An albumin-armed fibronectin fragment assembly on stents enables immune evasion, thrombosis prevention, and self-adaptive vascular cell selectivity

Abstract

Reconstruction of a healthy endothelium following stent implantation is critical for enhancing vascular healing process. Surface engineering of stents with endothelial cell (EC)-selective molecules represents one of the most effective strategies to promote re-endothelialization. However, the adsorption of blood components accompanied by stenting inevitably mask the surface-modified bioactive molecules, leading to a reduction or alteration in the EC selectivity. Herein, we develop a serum albumin (SA)-armed, extracellular-matrix-derived, EC-selective peptide (i.e., REDV) (REDV-SA) self-assembling amyloid-like coating to address this issue. Our results show that the intrinsic biological properties of SA endow the REDV-SA-functionalized stents with immune escape capabilities and resistance to the adsorption of blood components, thereby creating a favorable microenvironment for REDV to exhibit its high EC selectivity. Of particular significance is the adaptive synergy between SA and REDV, which significantly reinforces EC selectivity by inhibiting smooth muscle cell (SMC) adhesion and preferentially promoting EC over SMC migration. Owing to its high EC selectivity, the stent functionalized with REDV-SA coating achieves nearly complete endothelial coverage within one week in vivo, markedly reduces inflammation, and effectively inhibits neointimal hyperplasia, implying great application potential in vascular implants.

Graphical abstract

The albumin-armed fibronectin fragment self-assembling amyloid-like coating not only endows the vascular stents with immune escape capabilities and resistance to the adsorption of blood components, but also reinforces endothelial cell (EC) selectivity by inhibiting smooth muscle cell (SMC) adhesion and preferentially promoting EC over SMC migration, enabling rapid endothelization in vivo.

Highlights

• •An albumin-armed REDV amyloid-like self-assembling coating was constructed to reduce protein-corona masking and maintain endothelial selectivity on stent surfaces.
• •Albumin-REDV adaptive synergy confers immune evasion, inhibits SMC adhesion, and drives EC-preferential migration and re-endothelialization.
• •REDV-SA-functionalized stents accelerate re-endothelialization and inhibit neointimal hyperplasia, mitigating in-stent restenosis.

1. Introduction

Cardiovascular stents have been widely utilized as primary interventions to restore the patency of stenotic blood vessels [[1], [2], [3], [4]]. However, the failure to promptly re-establish a functional endothelial layer following implantation often result in severe complications, including thrombosis, chronic inflammation, and abnormal proliferation of smooth muscle cells (SMCs), ultimately leading to in-stent restenosis [5,6]. Therefore, promoting rapid and complete endothelialization of the stent surface has been recognized as a critical strategy for improving long-term clinical outcomes [[7], [8], [9]].

To this end, a variety of surface modification approaches have been explored, including the incorporation of bioactive molecules (e.g., heparin, hyaluronan (HA), anti-inflammatory recombinant proteins/peptides, exosomes) and physical cues (e.g., micro/nanopatterning, nanotubes, wettability/stiffness tuning, and shear- or stimulus-responsive surfaces) [[10], [11], [12], [13], [14], [15]]. Among these, the immobilization of endothelial cell (EC)-selective peptides has emerged as one of the most effective strategies to promote EC adhesion and migration [16,17]. In particular, short peptides derived from the extracellular matrix (ECM), such as Arg-Glu-Asp-Val (REDV, a short peptide fragment derived from the CS5 domain of fibronectin) peptide, can specifically interact with integrins on ECs, thereby enhancing EC targeted adhesion. It is worth noting that although REDV can effectively promote EC adhesion, it often restricts EC migration, which represents a significant limitation for complete endothelialization of stents [[18], [19], [20]]. Moreover, their biological efficacy is further compromised under physiological blood-contact conditions. Upon implantation, rapid nonspecific adsorption of plasma proteins, platelets, and red blood cells can mask the immobilized peptides, diminishing or even abolishing their selectivity [21,22]. Furthermore, the adsorbed blood components may trigger coagulation and immune responses, further impeding endothelial regeneration [[23], [24], [25]]. Specifically, thrombus formation creates a physical barrier to EC adhesion and migration, while inflammation exacerbates local tissue damage through cytokine release and immune cell recruitment, thereby promoting SMC proliferation and increasing the risk of restenosis [[26], [27], [28]]. Therefore, constructing a surface microenvironment that resists nonspecific adsorption is essential for preserving peptide functionality and enabling efficient endothelialization on stent surfaces.

Serum albumin (SA), one of the most abundant plasma proteins, has been widely explored in biomedical surface engineering owing to its excellent biocompatibility, low immunogenicity, and intrinsic antifouling properties [[29], [30], [31]]. Previous studies have shown that direct albumin functionalization of implant surfaces can not only alleviate foreign body response in vivo, but also improve endothelial cell selectivity by suppressing the adhesion of undesired cells, highlighting its significant potential for promoting endothelialization of vascular implants. In addition, the low-interference environment provided by albumin interfaces creates an opportunity for the subsequent integration of bioactive recognition functions. However, achieving efficient and stable incorporation of such functional components onto stent surfaces, while balancing interfacial protection and biological recognition under complex blood-contacting conditions, remains a pivotal challenge.

Herein, we developed a vascular cell selective peptide-engineered albumin armor that integrates anti-fouling properties with endothelial-targeting capability for achieving rapid and efficient endothelialization. SA was selected as the structural scaffold, and the EC-selective REDV peptides were engineered onto SA via click chemistry. The resulting REDV-engineered SA underwent oxidation-induced conformational changes and self-assembled into an amyloid-like coating (REDV-SAC) on stents. In this construct, the SA matrix forms an inert interfacial barrier that prevents nonspecific adsorption of blood components after implantation, thereby creating a favorable microenvironment for the REDV peptides to selectively promote EC growth, ultimately facilitating rapid endothelialization on stent surfaces. We found that the REDV-SAC coating effectively resisted fibrin and platelet adsorption and reduced immune cell adhesion, thereby preserving the EC selectivity of REDV. Moreover, SA further enhanced EC selectivity by inhibiting SMC adhesion and preferentially promoting EC over SMC migration. As a result, REDV-SAC modified stents achieved rapid and efficient endothelialization in vivo. This strategy provides a promising avenue for the development of advanced vascular stents and holds strong potential for clinical translation.

2. Experimental section

2.1. Materials

Bovine serum albumin (BSA), Hydrochloric acid (HCl), Ultra centrifugal filter, and Human fibrinogen were obtained from Sigma-Aldrich. Sodium persulfate (SP), Phosphate buffered saline (PBS), 1-Anilinonaphthalene 8-sulfonate (ANS), Thioflavin-T (ThT), and FITC-PEG-succinimidyl easter (FITC-PEG-NHS) were purchased from Aladdin Bio-Chem Technology Co., Ltd. (Shanghai, China). N₃-PEG4-REDV (N₃-REDV) was synthesized from ChinaPeptides Co., Ltd. (Shanghai, China). DMEM-F12, DMEM-High glucose, and fetal bovine serum (FBS) were purchased from GE Healthcare Life Sciences. Endothelial cell medium (ECM) and smooth muscle cell medium (SMCM) were purchased from ScienCell (USA). The cell counting kit-8 (CCK-8) was purchased from DOJINDO. Human umbilical artery smooth muscle cells (HUASMCs), Human umbilical vein endothelial cells (HUVECs) and murine macrophage line RAW 264.7 (MCs) were purchased from the GuangZhou Jennio Biotech Co., Ltd.

