppressed this increase (b). (Statistical significance was confirmed by HC3‐robust interaction model with 6 biological replicates.) B, The ATP‐enhanced migration of Sca‐1⁺ cells was significantly abolished by IK_Ca channel inhibition with shKCNN4 (P=0.0360, HC3‐robust interaction model, n=5). C, The proliferation of Sca‐1⁺ cells induced by ATP (a, b) and PDGF‐BB (c) was significantly suppressed by inhibition of IK_Ca channels with TRAM34 or shKCNN4 (P values are indicated on the graph; HC3‐robust interaction model, n=5). ATP indicates adenosine triphosphate; CON, control; HC3, heteroskedasticity‐consistent robust SE; IK_Ca, intermediate conductance calcium‐activated potassium channel; MCP‐1, monocyte chemoattractant protein‐1; PDGF‐BB, platelet‐derived growth factor‐BB; Sca‐1⁺, stem cell antigen‐1 positive; shKCNN4, a short hairpin RNA construct specifically designed to target and silence the expression of the KCNN4 gene; and TRAM34, 1‐ [(2‐chlorophenyl) diphenylmethyl]‐1h‐pyrazole.

MCP‐1 was also used to induce Sca‐1⁺ cell migration. The results (Figure 5A‐b) showed that MCP‐1 promoted the migration of Sca‐1⁺ cells, similar to the effect of ATP. MCP‐1 ‐significantly induced an increase in the migration of Sca‐1⁺ cells by 31.19%±1.48% (P=0.0000, n=6 independent biological replicates), while TRAM34 inhibited this MCP‐1‐induced Sca‐1⁺ cell migration by 23.69%±2.29% (P=0.0000, n=6 independent biological replicates, interaction P=0.0000). The shRNA experiments were performed to elucidate the function of IK_Ca. The results demonstrate that when IK_Ca channels were specifically silenced in Sca‐1⁺ cells using shRNA (Figure S7), the migration rate of Sca‐1⁺ cells induced by ATP was reduced significantly from 10.05%±2.87% to 8.30%±3.06% (P=0.0360, n=5) (shown in Figure 5B).

IK_Ca Is Involved in the Proliferation of Sca‐1⁺ Cells Induced by ATP or PDGF‐BB

As shown in Figure 5C, TRAM34 had no effect on the proliferation of Sca‐1⁺ cells under basal conditions, but ATP increased proliferation of Sca‐1+ cell in 19.25%±2.37% (P=0.0000. n=6 independent biological replicates). Blocking IK_Ca channels by TRAM34 reduced ATP‐induced proliferation by 52.90%±6.13% (P=0.0000, n=5 independent biological replicates, interaction P=0.0000) (Figure 5C‐a). After specific knockdown of IK_Ca channels in Sca‐1⁺ cells using shRNA, the knockdown of IK_Ca channels reduced the ATP‐induced proliferation rate of Sca‐1⁺ cells from 39.29%±2.68% (n=5 independent biological replicates) to 30.18%±1.15% (P=0.0000, n=5 independent biological replicates, interaction P=0.0195) (Figure 5C‐b).

In this study, PDGF‐BB was used to induce the proliferation of Sca‐1⁺ cells, and the impact of IK_Ca channels on Sca‐1⁺ cell proliferation was investigated. Our results showed that PDGF‐BB promoted the proliferation of Sca‐1⁺ cells by 53.21%±8.61% (P=0.0000, n=5 independent biological replicates). Blocking IK_Ca channels by TRAM34 reduced PDGF‐BB‐induced proliferation by 29.88%±13.17% (P=0.0012, n=5 independent biological replicates) (Figure 5C‐c).

Functional Cooperation Between IK_Ca and TRPC1 Channels in the Migration and Proliferation of Sca‐1⁺ Cells

