Activation of the YAP1/pSTAT3/NRP1 axis in peritendinous sensory nerves promotes tendon healing

Jiayi Wang; Fan Wang; Jingwen Liu; Yao Xiao; Zhaoyang Li; Xiaonan Liu; Peilin Zhang; Fei Wang; Wenguo Cui; Shen Liu

doi:10.1126/sciadv.aec1272

. 2026 Feb 18;12(8):eaec1272. doi: 10.1126/sciadv.aec1272

Activation of the YAP1/pSTAT3/NRP1 axis in peritendinous sensory nerves promotes tendon healing

Jiayi Wang ^1,^†, Fan Wang ^2,^†, Jingwen Liu ^1,^†, Yao Xiao ^1,^†, Zhaoyang Li ¹, Xiaonan Liu ¹, Peilin Zhang ¹, Fei Wang ^2,^*, Wenguo Cui ^2,^*, Shen Liu ^1,^*

PMCID: PMC12915599 PMID: 41706840

Abstract

Chemotactic migration of peritendinous nerves is essential for tendon regeneration, yet the underlying neuroelectrical mechanisms remain unclear. Here, we identify an electrically responsive yes-associated protein 1 (YAP1)/phosphorylated signal transducer and activator of transcription 3 (pSTAT3)/neuropilin-1 (NRP1) signaling axis in sensory neurons. Electrical stimulation enhances YAP1-pSTAT3 interaction, promotes pSTAT3 nuclear translocation and transcriptional activity, and up-regulates NRP1 to support growth of calcitonin gene-related peptide (CGRP)–positive sensory fibers. Guided by these findings, we engineered a bifunctional piezoelectric patch composed of poly(vinylidene difluoride-trifluoroethylene) [P(VDF-TrFE)] and regenerated silk fibroin@poly(3,4-ethylenedioxythiophene):polystyrene sulfonate (RSF@P:P), coupling mechanically induced electrical cues with dynamic lubrication. Under ultrasound activation, the P(VDF-TrFE) layer generates localized electrical signals that facilitate sensory-nerve and vascular ingrowth, while the RSF@P:P layer undergoes piezoelectric-triggered gel-sol transition to form a low-friction interface and reduce adhesion. In rat and Bama minipig models, the patch markedly enhanced tendon regeneration and decreased adhesion scores by ~50%. These findings establish a neuroelectrically guided strategy for enhancing tendon healing.

A responsive patch harnesses electro-activated nerve signaling to accelerate tendon regeneration and prevent adhesions.

INTRODUCTION

Tendons serve as essential force-transmitting structures that link muscles to bones and enable locomotion. However, the intrinsically low cellularity, limited vascular supply, and sparse innervation of tendons severely constrain their healing potential following acute rupture or chronic degeneration (1), rendering these injuries among the most disabling musculoskeletal disorders. In the United States alone, over 300,000 tendon repair surgeries are performed annually (2), but outcomes often fall short of expectations (3), with the most prominent issues including reduced mechanical strength of the tendon postsurgery, increased risk of re-rupture, and persistent tendon adhesions, leading to joint dysfunction, delayed rehabilitation, and chronic pain (4, 5). These challenges largely stem from the tendon’s inherently low regenerative potential and the disruption of the neurovascular network following injury, which impairs orderly collagen remodeling and functional restoration.

Tendon healing is a complex, staged process involving inflammatory, proliferative, and remodeling phases (6). Effective recruitment of sensory nerve fibers (SeNFs) and their associated vascular networks to the injury site is now recognized as a critical biological determinant of successful repair (7, 8). Emerging evidence highlights that maintaining a favorable ratio of sensory nerves to sympathetic nerves in peritendinous tissue during the early healing phases optimizes outcomes (9). SeNFs form a multidimensional regulatory network by releasing various neuropeptides [such as calcitonin gene-related peptide (CGRP)]: In terms of vascularization, CGRP acts as a potent vasodilator, notably up-regulating vascular endothelial growth factor (VEGF) expression, promoting the directed growth of new blood vessels into ischemic injury areas, and improving early microcirculation and nutrient supply; at the level of tendon cell function, neuropeptides directly stimulate cell proliferation, migration, and the synthesis and secretion of key extracellular matrix components (especially type I collagen) by activating their membrane-specific receptors [such as tropomyosin receptor kinase A (TrkA) and neurokinin-1 receptor (NK1R)], thereby driving structural reconstruction. Conversely, excessive sympathetic nerve activity and norepinephrine release induce vasoconstriction and metabolic stress, impairing repair. Thus, establishing and activating SeNF signaling networks represents a key regulatory axis for intrinsic tendon regeneration—a process for which no effective clinical strategies currently exist.