2.2. Synthesis of BSA-REDV

BSA-REDV was synthesized via a two-step strategy. In the first step, BSA (62.37 mg, dissolved in 25 mL PBS, pH 8.0) was reacted with DBCO-PEG₄-NHS (0.58 mL, 7.7 mM in DMSO) at room temperature in the dark for 2 h, followed by purification with ultrafiltration (MWCO 3 kDa) and washing with PBS to remove unreacted reagents, yielding BSA-DBCO. In the second step, N₃-REDV (39.8 mg, dissolved in 5 mL PBS, pH 8.0) was added to the BSA-DBCO solution, and the strain-promoted azide-alkyne cycloaddition (SPAAC) reaction was performed at room temperature in the dark for 4 h. The reaction mixture was purified by ultrafiltration and lyophilized to obtain a white BSA-REDV powder, which was stored at −20 °C.

2.3. Characterization of BSA-REDV

The MALDI-TOF-MS was utilized with a microflex LRF-MALDI TOF MS (Bruker) to analyze the mass spectrum of BSA-REDV using sinapinic acid as the matrix. The CD spectra of BSA-REDV in the PBS were analyzed using a J-1500 Spectrometer (Jasco, Japan). Spectra were recorded in the range of 190 to 260 nm at 1 nm intervals. Native BSA was included for comparative analysis as a control. The quantitation of secondary structural content was performed using the analysis software provided by the spectrometer manufacturer.

2.4. Preparation of SAC and REDV-SAC coatings

To prepare SAC and REDV-SAC coatings, 316L stainless steel (SS) substrates and vascular stents were immersed in an aqueous solution of sodium persulfate (2 mg/mL), which was mixed in a 1:1 vol ratio with a BSA solution (1 mg/mL in deionized water, pH adjusted to 5.0 using 0.1 M HCl), and reacted at 37 °C for 24 h. After the reaction, the 316L SS substrates were rinsed with DI water and dried by nitrogen gas for future use, marked as SAC. REDV-SAC were treated with sodium persulfate (2 mg/mL) following the same protocol, and labeled as REDV-SAC. To benefit the demand of different tests, various substrates were coated with SAC and REDV-SAC following above protocol too. To evaluate the substrate versatility of this albumin-based coating, the same protocol was applied to different materials (e.g., silicon (Si), titanium (Ti), polyurethane (PU), poly(vinyl chloride) (PVC), and silicone rubber (SR)), which were correspondingly referred to as SAC.

2.5. Characterization of the SAC or REDV-SAC coatings

To investigate the secondary structure of the assemblies, SAC or REDV-SAC coatings were prepared on quartz substrates by immersing the quartz plates in aqueous mixtures of BSA or BSA-REDV (1 mg/mL) and sodium persulfate (2 mg/mL) at 37 °C for 6 h. After the reaction, the coated substrates were rinsed and dried. Circular dichroism (CD) spectra were then recorded using a J-1500 spectropolarimeter (Jasco, Japan) with a path length of 0.1 mm and a sample volume of 3 mL. Native BSA was set as a control. The quantitation of secondary structural content was performed as previously described.

To evaluate the exposure of hydrophobic residues, ANS fluorescence analysis was conduced. Briefly, native BSA or BSA-REDV (2 mL, 1 mg/mL) was first incubated with ANS (300 μL, 100 μM) for 2 h in the dark, followed by the addition of sodium persulfate (SP, 2 mL, 2 mg/mL). After 6 h reaction, the fluorescence intensity of ANS was quantified using an FLS-920 fluorescence spectrophotometer (Edinburgh Instruments). Emission spectra were collected in the range of 410 to 600 nm with the excitation wavelength set at 355 nm. A mixture of native BSA and ANS without SP served as the control.

The presence of amyloid structure in the SAC or REDV-SAC assemblies was evaluated using ThT as a fluorescent probe. Native BSA or BSA-REDV (2 mL, 1 mg/mL) was mixed with an equal volume of SP (2 mg/mL) and incubated at 37 °C for 6 h. Subsequently, ThT solution (2 mL, 1 mM) was incubated with samples for 15 min in the dark. A mixture of native BSA and ThT without SP served as the control. Then, the samples were analyzed using a fluorescence spectrophotometer, with spectra recorded in the range of 450 to 600 nm and an excitation wavelength of 484 nm.

Dynamic light scattering (DLS) was performed with a zetasizer nano zs90 (Malvern Panalytical) to test the size of particles in mixture of BSA (1.5 mL, 1 mg/mL, pH 5) and SP (1.5 mL, 2 mg/mL, pH 5). As a control, a mixture of native BSA (1.5 mL, 1 mg/mL, pH 5) and DI water (1.5 mL, pH 5) was used, named as BSA.

2.6. Characterization of the SAC or REDV-SAC coatings

The Acid Orange colorimetric method was employed to quantify the density of amino groups on the SAC coating. Briefly, the samples were immersed in an acid Orange II (AO II) solution (500 mM in HCl, pH 4) for 12 h at 37 °C, followed by rinsing with a HCl solution (pH 4) and drying with nitrogen gas. Then, a NaOH solution (pH 12, 200 μL) as a destaticizer was applied to the sample surface for 15 min to release AO II, which binding to the amine group of the coatings though electrostatic binding. The concentration of AO II in the NaOH solution was determined at 485 nm using a microplate reader (BioTek Epoch2). The density of the amine groups was then calculated by using a standard curve based on measurements of AO II at different concentrations.

The SAC or REDV-SAC coatings were also deposited onto mirror-polished 316L SS steels and gold-coated substrates for further characterization. Surface morphology was examined using scanning electron microscope (SEM) (JEOL JSM 7800F). The coatings thinkness was determined by Spectroscopic ellipsometer (W-VASE, J.A. Woollam, USA). The water contact angle (WCA) was measured using a DSA100 drop shape analysis system (KRuSS GmbH Company, Hamburg, Germany). Grazing angle attenuated total reflection Fourier transform infrared spectra (GATR-FTIR) were collected using a Nicolet model 5700 instrument. X-ray photoelectron spectroscopy (XPS) (K-alpha, ThermoFisher, USA) was used to analyze the chemical composition of the coatings and the experimental data were fitted with Avantage software (v.5.976). The C1s peak at 284.8 eV was used as the reference for charge calibration. Raman spectra were collected using a Lab RAM HR Evolution Raman microscope (air cooled frequency doubled Nd:YAG laser wavelength 532 nm), scannig from 100 to 3200 cm⁻¹. Surface zeta potential measurement was performed by SurPASS electrokinetic analyzer (Anton Paar GmbH, Austria). Atomic force microscopy (AFM) was performed by Dimension ICON (Brucker).