Using Western blot, we detected TRPC1 protein expression in Sca‐1⁺ cells, as shown in the representative Figure S8A. Expression was observed in passage 3 (P3), passage 4 (P4), and passage 14 (P14) cells, indicating TRPC1 presence across different cell passages. In our study, the nonselective cation channel blocker 2‐APB was observed to modulate the permeability‐mediated Ca²⁺ influx that occurs subsequent to extracellular calcium entry induced by ionomycin, a calcium ionophore. As shown in Figure S8B, application of ionomycin elicited a pronounced transient elevation in intracellular calcium concentration ([Ca²⁺]i, reported as F340/F380 ratio) in Sca‐1⁺ cells, which was followed by a gradual decline to a sustained plateau phase. Subsequent addition of 100 μM 2‐APB during this plateau phase induced a distinct turning point, demonstrating inhibition of extracellular Ca²⁺ influx. Furthermore, this experiment revealed that 2‐APB significantly inhibited both MCP‐1‐induced migration and PDGF‐BB‐induced proliferation of Sca‐1⁺ cells (Figure S8C and S8D), indicating the involvement of nonselective cation channels in Sca‐1⁺ cell migration and proliferation activities. To further identify the specific ion channel involved in regulating intracellular Ca²⁺ in Sca‐1⁺ cells, we investigated the effect of the TRPC1 inhibitor Pico145 on ionomycin‐induced extracellular calcium influx and the subsequent permeability‐mediated Ca²⁺ influx. The results demonstrated a similar inhibition of Ca²⁺ influx as observed with 2‐APB (indicated by the arrow in Figure S8E).

The combined effects of IK_Ca and TRPC1 channels were investigated. The migration assay results (Figure 6A) showed that compared with the control group (MCP‐1 untreatment), MCP‐1 increased Sca‐1⁺ cell migration by 26.1% (n=6). In the presence of MCP‐1, the addition of TRAM34 reduced the cell migration rate by 24.9% (n=6), whereas blocking TRPC1 with Pico145 inhibited MCP‐1‐induced Sca‐1⁺ cell migration by 20.1% (n=6). The proliferation assay results (Figure 6B) indicated that following PDGF‐BB activation of Sca‐1⁺ cells, both the IK_Ca inhibitor TRAM34 and the TRPC1 inhibitor Pico145 suppressed cell proliferation, inhibiting it by 23.3% (n=3) and 27.2% (n=3), respectively. In Sca‐1⁺ cells overexpressing IK_Ca, administration of TRAM34, Pico145, or both inhibitors almost equally suppressed proliferation, inhibiting it by 20.0% (n=7), 30.7% (n=7), and 34.2% (n=7), respectively (interaction P=0.0309, Figure 6C).

Figure 6. Functional cooperation between IK_Ca and TRPC1 channels in the migration and proliferation of Sca‐1⁺ cells.

A, Migration assays showed that TRAM34 and Pico145 significantly inhibited MCP‐1‐induced migration of Sca‐1⁺ cells (P=0.0015 and P=0.0021, respectively; HC3‐robust interaction model, n=6, interaction P=0.0240). B, Treatment with TRAM34 or Pico145 following PDGF‐BB activation led to a significant suppression of Sca‐1⁺ cell proliferation (P=0.0053 and P=0.0014, respectively interaction P=0.0346). Data are from 6 independent biological replicates and were analyzed by HC3‐robust interaction model. C, In IK_Ca‐overexpressing Sca‐1⁺ cells, treatment with TRAM34, Pico145, or their combination significantly suppressed proliferation (P=0.0119, P=0.0000 and P=0.0000, respectively; HC3‐robust interaction model, n=7 independent biological replicates). Significant interaction effect was observed between TRAM34 and Pico145 (interaction P=0.0309). adCN indicates viral vector devoid of any genetic material; CON, the group serves as the untreated control group; CTR, the baseline control condition for each respective group prior to treatment; HC3, heteroskedasticity‐consistent robust SE; IK_Ca, intermediate conductance calcium‐activated potassium channel; MCP‐1, monocyte chemoattractant protein‐1; PDGF‐BB, platelet‐derived growth factor‐BB; Sca‐1⁺, stem cell antigen‐1 positive; TRAM34, 1‐ [(2‐chlorophenyl) diphenylmethyl]‐1h‐pyrazole; and TRPC1 channel, transient receptor potential canonical channel.

MAPK Signaling Pathway Mediates the Effect of IK_Ca on Migration and Proliferation of Sca‐1⁺ Cells

After stimulating the cells with ATP, MCP‐1, and PDGF‐BB, we investigated the involvement of MAPK signaling pathways in facilitating the effects of IK_Ca on the migration and proliferation of Sca‐1⁺ cells.