Electrical stimulation (ES) as a nonpharmacological, precisely controllable physical intervention method for modulating neural activity demonstrates unique therapeutic potential (10, 11). Notably, ES has been shown to promote SeNF growth and neuropeptides, making it an ideal strategy for optimizing the “low sympathetic/high sensory” neural ratio and activating the neural-tendon repair axis (12, 13). This is primarily due to the fundamental differences between SeNFs and sympathetic nerve fibers (SyNFs) in terms of excitation thresholds, electrophysiological responses, nutrient factor dependency, and regenerative mechanisms. Under identical electrode-tissue conditions, large-diameter myelinated fibers (such as Aβ fibers primarily associated with sensory function) have a lower activation threshold than small-diameter unmyelinated C fibers. Therefore, low-intensity ES is more likely to preferentially activate SeNFs rather than SyNFs. This recruitment order has been demonstrated in animal and modeling studies, showing that larger-diameter fibers are typically activated earlier during ES (14, 15). In clinical and experimental contexts, “microcurrent” is generally defined as current of <1 mA, most commonly tens to hundreds of microamperes, which is sufficient to modulate neuronal excitability without thermal or electrochemical damage (16). At these intensities, sensory afferents exhibit robust responses, with downstream activation of phosphatidylinositol 3-kinase (PI3K)/Akt (protein kinase B) and, in many paradigms, extracellular signal–regulated kinase (ERK)/mitogen-activated protein kinase (MAPK) signaling that supports Schwann-cell migration, axon extension, and growth-associated protein expression (12, 17, 18). By contrast, selective activation of small-diameter C-fiber populations typically requires higher intensities and/or longer pulse durations; optimized paradigms for C-fiber recruitment use substantially greater charge than microcurrent levels, supporting the view that sympathetic C fibers show limited engagement under sub-milliampere stimulation (15, 19). This unique neuroselectivity makes ES an attractive strategy to restore neurovascular homeostasis and drive tendon early healing. However, the molecular mechanisms linking ES-mediated SeNFs activation to downstream processes such as angiogenesis and collagen remodeling remain poorly defined. For instance, intermittent theta-burst stimulation (iTBS) has been reported to activate signal transducer and activator of transcription 3 (STAT3), which is closely associated with nerve fiber growth in rats, yet the intrinsic regulatory machinery remains unclear (20, 21). Furthermore, clinical ES techniques typically rely on external power sources, which are impractical for deep tissues like tendons and are associated with infection risk, low compliance, and variable efficacy (22, 23). These limitations highlight the urgent need to develop implantable systems that avoid external power sources while enabling localized, dynamic neuromodulation.

Based on the core mechanism of ES promoting SeNF growth and its limitations in deep tissue applications (such as external power dependency issues), this study first focuses on revealing the specific molecular signaling networks mediated by ES that drive SeNF growth and its downstream repair effects. Through in-depth exploration, we identified the yes-associated protein 1 (YAP1)/phosphorylated STAT3 (pSTAT3)/neuropilin-1 (NRP1) signaling axis as playing a key role in ES-driven SeNF growth and neurotrophic factor expression. The activation of this signaling pathway serves as the crucial molecular basis for ES to optimize the “low sympathetic/high sensory” nerve ratio and coordinate downstream repair processes such as angiogenesis and collagen remodeling. Guided by this discovery, we designed a force-electricity–coupled dual-layer patch poly(vinylidene difluoride-trifluoroethylene) [P(VDF-TrFE)]/regenerated silk fibroin@poly(3,4-ethylenedioxythiophene):polystyrene sulfonate (RSF@P:P) that converts external ultrasound stimulation into therapeutic piezoelectric signals, thereby eliminating the reliance on external power sources. The tendon-facing P(VDF-TrFE) layer generates piezoelectric currents during movement, activating the YAP1/pSTAT3/NRP1 axis and promoting directed ingrowth of SeNFs and their accompanying vessels into the injury site. Simultaneously, piezoelectric stimulation triggers a gel-sol transition in the RSF@P:P hydrogel layer, creating a lubricating interface that reduces peritendinous adhesions. In vitro and in vivo studies confirm that this bioelectronic system synergistically enhances neurovascular regeneration while mitigating adhesion formation, culminating in improved structural and functional recovery of repaired tendons.

RESULTS

Neurovascular remodeling and STAT3-YAP1 axis activation in tendon repair

Initial histological evaluations of Achilles tendon injury (TI) at 2 weeks postsurgery revealed pronounced ingrowth of SeNFs (Fig. 1A) and vasculature (Fig. 1B) in the peritendinous region compared to sham-operated controls, consistent with prior observations of neurovascular involvement in tendon repair (7). To delineate the molecular mechanisms underlying the neurite outgrowth, we performed transcriptomic RNA sequencing (RNA-seq) coupled with expression profiling. Global heatmap analysis demonstrated distinct gene expression patterns between the sham group and TI group (fig. S1). Volcano plot analysis identified 3454 up-regulated and 3296 down-regulated genes following TI (Fig. 1C). Principal components analysis further confirmed intergroup transcriptional divergence, providing a robust framework for subsequent differential gene expression and clustering analyses (fig. S2). Gene ontology (GO) enrichment analysis highlighted the top 14 biological processes associated with neuronal architecture and function, including neuron part, neuron projection, and axon development (fig. S3). These findings further confirm the ingrowth of nerve fibers after TI.