The SAC and REDV-SAC were incubated with FITC-PEG-NHS, respectively. After incubation, the samples were washed with distilled water three times to remove unbound fluorescent markers. Then, the sample surfaces were examined using a laser confocal microscope with excitation wavelengths of 488 nm.

To evaluate the adhesive strength and mechanical stability of SAC and REDV-SAC coatings for surface engineering of cardiovascular stents, a balloon dilatation test was conducted. Briefly, a bare 316L SS vascular stent with dimensions of Φ1.65 mm × 18 mm was coated with either SAC or REDV-SAC, and then mounted onto an angioplasty balloon using a stent crimper. The dilation process involved increasing the diameter to Φ3 mm at a pressure of 9 atm for 1 min. Finally, SEM analysis was performed to examine the surface morphology of post-expansion stents.

The long-term stability of REDV-SAC coatings was evaluated by immersing the samples in PBS at 37 °C, with daily repacement of the PBS solution. Samples were collected at different time points, including 0, 1, 2, 3, 4 weeks for weighing (using a Sartorius ultra-micro balance), SEM analysis, and in vitro hemocompatibility evaluation.

Cell culture: HUVECs were cultured in DMEM-F12 medium supplemented with 50% (v/v) ECM and 10% (v/v) FBS. HUASMCs were cultured in DMEM-F12 medium supplemented with 50% (v/v) SMCM and 10% (v/v) FBS. Both medium were supplemented with 1% (v/v) penicillin/streptomycin. HUVECs and HUASMCs were cultured in DMEM-F12 medium supplemented with 10% (v/v) FBS, while macrophages (MCs) were cultured in DMEM-High glucose medium supplemented with 5% FBS. For all tests, HUVECs and HUASMCs were used between the 3rd and 7th passages, and were detached using 0.25% trypsin after reaching 80% confluence.

2.7. Biological performances of cells

The HUVECs, HUASMCs and MCs cells were seeded at a density of 2 × 10⁴ cells/cm² and cultured for 2, 24, and 72 h, respectively. Cell viability was assessed using a CCK-8 kit after 24 and 72 h of incubation, with absorbance measured at 450 nm using a microplate reader. The measurement of cytokines release of MCs were performed using ELISA kits (MULTISCIENCES). The HUVECs cells on the 24-h samples were stained with anti-Vinculin antibody (Cat. No.: FNab09800, FineTest, 1/100 dilution in PBS), followed by Alexa Fluor® 488-conjugated antibody (Cat. No.: FNSA-0055, FineTest, 1/500 dilution in PBS), phalloidin-TRITC (1 μg/mL in PBS) and DAPI (5 μg/mL in PBS). The HUASMCs cells on the 24-h samples were stained with anti-αSMA monoclonal antibody (Cat. No.: bsm-33187M, Bioss, 1/500 dilution in PBS) and Alexa Fluor® 647-conjugated polyclonal antibody (Cat. No.: bs-0296G-AF647, Bioss, 1/500 dilution in PBS), followed by DAPI (5 μg/mL in PBS). MCs cultured on the samples were stained with anti-CD86 polyclonal antibody and anti-CD206 polyclonal antibody (Invitrogen, Carlsbad, USA, 1/500 dilution in PBS), followed by DAPI (5 μg/mL in PBS). Images were captured using a confocal laser-scanning microscope (CLSM, TCS SP5 II, Leica, Germany; software: Leica Application Suite X)0.2.6 Co-culture of HUVECs and HUASMCs.

For the co-culture experiment, HUVECs and HUASMCs were pre-labeled with CMFDA (green, Thermo Fisher) and CMTMR (Orange, Thermo Fisher) cell tracker, respectively. These two types of cells were then mixed at a ratio of 1:1 with a final concentration of 2 × 10⁴ cells/cm². After 2 h of culture, the cells were photographed using an Olympus IX51 fluorescence microscope. The cell counts and ratio of cells were calculated using the ImageJ software (software version 1.53C, NIH, USA).

For migration assay, the L-shaped gravity migration method was employed. Briefly, L-shaped 316L SS (10 × 20 × 0.05 mm) foil with one coated plane and one bare plane (10 × 10 × 0.05 mm for each plane). Then, the bare plane was seeded at a density of 5 × 10⁴ cells/cm² for 24 h to form a complete cell layer. After that, the sample was flipped over and transferred to new cell culture plates with fresh culture medium, causing the cell migration from the bare plane to the coated plane. After 24 h, the migrated cells were stained with Rhodamine 123 and visualized using an Olympus IX51 fluorescence microscope. The cell migration distance was calculated using the ImageJ software. More than 5 photos were selected for each group and analysis was repeated 3 times.

2.8. In vitro hemocompatibility evaluation

To assess hemocompatibility, human fibrinogen solution (Sigma-Aldrich, F3879, 1 g/L in PBS) was labeled with FITC using FITC-labeled goat anti-human fibrinogen (Solarbio, SF108, 1/4 dilution in PBS). Then, Samples were incubated in 24-well plates with the FITC-labeled fibrinogen solution at 37 °C for 2 h. Subsequently, the samples were gently rinsed with PBS three times and then observed by the laser confocal microscope. The fibrinogen particle counts were calculated using the ImageJ software.

For the evaluation of blood compatibility, samples were incubated in BSA-blocked wells with partially heparinized whole rabbit blood (0.25 U/mL) for 1 h on an orbital shaker. After incubation, samples were washed three times in normal saline (0.9% sodium chloride) and fixed for 1 h with a 2.5% glutaraldehyde, and then labeled with anti-fibrinogen antibody (ab34269) and goat anti-rabbit IgG H&L (Alexa Fluor® 488) (ab150077) to visualize polymerized fibrin networks using laser confocal microscopy. Meanwhile, the remaining fibrinogen in whole blood was measured using the fibrinogen (FIB) assay kit (Sunbiote, 1035), with fresh whole blood as a control.

Platelet-rich plasma (PRP) and platelet-poor plasma (PPP) were prepared as described previously. One group of samples were incubated with 0.3 mL of PRP at 37 °C for 30 min, then were rinsed three times with a saline and subsequently fixed with a 2.5% glutaraldehyde. After further dehydration, the platelets on the surfaces of the samples were observed using a SEM. Another group of samples were incubated with 0.3 mL of PRP at 37 °C for 2 h, then, the P-selectin and LDH expression levels of the platelets adhered to each sample were measured using ELISA kits.