Our results demonstrated that treatment of Sca‐1⁺ cells with PDGF‐BB significantly increased the levels of p‐ERK (phosphorylated ERK; P=0.0000), and this effect was significantly inhibited by the IK_Ca channel inhibitor TRAM34 (P=0.0375, n=5) (Figure 7A). Meanwhile, we further investigated the effect of TRPC1 addition on p‐ERK expression levels. As shown in Figure 7B, adding PDGF‐BB significantly increased p‐ERK expression (P=0.0000). Furthermore, the addition of TRAM34 and Pico145 significantly inhibited p‐ERK expression (P=0.0016 and P=0.0002, n=3, respectively).

Figure 7. Signaling pathways mediated the effect of IK_Ca on the migration and proliferation of Sca‐1⁺ cells.

A, PDGF‐BB stimulation significantly increased ERK phosphorylation. This effect was significantly attenuated by IK_Ca blockade (P=0.0375, HC3‐robust interaction model; n=5 independent biological experiments). B, PDGF‐BB stimulation significantly increased ERK phosphorylation. This increase was significantly reduced by blockade of either IK_Ca (P=0.0016) or TRPC1 (P=0.0002; HC3‐robust interaction model; n=3). C, PDGF‐BB stimulation also significantly increased p38 phosphorylation, an effect significantly reduced by IK_Ca blockade (P=0.0101; HC3‐robust interaction model; n=6, interaction P=0.0437). D, MCP‐1 stimulation increased AKT phosphorylation. This increase was significantly attenuated by IK_Ca blockade (P=0.0043, HC3‐robust interaction model; n=3 independent biological experiments). AKT indicates serine–threonine protein kinase; CON, control; CTR, the baseline control condition for each respective group prior to treatment; ERK, extracellular regulated protein kinase; GAPDH, glyceraldehyde‐3‐phosphate dehydrogenase; HC3, heteroskedasticity‐consistent robust SE; IK_Ca, intermediate conductance calcium‐activated potassium channel; MCP‐1, monocyte chemoattractant protein‐1; PDGF‐BB, platelet‐derived growth factor‐BB; p‐ERK, phosphorylated extracellular regulated protein kinase; Sca‐1⁺, stem cell antigen‐1 positive; TRAM34, 1‐ [(2‐chlorophenyl) diphenylmethyl]‐1h‐pyrazole; and TRPC1 channel, transient receptor potential canonical channel.

The level of p‐P38 in Sca‐1⁺ cells was significantly elevated following PDGF‐BB treatment (P=0.0000, n=6), and this effect was significantly inhibited by TRAM34 (P=0.0101, n=6) (Figure 7C).

Our findings indicate that the role of IK_Ca in the activity of MCP‐1 may not be linked to MAPK, as our data did not demonstrate a significant relationship. However, it is possible that this effect is associated with changes in the phosphorylation of AKT (protein kinase B), as illustrated in Figure 7D.

DISCUSSION

This is the first study to investigate the role of ion channels in vascular Sca‐1⁺ cells during a vascular restenosis. Specifically, our research focused on studying the characteristics and activation of IK_Ca channels. Our research findings indicate that under normal circumstances, Sca‐1⁺ cells have a minimal level of IK_Ca channel activation but show promising potential for activation. Additionally, our study highlights the cooperative relationship between IK_Ca and TRPC1 channels in regulating membrane potential and intracellular Ca²⁺ levels. This study also explored the role of IK_Ca channels in the regulation of Sca‐1⁺ cell migration and proliferation during vascular remodeling. The activation of Sca‐1⁺ cells with substances such as ATP, PDGF‐BB, or MCP‐1 enhances their migration and proliferation. On the contrary, inhibition or reduction of IK_Ca channels decreases the migration and proliferation induced by these stimuli, and this process involves the joint role of IK_Ca and TRPC1 channels. Our findings also demonstrate that IK_Ca channels play a role in the proliferation and migration of Sca‐1⁺ cells through the classic MAPK signaling pathway. In a mouse model of femoral artery anastomosis, we observed notable intimal lesions and an increase in tdTomato‐labeled Sca‐1⁺ cells. However, the impact of these effects was significantly diminished in KCNN4‐flox× Sca‐1‐Cre×tdTomato mice (IK_Ca ^−/− mice) following elimination of IK_Ca channels in Sca‐1⁺ cells using tamoxifen. Additionally, the proportion of SMMHC‐labeled mature SMCs derived from Sca‐1⁺ cells was reduced in IK_Ca ^−/− mice. These findings underscore the role of Sca‐1⁺ cells in vascular remodeling and suggest that blocking IK_Ca channels in these cells could be beneficial in preventing neointimal overgrowth and improving outcomes in pathological vascular restenosis.