2.9. Ex vivo thrombogenicity

All animal experiments were approved by the Dongguan People's Hospital Laboratory Animal Welfare and Ethics Committee (AWEC Issue No. IACUC-AWEC-202410001). In this study, a total of four adult New Zealand white rabbits weighing between 2.5 and 3 kg were anesthetized by intravenous injection of 30 mg/mL pentobarbital sodium solution, with a dosage of 1 mL/kg. A thrombosis formation assay was conducted ex vivo using a model of arteriovenous (AV) shunt in New Zealand White rabbits [32,33]. The test samples (0.8 mm × 1.5 mm × 0.02 mm) were securely attached to the inner surface of an extracorporeal circulation (ECC) system, while the catheters were connected to the left carotid artery and right jugular vein, forming a continuous loop. After circulation 0.5 h and 1 h, samples (bare, SAC, and REDV-SAC) were removed, rinsed with saline solution, and evaluated for occlusion rate by capturing images of cross-sections and measuring lumen cross-sectional areas pre- and post-circulation. The blood flow rate at the end of circulation was normalized to that prior to starting circulation, under the identical pressure pump conditions. Subsequently, the samples were retrieved, photographed, weighed, and fixed. Finally, dehydration was carried out before SEM testing.

2.10. In vivo stent implantation

For stent implantation, sixteen adult New Zealand white rabbits weighing between 2.5 and 3 kg were used in the experiments. Four rabbits were implanted with bare 316L SS vascular stents, SAC-, and REDV-SAC-coated stents in a random manner (with each group, n = 4) into the iliac arteries on both sides using a percutaneous transluminal angioplasty (PTA) balloon catheter. The incision made on the femoral arteries was closed using a 9-0 suture to ensure sufficient blood supply. Following a circulation period of 2 h, the arteries containing the stents were collected and gently flushed with heparin saline solution (50 U/mL). Subsequently, they were fixed using a 2.5% glutaraldehyde solution and longitudinally cut for SEM analysis.

The remaining twelve rabbits were implanted with bare stents and REDV-SAC stents into the bilaterally iliac arteries. After 1 week, 1 month, and 3 months of implantation, the arteries containing the stents were harvested and gently flushed with heparinized saline (each time point 4 animals). For 1-week samples, the arteries containing the stents were collected for SEM and immunofluorescence. The stented arteries were longitudinally bisected; one half was fixed with 2.5% glutaraldehyde, sequentially dehydrated, and observed using SEM. Another part was fixed with a 4% paraformaldehyde solution for 6-12 h, incubated with CD 31 antibody (Novus Biologics) at 4 °C overnight, followed by incubation with Goat Anti-Mouse IgG (H + L)-Alexa Fluor® 488 antibody (Absin Bioscience) at room temperature for 6 h, then, stained with YF®555-Phalloidin for 4 h and with DAPI for 30 min, observed using a laser scanning confocal microscope. For 1 and 3 months-samples, the arteries were collected and cut into three parts for SEM, VG analysis. The VG staining samples were first fixed in 4% paraformaldehyde for 6-12 h, embedded in a solidifying solution (methyl methacrylate and dibutyl phthalate, 4:1 v/v, and benzoyl peroxide, 2% w/v) at 37 °C for 24 h, sectioned into 170 μm thin slices using a hard tissue microtome (BQ1600, Lan Ming Medical Treatment, China), stained with van Gieson stain (S-1515, Exon Biotechnology Co., Ltd., Guangzhou, China), and imaged using an optical microscope (CKX41, Olympus, Japan). Tissue morphometry analysis was conducted using ImageJ software.

2.11. Statistical analysis

The results are presented as the mean ± standard deviation of the mean for each sample. All experiments were independently repeated at least three times, unless otherwise specified. Statistical analysis was performed using one-way analysis of variance (ANOVA) and Student's t-test in GraphPad Prism 8.0 software (GraphPad Prism Software, San Diego, CA, USA) to compare data between different groups.

3. Results and discussion

3.1. Preparation and characterization of SAC

Conformational transition-driven amyloid-like protein coatings have shown great potential in biomedical applications. In this work, we developed an amyloid-like SA coating (SAC) by inducing SA conformational transition with sodium persulfate (SP). The oxidation process disrupts the disulfide (S-S) bonds in SA, enabling the phase transformation and subsequent coating formation (Fig. 1A–C). The disruption of S-S was evidenced by the Fourier-transform infrared spectroscopy (FTIR) and the Raman spectra. As observed, the characteristic O=S=O peak in FTIR and the loss of the S-S signal in Raman spectra confirmed the oxidative cleavage of disulfide bonds (Fig. S1). Further evidence was provided by X-ray photoelectron spectroscopy (XPS) analysis. In the S2p high-resolution spectrum, a relative decrease in the S2p₃/₂ peak at 163.3 eV (corresponding to S-S/S-H) was observed in SAC (Fig. 1D). Meanwhile, two new peaks appeared at 164.3 eV and 167.9 eV, which were assigned to R-SO₃^- S2p_3/2 and R-SO-S-R S2p_3/2, respectively. These results suggest that the disulfide bonds in SA were cleaved and primarily oxidized into sulfonate groups following treatment with SP.

Fig. 1.

Fabrication and characterization of SAC. (A) Reactants used for the fabrication of the SAC coating. (B) Partial S-S bonds breakdown of BSA using oxidant SP. (C) Schematic illustration of the self-assembly mechanism. Formation of nucleation platforms through hydrophobic assembly of BSA, followed by the self-assembly of BSA onto these platforms, then followed by inter-particle fusion and brick packing. (D) High-resolution S2p spectra of BSA and SAC, (E) Quantitative analysis of structural content in BSA and SAC according to the CD analysis. (F) ThT fluorescence and (G) ANS fluorescence spectra of BSA and SAC. (H) Size distributions of BSA and SP mixture against incubation time. (I) Thickness of SAC against incubation time. (J) Surface morphology of SAC after different incubation times. (K) Fluorescence images of the different substrates after labeling by NHS-FITC with green fluorescence. (L) Photographs and SEM image of SAC coated stent before and after balloon dilation.

Following the breakdown of disulfide (S-S) bonds, conformational changes and protein aggregation of SA were triggered. Far-UV circular dichroism (CD) spectroscopy revealed a marked loss of α-helix content for SP treated SA, as indicated by diminished signals at 208 and 222 nm compared to native BSA (Fig. 1E and Fig. S2). Meanwhile, Thioflavin T (ThT) fluorescence spectra showed a significant increase in β-sheet content of SP treated SA (Fig. 1F), indicating protein unfolding and the exposure of hydrophobic residues. However, 8-anilino-1-naphthalenesulfonic acid (ANS) binding assays revealed a substantial reduction hydrophobicity (Fig. 1G). This apparent contradiction can be attributed to the assembly of unfolded SA via hydrophobic interactions, during which hydrophobic residues become buried inside the aggregates, while hydrophilic domains are preferentially exposed on the surface.

To understand the self-assembly process of SA into a coating, we analyzed the physicochemical properties of SAC at different deposition times. As shown in Fig. 1H, the oxidation-mediated self-assembly of SAC proceeds through two distinct stages. Within the first hour, the particle size remained largely unchanged (∼1.5 nm), suggesting that BSA underwent slow conformational transitions and the formation of hydrophobic nucleation sites. This early phase did not contribute significantly to the coating formation, as evidenced by minimal changes in coating thickness (Fig. 1I) and water contact angle (WCA) (Fig. S3). Thereafter, a sharp increase in particle size (∼45.2 nm) was observed, which can be attributed to the attainment of a critical nucleation threshold, beyond which unfolded SA rapidly aggregated via cooperative hydrophobic interactions [34]. As the process continued, these growing SA aggregates deposited onto the substrate and gradually fused into a coating. This was evidenced by a marked increase in coating thickness, from 13.91 nm at 17 h to 5163.81 nm at 48 h, along with concurrent increases in WCA and β-sheet content, indicating progressive film formation. Finally, the SAC coating after 24 h composition was chosen for the following research due to its complete coverage, appropriate thickness, and properly time-cost (Fig. 1J).

The uniform deposition of the coating on the surface of the vascular stent with the complex geometries is crucial for clinical applicability. After confirming that SAC could be formed on a variety of clinically relevant substrates (316L SS, Si, Ti, PU, PVC, and SR) (Fig. 1K), SAC coatings were further applied onto 316L SS cardiovascular stents (Φ1.65 mm × 18 mm). The coated stents were mounted onto angioplasty balloons using a stent crimper and dilated to Φ3 mm at 9 atm pressure for 1 min (Fig. 1L). SEM analysis showed that the SAC coating exhibited excellent mechanical robustness, retaining its structural integrity without delamination or cracking. Additionally, EDS mapping revealed a homogeneous distribution of N signals on stent surface, confirming the presence and uniformity of the SAC coating (Fig. S4). The findings highlight the remarkable adhesive strength and flexibility of SAC coatings, underscoring their potential for surface engineering applications in blood-contacting biomaterials.

3.2. In vitro anticoagulant performance and cytocompatibility of SAC

After implantation, the exposed stent surface rapidly adsorbs blood components such as fibrinogen and platelets, which can trigger complications including acute thrombosis and inflammation, ultimately compromising surface functionality [35]. Therefore, we first evaluated the effectiveness of the SAC coating in inhibiting adhesion and activation of fibrinogen and platelet (Fig. 2A). Firstly, the SAC coated 316L SS foils were exposed to the FITC-labeled fibrinogen solution and fresh whole blood. As shown in Fig. 2B, fluorescence imaging revealed significantly reduced fibrinogen adhesion on the SAC-coated surface compared with the bare 316L SS surface, which exhibited extensive fibrinogen adsorption. To better mimic physiologically relevant conditions, the samples were incubated with heparin-anticoagulated whole blood. Under these conditions, the bare SS surface developed a dense fibrin network, indicating a high risk of thrombus formation. In contrast, no fibrin network was observed on the SAC surface. Further quantitative analysis (Fig. 2C) demonstrated that the SAC coating significantly reduced fibrinogen adhesion and fibrin formation by approximately 3-fold and 4-fold, respectively, compared to the control. For evaluating platelet adhesion and activation on the SAC surface, bare and SAC-coated 316L SS samples were incubated in platelet-rich plasma (PRP) at 37 °C for 1 h and analyzed by SEM. Platelet activation was assessed based on morphological classification into five states of increasing activation: round, dendritic, spread dendritic, spreading, and fully spread. As shown in Fig. 2D, the bare SS surface exhibited a high density of spread dendritic or fully spread platelets, indicating substantial activation. In contrast, only a few round, non-activated platelets were observed on the SAC-coated surface. Quantitative analysis further confirmed that SAC significantly reduced platelet adhesion and activation by approximately 8-fold and 54-fold, respectively, compared to the bare SS surface (Fig. 2E). These results together suggest that the SAC has an inhibitory effect on fibrin polymerization and effectively hinders platelet adhesion and activation.

Fig. 2.

In vitro anticoagulant performance and cytocompatibility of SAC armor. (A) Schematic illustration of the selective anti-adhesion and endothelial-favoring properties of the coating. (B) Fluorescence images of fibrinogen and fibrin network formation on different surfaces. (C) Quantitative analysis of fibrinogen particles density (n = 4, mean ± SD) and fibrinogen consumption (n = 3, mean ± SD). (D) SEM images of platelet on different surfaces and corresponding (Yellow pseudo-colors indicates platelets). (E) quantitative analysis of platelet adhesion and activation. (F) SEM and fluorescence images of blood cell adhesion on different sample surfaces after whole blood incubation (Red pseudo-colors indicates red blood cells, white indicates white blood cells, and yellow indicates platelets). (G) Representative fluorescence staining images of MCs cultured for 24 and 72 h on the 316L SS before and after coated with BSA. (H) Cell viability of RAW264.7 MCs grown on different surfaces after 24 and 72 h of culture determined by CCK-8 assay. (I) Fluorescence images and (J) migration distance of HUVECs and HAVSMCs on the different samples for 24 h. (K) Competitive adhesion of HUVECs (green) and HUASMCs (red) for co-culture on different surfaces after 2 h. (L) Quantification of the HUVEC-to-HUASMC adhesion ratio after 2 h of co-culture. One-way ANOVA followed by Tukey's multiple comparisons test was used for statistical analysis with ∗p < 0.05, ∗∗p < 0.01 and ∗∗∗p < 0.001.

To further evaluate the hemocompatibility of the SAC coating under physiologically relevant conditions, whole blood incubation assays were conducted and analyzed by SEM and DAPI nuclear staining. As shown in Fig. 2F (left), the bare SS surface exhibited substantial adhesion of various blood components, including a large number of erythrocytes, activated platelets and leukocytes, which was indicated by their irregular morphology and surface spreading. These observations suggest that the bare metal surface readily interacts with blood cells and may trigger immune activation. In contrast, the SAC-coated surface exhibited minimal blood cell adhesion, indicating its strong resistance to nonspecific adsorption of blood components and immune cells. DAPI staining further supported these findings (Fig. 2F, right), revealing the presence of nucleated cells on the SS surface, while the SAC-coated surface exhibited almost no nuclear signal. Then, we assessed the attachment and cytokine release of RAW 264.7 MCs on the SAC to evaluate the inflammatory response. It can be observed that the SAC show a significant reduction in the adhesion and proliferation of MCs after 24 and 72 h incubation compared to SS (Fig. 2G, H and Fig. S5). These results demonstrate that the SAC coating significantly inhibits leukocyte adhesion and activation, suggesting its immune-evasive property.

The attachment and proliferation of vascular cells on interventional devices, particularly vascular stents, also necessitate focused attention. To assess the effects of the SAC coating on vascular cells, HUVECs and HUASMCs were separately seeded on SAC-coated substrates. As shown in Fig. S6, the SAC coating significantly suppressed the proliferation of both cell types. In addition, the migration distances of HUVECs and HUASMCs on the SAC surface were reduced by approximately 1.6-fold and 2.7-fold, respectively, compared to those on 316L SS (Fig. 2I and J), indicating that the SAC coating exhibits strong anti-fouling behavior against both endothelial and smooth muscle cells. Notably, the inhibitory effect of SAC appeared more pronounced for HUASMCs than for HUVECs. To further investigate this selectivity, we conducted a co-culture experiment to examine the competitive adhesion behavior of the two cell types. HUVECs and HUASMCs were pre-labeled with different fluorescent tracking dyes and co-seeded at a 1:1 ratio on SAC and SS surfaces. After 2 h of co-incubation (Fig. S7), the SAC coating significantly reduced the adhesion of both cell types, but the inhibition of HUASMCs was substantially stronger than that of HUVECs (Fig. 2K and L), indicating a cell-type selective anti-adhesive effect. Previous studies have shown that the cell selectivity of SA-based coatings primarily arises from nanoscale interstitial structures formed during self-assembly. These structures restrict the nonspecific adsorption of large plasma proteins while allowing small serum proteins to permeate, thereby creating a low-adhesion microenvironment. Endothelial cells (ECs), due to their ability to extend lamellipodia in multiple directions, can form stable focal adhesions even at sparse attachment points, allowing for effective surface colonization. In contrast, SMCs, characterized by high myosin II activity and strong intracellular contractile forces, depend on dense and stable adhesion zones to maintain cell spreading [36].