KCNN4 encodes IK_Ca channels that are found in both humans and mice. These channels have a single‐channel conductance of ∼40 pS. ²⁸ Although channel activity is mostly unaffected by membrane voltage, it is extremely responsive to variations in [Ca²⁺]_i. IK_Ca channels are expressed in various tissues in the body and are present in cells undergoing activation, such as stem cells that transition from a dormant state to active proliferating, migrating, and differentiating. In undifferentiated rat mesenchymal stem cells, a study revealed the presence of IK_Ca channel currents that are sensitive to clotrimazole. ²⁹ IK_Ca channel currents have also been shown in undifferentiated mesenchymal stem cells derived from rabbit bone marrow have also shown evidence of IK_Ca channel currents. ³⁰ Furthermore, Weisbrod et al. ³¹ found that IK_Ca channels were present in human embryonic stem cell‐derived cardiomyocytes. IK_Ca channels are also implicated in vascular pathology. Under normal conditions, VSMCs exhibit a contractile phenotype with minimal IK_Ca activity. However, upon vascular injury or disease, VSMCs shift to a proliferative state, and IK_Ca channels help regulate calcium‐dependent signaling that drives proliferation. Inhibiting IK_Ca channels can suppress VSMC growth by limiting [Ca²⁺]_i increases and reducing phosphorylation of CREB (cAMP response element‐binding protein), c‐Fos, and NOR‐1 (neuron‐derived orphan receptor‐1). ¹⁹ In a rat balloon catheter injury model, IK_Ca expression was upregulated 2 weeks post injury. Daily administration of TRAM34, a selective IK_Ca blocker, reduced intimal hyperplasia by ∼40%. ²⁰ These results suggest that IK_Ca overexpression may contribute to excessive VSMC proliferation in neointima formation, and that targeting IK_Ca could offer a novel therapeutic strategy to prevent restenosis after angioplasty. In our study, we used a differentiation assay to investigate the response of Sca‐1⁺ cells to TGFβ‐1 and found that the Sca‐1⁺ cells used in our study exhibit fundamental stem cell characteristics. To further examine IK_Ca currents in Sca‐1⁺ cells, we conducted patch‐clamp experiments. Using the specific inhibitor, TRAM34, we successfully detected TRAM34‐sensitive IK_Ca channel currents. Interestingly, patch‐clamp recordings of total K⁺ currents in Sca‐1⁺ cells revealed 2 distinct forms of K⁺ currents. The form with a significant inward component was predominantly observed in Sca‐1⁺ cells that had been cultured for >15 passages (Figure S5C‐c and S5C‐d). This observation highlights the diverse nature of Sca‐1⁺ cells and suggests that their electrophysiological properties may vary due to the presence of different cell types, such as fibroblasts, adipocytes, and mesenchymal cells, within the Sca‐1⁺ cell population. Additionally, it is possible that the electrophysiological characteristics of the cells changed as Sca‐1⁺ cells underwent differentiation over successive generations of culture. To ensure the quality of the cells, immunomagnetic bead screening was performed every 5 passages to monitor cell purity.