3.3. Synthesis of REDV-functionalized BSA and construction of the REDV-SAC coatings

Given the demonstrated antifouling and immune-evasive properties of SAC, it was employed as a bioinert carrier for the REDV peptide. REDV was engineered to BSA via strain-promoted azide-alkyne cycloaddition (SPAAC), thereby obtaining REDV-engineered BSA (BSA-REDV) (Fig. 3A). The resulting BSA-REDV, with an average grafting ratio of 18:1 (REDV:BSA), was confirmed by matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry (Fig. 3B). Circular dichroism (CD) was further carried out to investigate the secondary structural changes of BSA after REDV conjugation. The CD spectra (Fig. 3C) and corresponding quantitative analysis (Fig. S8) revealed a slight decrease in α-helix and random coil content, indicating minor conformational changes with preserved BSA structure.

Fig. 3.

Synthesis of REDV-functionalized BSA and construction of the REDV-SAC coating. (A) Molecular structures of N₃-REDV peptide and DBCO-NHS, and schematic illustration of grafting REDV onto BSA through the disulfide exchange and the Michael addition reaction. (B) MALDI-TOF-MS spectra and (C) CD spectra of native BSA and REDV-grafted BSA. (D) Schematic illustration of the self-assembly mechanism of the REDV-SAC coating. (E) CD spectra and corresponding (F) quantitative analysis of BSA-REDV and REDV-SAC. (G) Fluorescence images (left), SEM images (middle), and AFM images (right) of SAC and REDV-SAC. (H) FTIR spectra, (I) High-resolution S2p spectra, (J) High-resolution C1s spectra, and (K) surface zeta potential of SAC and REDV-SAC. (L) Photographs of REDV-SAC uncoated and coated stent before and after balloon dilation. (M) SEM images of REDV-SAC coated stent after balloon dilation. (N) Surface morphology and corresponding (O) weighing analysis of REDV-SAC before and after treatments by PBS for different week. Data presented as mean ± SD (K, n = 3; O, n = 4).

The SP-triggered conformation transition and aggregation of BSA-REDV followed a similar α-helix to β-sheet shift as observed in native BSA (Fig. 3D–F). Based on this mechanism, BSA-REDV coating (termed REDV-SAC) was prepared by SP-induced assembly of BSA-REDV. The SAC and REDV-SAC coatings on the 316L SS both exhibited dense and complete coverage (Fig. 3G, right), with similar morphologies consisting of fused nanoscale-particles (Fig. 3G, middle) and comparable roughness (RA: 100 nm) (Fig. 3G, left). The chemical composition and structure of SAC and REDV-SAC were detected by FTIR and XPS. As shown in Fig. 3H–J, both SAC and REDV-SAC exhibited characteristic peaks at 1650, 1537, 1245, and 1042 cm⁻¹, corresponding to Amide I, II, III, and O=S=O groups, respectively, indicating their similar chemical structure. However, REDV-SAC exhibited higher atomic contents of nitrogen, oxygen, and sulfur, as well as elevated O/C, N/C, and S/C ratios (Table S1), suggesting successful integration of REDV peptides. This was further supported by XPS fine-scan spectra, which revealed increased proportions of disulfide (S-S) and sulfonate (R-SO₃^-) bonds, as well as elevated carboxyl (COOH) content in REDV-SAC compared to SAC (Fig. 3J and K). Additionally, REDV integration decreased the WCA by 15.68° and increased the surface potential from 13.95 ± 11.3 to 41.72 ± 13.77 μV (Fig. S9 and Fig. 3K).

We further verified the adhesion and mechanical properties of the REDV-SAC on cardiovascular stents. REDV-SAC was applied to 316L stainless steel (SS) stents (Φ1.65 mm × 18 mm), resulting in a visible color change from metallic silver to yellow (Fig. 3L). SEM analysis confirmed that the REDV-SAC coating retained its surface integrity after dilation, with no signs of peeling or cracking on the stent struts (Fig. 3M). This was further corroborated by EDS, which showed a homogeneous distribution of N element across the stent surface (Fig. S10). To assess long-term stability, REDV-SAC-coated samples were immersed in PBS for 4 weeks. SEM analysis revealed that the REDV-SAC remained intact throughout the immersion period. Notably, after 1 week, the initially distinct particles fused into a continuous film, with particle boundaries gradually blurring over time (Fig. 3N). In terms of mass loss, the coating exhibited an initial mass loss of ∼7.26% (17.3 ± 6.4 μg) in the first week, followed by minimal additional loss (∼2.56%, 6.0 μg) over the subsequent three weeks (Fig. 3O). These results highlight the robustness of the REDV-SAC coating, supporting its potential for long-term application in cardiovascular stent surface engineering.

3.4. In vitro and ex vivo evaluation of the anti-thrombogenic and anti-inflammatory properties of REDV-SAC coatings

To preliminarily evaluate the hemocompatibility of REDV-SAC, we investigated its regulatory effects on fibrinogen and platelet adhesion and activation. Immunofluorescence staining [37] revealed that both REDV-SAC and SAC significantly inhibited fibrinogen adhesion and fibrin network formation compared to 316L SS (Fig. 4A and B). In addition, platelet adhesion and activation assays demonstrated that REDV-SAC effectively inhibited platelet adhesion and activation, as evidenced by reduced platelet deposition, lower LDH release, and decreased expression of the activation marker P-selectin (Fig. 4C, D and Fig. S11). Under whole-blood conditions, REDV-SAC further reduced the adhesion of blood components, including red blood cells and leukocytes, indicating improved hemocompatibility (Fig. 4E). Taken together, REDV-SAC maintained comparable hemocompatibility to the SAC coating. It is well established that thrombosis and inflammation form a self-reinforcing loop at blood-material interfaces. Platelet activation and fibrin scaffolding provide adhesive and biochemical cues, such as P-selectin/PSGL-1 engagement and complement activation, that recruit leukocytes, amplify cytokine release, and sustain inflammatory signaling. Platelet-derived mediators together with fibrin(ogen) further prime macrophage adhesion and activation, biasing them toward a pro-inflammatory phenotype [38]. Given this thrombo-immune coupling, we evaluated the adhesion and cytokine expression of RAW 264.7 on REDV-SAC. Fluorescence imaging and CCK-8 assay (Fig. S12) showed significantly higher MCs adhesion and proliferation on bare 316L SS, while both SAC and REDV-SAC exhibited minimal cell attachment with no statistical difference between them. Immunofluorescence staining further showed that (Fig. 4F), compared with bare 316L SS, both SAC and REDV-SAC markedly reduced the number of adherent macrophages, with a corresponding decrease in the number of visible CD86-positive adherent cells, which is consistent with the low-fouling and anti-adhesive characteristics of the albumin-based surfaces. Consistently, ELISA analysis revealed a marked reduction in TNF-α expression on REDV-SAC compared with 316L SS, while no significant change was observed in IL-10 expression (Fig. 4G and H). Collectively, these findings suggest that the albumin-based coating mainly functions as a low-fouling, anti-adhesive interface that reduces macrophage attachment, rather than actively promoting macrophage polarization.