Research has shown that the K⁺ current in endothelial cells and SMCs, which make up blood vessels, typically ranges from 200 to 300 pA. ³² , ³³ In this experiment, it was found that the K⁺ current in Sca‐1⁺ cells was <100 pA. However, when [Ca²⁺]_i was elevated by ATP or ionomycin stimulation, this small current had the potential to increase by several hundred times. This indicates that the K_Ca present in Sca‐1⁺ cells has a basal current only under normal physiological conditions but can be activated under stress or other stimulation conditions in order to fulfill its role. This phenomenon resembles the physiological characteristics of vascular wall stem cells, which tend to remain quiescent in order to maintain vascular homeostasis. However, under pathological conditions, such as atherosclerosis or surgical injury, vascular wall stem cells can be activated through ion channels and other mechanisms, allowing them to fully use their regenerative and repair abilities in vascular remodeling. Our experiments effectively uncovered the essential characteristics of IK_Ca channels in Sca‐1⁺ cells, as shown in Figures 3 and 4. This study investigated the role of IK_Ca in Sca‐1⁺ cells, spatiotemporal activation of IK_Ca by intracellular Ca²⁺, impact of K⁺ channels on membrane potential, and regulation of IK_Ca on membrane potential. These foundational studies form the basis for further exploration of the regulatory role of IK_Ca channels in stem cells. In our study, we observed that low IK_Ca activity was associated with inherent cellular characteristics. This suggests that elucidating the precise functions of these channels under low‐activity conditions may be challenging. To address this, we used factors frequently elevated in disease states or tissue injury—such as ATP, MCP‐1, and PDGF‐BB—to stimulate the migration and proliferation of Sca‐1⁺ cells. Here, we investigated the role of the IK_Ca channel in regulating these processes. As expected, inhibition of IK_Ca channel activity significantly suppressed the stimulus induced migration and proliferation of Sca‐1⁺ cells. Furthermore, our proliferation assay showed that specifically targeting IK_Ca channels with shRNA effectively inhibited the proliferation of Sca‐1⁺ cells, even in the absence of external stimulation (Figure 5C‐b). The findings of our permutation‐based 2‐factor tests reveal statistically significant interactions (interaction P<0.05) between TRAM34 and ATP, MCP‐1, and PDGF‐BB in biological processes associated with cell proliferation and migration. These results suggest that the influence of TRAM34 is contingent upon the activation of cells by ATP, MCP‐1, and PDGF‐BB. This finding highlights the crucial role of IK_Ca channels in promoting the proliferation and migration of Sca‐1⁺ cells under disease states or tissue injury. Transmembrane ion channels, such as IK_Ca channels, found within the plasma membrane, play a vital role in regulating cell proliferation and migration by controlling membrane potential, intracellular Ca²⁺ signals, and downstream signals. Our study, along with others, ¹³ , ³⁴ , ³⁵ , ³⁶ , ³⁷ , ³⁸ consistently supports the importance and mechanisms of IK_Ca channels in these cellular processes.

Multiple studies have emphasized the vital role of Sca‐1⁺ cells in the vascular remodeling process following injury. In cases of postoperative vascular restenosis, excessive VSMC proliferation is the primary cause of intimal hyperplasia. Research ³⁹ suggests that more severe arterial injury resulting from cutting and anastomosis triggers significant apoptosis in SMCs in the media layer. Adventitial cells expressing Sca‐1, CD44, CD34, and GLI1 differentiate into new VSMCs, contributing to the development of intimal hyperplasia. Furthermore, findings ⁴⁰ highlight the diverse nature of Sca‐1⁺ cells, including endothelial cells, adventitial cells, and subpopulations of adipocytes. Although these cells typically do not participate in SMC formation, they play a crucial role in the differentiation process in the presence of severe arterial injuries. Therefore, the current study used an experimental model of arterial anastomosis known for its effectiveness in observing the migration of adventitial cells into the neointima and their subsequent differentiation into SMCs, following more severe arterial injury. We used KCNN4‐flox×Sca‐1‐CreERT×Rosa26‐tdtomato transgenic mice to investigate the effect of IK_Ca channels on Sca‐1⁺ cells in vascular anastomosis. These mice enabled us to achieve selective knockout of the target IK_Ca gene in Sca‐1⁺ cells, while ensuring lineage tracing. Our hematoxylin and eosin assay and immunofluorescence tracing demonstrated the involvement of Sca‐1⁺ cells in vascular remodeling. Our findings revealed that adventitial Sca‐1⁺ cells labeled with tdTomato migrated from the adventitial layer and contributed to neointimal formation. Interestingly, knockdown of IK_Ca expression significantly reduced the migration process of Sca‐1⁺ cells. Moreover, the Sca‐1⁺ cells present within the neointima exhibited the capacity to differentiate into SMCs expressing αSMA and SHMMHC. Notably, our findings indicated a significant decrease in the proportion of mature SMCs that originated from Sca‐1⁺ cells in the IK_Ca ^−/− group (Figure 2C). In addition, IK_Ca knockout in Sca‐1⁺ cells did not have a significant effect on vascular endothelial repair (Figure S2C).

Some studies hypothesize that IK_Ca activation, increasing K⁺ efflux, induces membrane hyperpolarization, which provides the driving force for extracellular Ca²⁺ entry. This leads to an increase in [Ca²⁺]_i and subsequent activation of Ca²⁺‐dependent downstream signaling pathways. ²³ , ²⁵ , ⁴¹ As early as 2003, Nilius et al proposed that TRP channels provide pathways for Ca²⁺ influx. ²³ Research has found that TRPC1 is expressed in the vascular system and is speculated to be a candidate channel for intracellular Ca²⁺ handling. ⁴² Faouzi et al. ²⁵ reported that TRPC1 participates in IK_Ca activation‐induced Ca²⁺ entry in MCF‐7 cells, and Jia et al. ⁴³ confirmed functional synergy between IK_Ca and TRPC1 in mediating Ca²⁺ influx and ERK1/2 activation to regulate serum‐induced proliferation of SMCs. Our present results demonstrate that IK_Ca participates in regulating vascular remodeling after injury by mediating the proliferation and migration of Sca‐1⁺ cells. Furthermore, this study elucidates the mechanism of functional interaction between IK_Ca and TRPC1 in Sca‐1⁺ cells. These findings offer new experimental insights into the ion channel mechanisms underlying postinjury vascular remodeling. Furthermore, they contribute to the theoretical understanding of these remodeling mechanisms by supplementing existing theories with experimental evidence.

Limitations

One of the main limitations of this study is the insufficient exploration of IK_Ca channel activation and its specific role in promoting Ca²⁺ influx. IK_Ca channels exhibit unique activation properties, enabling them to regulate membrane potential and facilitate Ca²⁺ influx even at negative potentials. While our results confirm that IK_Ca activation induces membrane hyperpolarization in Sca‐1⁺ cells—a process that triggers subsequent TRPC1‐mediated Ca²⁺ influx—it remains unclear whether alternative Ca²⁺ influx pathways exist beyond TRPC1. Therefore, future studies will specifically focus on addressing this question to enhance our understanding in this field. Furthermore, regarding the molecular mechanisms explored, only the canonical MAPK pathway has been definitively implicated thus far. Future studies will systematically investigate other potential pathways, such as Notch, Wnt, or YAP/TAZ (yes‐associated protein/transcriptional coactivator with PDZ‐binding motif) signaling, to broaden our mechanistic insights.

CONCLUSIONS

Our experiments confirmed that IK_Ca knockdown in Sca‐1⁺ cells effectively blocked the intimal hyperplasia process caused by abnormal VSMC proliferation in postoperative restenosis. Importantly, our study reveals for the first time the role of IK_Ca current and its relationship with Ca²⁺ levels, which influence the cell membrane potential and trigger accompanying Ca²⁺ influx through TRPC1 (schematic diagram shown in Figure 8). We have also made significant progress in understanding how MAPK signaling and IK_Ca channels affect the migration and proliferation of Sca‐1⁺ cells. Our results suggest that exploring ion channel regulation in stem cells could provide a unique opportunity to develop more effective and reliable treatments for vascular restenosis.

Figure 8. Schematic diagram illustrating the role of IK_Ca channels in Sca‐1⁺ cells during vascular restenosis.

Activation of IK_Ca currents in Sca‐1⁺ cells promotes K⁺ efflux, inducing membrane hyperpolarization. This hyperpolarization provides the driving force for TRPC1‐mediated extracellular Ca²⁺ influx. The resulting increase in intracellular Ca²⁺ triggers a cascade of downstream signaling events that enhance cell proliferation and migration, thereby contributing to pathological vascular remodeling. ER indicates endoplasmic reticulum; IK_Ca intermediate conductance calcium‐activated potassium channel; MAPK, mitogen‐activated protein kinase; p‐AKT, phosphorylated serine–threonine protein kinase; p‐ERK, phosphorylated extracellular regulated protein kinase; Sca‐1⁺, stem cell antigen‐1 positive; SR, sarcoplasmic reticulum; and TRPC1 channel, transient receptor potential canonical channel.

Sources of Funding

This work was supported by the Sichuan Provincial Science and Technology Support Program (grant numbers 2025HJRC0035, 2022YFS0607, 2026NSFC0499, and 2024 NSFSC0709), National Natural Science Foundation of China (grant numbers 31972909, 82070502, and 32171099), Science and Technology Strategic Cooperation Programs of Luzhou Municipal People’s Government and Southwest Medical University (grant number 2024LZXNYD021).

Disclosures

None.

Supporting information

Figures S1–S8

Unedited Gels