Fig. 4.

In vitro and ex vivo evaluation of the anti-thrombogenic and anti-inflammatory properties of REDV-SAC coatings (A) Fluorescence images of fibrinogen and fibrin network on different surfaces and corresponding (B) quantitative analysis of adhesion fibrinogen particles density (n = 7, mean ± SD) and decrease of fibrinogen (n = 3, mean ± SD). (C) SEM images of platelet on different surfaces and corresponding (Yellow pseudo-colors indicates platelets.) (D) quantitative analysis of platelet adhesion and activation. (E) SEM images of whole blood cell adhesion on different sample surfaces after whole blood incubation. (Red pseudo-colors indicates red blood cells, white indicates white blood cells, and yellow indicates platelets.) (F) Immunofluorescence images of RAW 264.7 macrophages on different sample surfaces after incubation. (CD86 (M1, green), CD206 (M2, red), and nuclei (blue)) (G) TNF-α and (H) IL-10 expression of MCs analyzed by ELISA assay. (I) Schematic illustration of rabbit AV shunt model for the ex vivo blood compatibility test. Photographs of (J) cross-sectional views and (K) lumen side thrombus of different surfaces after 1 h circulation. (L) Representative SEM images of the lumen side of different surfaces after circulation. Measurement of (M) occlusion ratio, (N) thrombus weight, and (O) blood flow rate after circulation. Data presented as mean ± SD (D, n = 3; I to K, n = 4). One-way ANOVA followed by Tukey's multiple comparisons test was used for statistical analysis with ∗p < 0.05, ∗∗p < 0.01 and ∗∗∗p < 0.001.

We further evaluated the effectiveness of the REDV-SAC in inhibiting the thrombogenicity ex vivo (Fig. 4I). After 1 h blood circulation, 316L SS exhibited severe thrombus formation, whereas both SAC and REDV-SAC showed only minimal clot deposition (Fig. 4J and K). SEM analysis revealed that 316L SS surfaces were densely covered with fibrin networks, activated platelets, and trapped red blood cells within 1 h, while SAC and REDV-SAC surfaces retained their original nanoparticulate morphology with minimal protein deposition (Fig. 4L). Quantitative analysis revealed a dramatic reduction in occlusion rates (9% and 7.5% for SAC and REDV-SAC vs. 100% for 316L SS), thrombus weights (14.1- and 16.4-fold lower, respectively), and fully preserved blood flow for both SAC and REDV-SAC group (Fig. 4M − O). These results implied the promising anti-thrombogenic property of the REDV-SAC surface.

3.5. Regulation of the behaviors of vascular cells on the REDV-SA coatings

Rapid endothelialization and inhibition of excessive smooth muscle cell (SMC) proliferation are essential for ensuring long-term efficacy of cardiovascular implants. Therefore, we further evaluated the regulatory effects of different surfaces on HUVECs and HUASMCs. As shown in Fig. 5A–C, immunofluorescence staining of vinculin and F-actin revealed that both SAC and REDV-SAC enhanced HUVEC adhesion and spreading compared with 316L SS, as indicated by more organized stress fibers and increased vinculin patches. This result suggests that the albumin-based interface itself already provides a more favorable microenvironment for endothelial adhesion and cytoskeletal organization [36]. Notably, REDV-SAC showed a further enhancement, and quantitative analysis confirmed that the REDV-SAC group exhibited the highest vinculin expression per cell. On this basis, the further enhancement observed on REDV-SAC can be attributed to the additional endothelial-targeting effect of the REDV motif. In addition, CCK-8 assay results demonstrated that REDV-SAC significantly promoted HUVEC proliferation at both 24 and 72 h (Fig. 5B), further indicating a more favorable microenvironment for endothelial growth. Furthermore, in the case of HUASMCs, REDV-SAC markedly inhibited cell proliferation compared to 316L SS (Fig. 5D and E). We further investigated the comparative growth patterns of HUVECs and HUASMCs on REDV-SAC-coated surfaces. After 2h coculture, the REDV-SAC selectively promoted the adhesion and spreading of HUVECs relative to HUASMCs, exhibiting highest HUVECs/HUASMCs ratio among groups (Fig. 5F, G and Fig. S13). To investigate the potential of rapid endothelialization, we assessed the migratory behavior of HUVECs and HUASMCs on REDV-SAC surface by quantifying the migration distances. Notably, REDV-SAC promoted HUVECs migration and maintained inhibition on HUASMCs migration compared to the 316L SS and SAC (Fig. 5H and I). This result demonstrates that our design effectively overcomes the inherent limitation of conventional REDV-grafted surfaces, where the immobilized REDV peptides often restrict EC migration [21]. The enhanced migration of ECs in this work can be attributed to the albumin-based low-adhesion interface, which facilitates effective EC colonization by supporting lamellipodia extension and focal adhesion formation, as previously discussed [36].

Fig. 5.

Regulatory effects of the REDV-SAC coatings on vascular cells. (A) CLSM images of HUVECs stained with Vinculin (green), F-actin (red) and DAPI (blue) on different surfaces after 1 day of culture. Cell viability of HUASMCs grown on different surfaces after (B) 24 and 72 h of culture determined by CCK-8 assay. Graphs showing the quantification of confocal images for (C) number of vinculin patches. and (D) CLSM images of HUASMCs stained with α-SMA (red) and DAPI (blue) on different surfaces after 1 day of culture. (E) Cell viability of HUASMCs grown on different surfaces after 24 and 72 h of culture determined by CCK-8 assay. (F) Competitive adhesion of HUVECs (green) and HUASMCs (red) for co-culture on different surfaces after 2 h. (G) The ratios of the number of HUVECs to the number of HUASMCs adhered on different surfaces after 2 h of co-culture. (H) Migration of HUVECs on different surfaces after 1 day of culture. (I) Migration of HUASMCs on different surfaces after 1 day of culture. Data are presented as mean ± SD (n = 6). One-way ANOVA followed by Tukey's multiple comparisons test was used for statistical analysis with ∗p < 0.05, ∗∗p < 0.01 and ∗∗∗p < 0.001.

3.6. Vascular stent deployment in rabbit iliac arteries

Prior to stent implantation in rabbits, the in vivo blood compatibility of REDV-SAC was first evaluated to verify its anti-thrombogenicity and determine an appropriate anticoagulation protocol (Fig. 6A). First, SAC or REDV-SAC coated stents were parallelly placed into iliac arteries of New Zealand white rabbits with bare 316L SS stents, with low heparin (150 U/Kg) systemic anticoagulation (Fig. 6B). After 2 h implantation, the blood vessels containing stent were collected to evaluate acute thrombus formation. SEM analysis revealed that the bare 316L SS stent was covered by a layer of radially extending thrombus, consisting of dense fibrin networks and red blood cells. In contrast, only a few non-activated platelets were observed on the SAC and REDV-SAC stents (Fig. 6C). The result confirmed the in vivo anti-thrombogenic property of REDV-SAC. Notably, a reduced dose of heparin (150 U/Kg) (as compared to the conventional 300 U/kg) was sufficient for implantation of REDV-SAC stent.

Fig. 6.

Stenting in rabbit model. (A) The synergy between BSA and REDV enables multidimensional regulation of the vascular microenvironment, including the suppression of blood cell and inflammatory cell adhesion, inhibition of SMC activation and proliferation, and enhancement of EC adhesion and functional expression, thereby effectively promoting rapid re-endothelialization. (B) Schematic of stent implantation in the iliac arteries of rabbits. (C) Representative SEM images of thrombosis on the harvested stents after 2 h circulation. (D) SEM images of bare and REDV-SAC-coated stents after implantation for 1 weeks. (E) Immunostaining of ECs (green: CD31, red: F-actin, blue: DAPI) on bare and REDV-SAC-coated stents after 1-week implantation. (F) SEM images of bare and REDV-SAC-coated stents after implantation for 4 and 12 weeks. (G) Representative optical images of stented arteries cross sections after van Gieson staining and quantitative analysis of ISR after implantation for (H) 4 and (I)12 weeks. Data presented as mean ± SD (G, n = 4 independent animals for 316L SS, REDV-SAC, respectively). T-tests was used for statistical analysis with ∗p < 0.05, ∗∗p < 0.01 and ∗∗∗p < 0.001.

Then, we implanted the REDV-SAC-coated stents and bare 316L SS stent into the iliac arteries of rabbits. At 1, 4, and 12 weeks post-stent implantation, the implanted arteries were harvested and histologically evaluated. SEM revealed that, at 1 week, the bare stent surface was uniformly covered by a radially organized, fibrin-rich extracellular matrix (Fig. 6D). In contrast, distinct EC outlines were still clearly visible on REDV-SAC stents. Immunostaining further confirmed that the REDV-SAC stents were covered with an intact EC layer and maintained structurally integrated with the surrounding vessel walls. In comparison, the dense coverage observed on bare 316L SS stents consisted mainly of ECM with only sparse ECs, as indicated by the limited distribution of cell nuclei on the stent struts (Fig. 6E and Fig. S14). These findings highlight the excellent reendothelialization capacity of REDV-SAC-coated stents in vivo. Moreover, SEM images showed that ECs exhibited robust and oriented growth on the REDV-SAC stent at 4 weeks, which was well sustained until 12 weeks. Meanwhile, ECs exhibited migration on the bare 316L SS surface at 4 weeks, while a complete EC layer was not observed after 12 weeks (Fig. 6F). These results demonstrated that the REDV-SAC surface could significantly promote reendothelialization process on vascular stents.

The incidence of ISR remains a major concern for long-term stent implantation. SEM analysis showed that bare stents were obscured by a dense tissue layer suggestive of early neointimal hyperplasia at 4 weeks post-implantation, whereas REDV-SAC coated stents maintained a clearly visible strut outline even at 12 weeks (Fig. S15). As observed, the bare stents exhibited markedly thicker neointimal layers than the REDV-SAC coated counterparts (Fig. 6G). Quantitative analysis revealed that REDV-SAC coatings significantly suppressed neointimal hyperplasia. Compared with bare 316L SS stents, REDV-SAC coated stents reduced area stenosis from 37.8% to 23.8% at 4 weeks and from 21.5% to 8.5% at 12 weeks, and decreased neointimal thickness (NT) from 235.8 μm to 110.8 μm at 4 weeks and from 129 μm to 27.4 μm at 12 weeks (Fig. 6H and I).

4. Conclusion

In this study, we developed a REDV-engineered SA armor via a facile oxidation-induced conformational transition for achieving rapid and efficient endothelialization. By leveraging the intrinsic antifouling and immune-evasive properties of SA, this armor effectively protects REDV peptides from nonspecific adsorption of blood components, thus highly preserving their EC selectivity under physiological conditions. Moreover, the SA matrix synergistically enhanced this selectivity by inhibiting SMC adhesion and preferentially promoting EC over SMC migration, thereby effectively overcoming the intrinsic migration limitation of REDV-modified surface. Consequently, REDV-SA-coated stents achieved rapid endothelial coverage, reduced inflammation, and significantly attenuated neointimal hyperplasia in rabbit.

Compared with conventional vascular stents, including bare metal stents (BMS) and drug-eluting stents (DES), REDV-SA represents a distinct pro-regenerative bio-functional strategy. Rather than relying solely on mechanical support or anti-proliferative drug release, this design aims to suppress restenosis by promoting physiological vascular healing, particularly rapid endothelialization, while simultaneously reducing thrombo-inflammatory interference. In this regard, REDV-SA provides an alternative route to vascular repair based on restoration of vascular homeostasis. Such a pro-regenerative concept may offer several important advantages, including reduced interference with endothelial recovery, improved overall healing quality, and integration of antifouling protection, endothelial selectivity, and vascular healing regulation into a single bio-functional interface. Nevertheless, this strategy may also face challenges, particularly in highly proliferative or complex lesions where its efficacy may be less pronounced than that of current DES, and its long-term stability and functional persistence in vivo still require further evaluation. Overall, these findings highlight the REDV-SA coating as a promising strategy for next-generation vascular implants with enhanced biocompatibility and re-endothelialization performance.

CRediT authorship contribution statement

Qing Ma: Writing – original draft, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Wenxuan Wang: Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Xiaohui Mou: Writing – review & editing, Project administration, Investigation, Funding acquisition. Wentai Zhang: Writing – review & editing, Project administration. Yuting Huang: Methodology, Investigation. Siwei Yin: Methodology, Investigation. Wenjie Tian: Writing – review & editing, Project administration, Methodology, Funding acquisition, Conceptualization. Jianbing Zhu: Writing – review & editing, Project administration, Methodology, Funding acquisition, Conceptualization. Zhilu Yang: Writing – review & editing, Project administration, Funding acquisition, Conceptualization.

Ethics approval and consent to participate

All animal experiments adhered to the Laboratory Animal Administration Rules of China and were approved by the Dongguan People's Hospital Laboratory Animal Welfare and Ethics Committee (Approval No.: IACUC-AWEC-202410001).

Data availability statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Declaration of competing interest

Prof. Zhilu Yang is an editorial board member for Bioactive Materials and was not involved in the editorial review or the decision to publish this article. All authors declare that there are no competing interests.

Appendix A. Supplementary data

The following is the Supplementary data to this article.