The STAT family, a core component of transcriptional regulatory networks, has been demonstrated by recent studies to play critical roles in neurite outgrowth (24, 25). To elucidate the mechanisms underlying SeNF infiltration following TI, we first analyzed the STAT family based on differentially expressed genes (DEGs). The STAT family comprises critical transcription factors governing biological processes such as cell proliferation, differentiation, and neural repair. Comparative analysis revealed STAT3 as the most prominently altered member in the injured Achilles tendon group compared to sham-operated controls (Fig. 1D). Gene set enrichment analysis (GSEA) of cell growth pathways further identified STAT3 as a key transcriptional regulator implicated in neural growth modulation (Fig. 1E). Immunofluorescence staining corroborated these findings by showing markedly elevated pSTAT3 expression in the peritendinous sensory nerves of the TI group compared with the sham group (Fig. 1F).

To elucidate the molecular mechanisms by which STAT3 regulates neurite outgrowth, we established a STAT3-centric protein-protein interaction (PPI) network through STRING database analysis (Fig. 1G). Guided by previous reports highlighting the close functional association between STAT3 and YAP1 across multiple biological processes (26, 27), we then used molecular docking simulations to predict their specific binding interfaces and dynamic interactions. Cross-species conservation analysis of human homologs (used due to limited rat structural data) demonstrated high sequence conservation (fig. S4), with human STAT3-YAP1 docking achieving a score of −270.36 (91% confidence), indicative of stable binding (Fig. 1H). Immunoprecipitation (IP) assays biochemically confirmed this interaction, revealing physical complex formation (Fig. 1I). Complementary GSEA identified notable YAP1 expression modulation in injured versus control tendons (fig. S5A). Further quantitative immunofluorescence analysis revealed that YAP1 expression in peritendinous sensory nerves was markedly up-regulated after injury compared with the sham group (fig. S5B).

ES drives YAP1/pSTAT3 translocation and NRP1 up-regulation

To further elucidate the interaction between pSTAT3 and YAP1, we performed immunofluorescence colocalization analysis (fig. S6A) and quantitative protein assessment (fig. S6B) in nerve growth factor (NGF)–differentiated PC12 rat pheochromocytoma cells, a commonly used in vitro model for studying neurite growth–related signaling. The results revealed a high degree of spatial overlap between the two proteins, providing imaging-based evidence of their close association. Based on current research reports indicating that both STAT3 activation and ES can influence neural growth through promoting the release of neurotrophic factors (28, 29), we postulated a potential mechanistic link between these two modalities.

In subsequent experiments, we systematically investigated the effects of ES on pSTAT3 and YAP1 protein expression in dorsal root ganglion neurons (DRGn). Western blot analysis (Fig. 2, A and B) demonstrated that ES up-regulated the total and nuclear levels of pSTAT3 and YAP1 (figs. S7 and S8), whereas the total STAT3 levels remained unchanged. To identify potential pSTAT3 targets associated with nerve growth, we used databases such as hTFtarget, JASPAR, ENCODE, and MotifMap (Fig. 2, C to E), predicting targets including mitogen-activated protein kinase kinase kinase 13 (MAP3K13), NRP1, protein tyrosine phosphatase receptor type F (PTPRF), and glycogen synthase kinase 3 beta (GSK3B). These predictions were validated using quantitative reverse transcription polymerase chain reaction (qRT-PCR), which indicated that NRP1 showed the most pronounced up-regulation under ES (Fig. 2F). Subsequent Western blot analysis (Fig. 2G) and quantitative measurements (fig. S9) confirmed a statistically significant increase in NRP1 protein expression following ES.

Fig. 2. — (A) The total protein levels of YAP1, pSTAT3 Tyr⁷⁰⁵, and STAT3 with and without ES. β-actin was used as an internal control. n = 3. (B) The nuclear protein levels of YAP1 and pSTAT3 Tyr⁷⁰⁵ with and without ES. Histone H3 was used as an internal control. n = 3. (C) Intersection of neurite outgrowth–related genes across multiple databases. (D and E) Enrichment pathways of neurite outgrowth–related genes transcribed via pSTAT3 Tyr⁷⁰⁵, as predicted by GO and Kyoto Encyclopedia of Genes and Genomes (KEGG) analyses, respectively. (F) qRT-PCR results for the mRNA expression of neurite outgrowth–related genes transcribed via pSTAT3, including MAP3K13, NRP1, PTPRF, and GSK3B. n = 3. (G) Protein levels of NRP1 with and without ES. β-actin was used as an internal control. n = 3. (H) Predicted STAT3 binding motif sequence from the JASPAR database. (I) Dual-luciferase assays. n = 3. (J) Schematic representation of the NRP1 promoter region, highlighting the predicted STAT3 binding site (blue box) and the locations of forward (F) and reverse (R) primers used for ChIP-PCR. (K) ChIP-PCR analysis revealed that compared with the IgG control, the NRP1 promoter fragment was enriched in the chromatin precipitated by the anti-STAT3 antibody, confirming that STAT3 occupied the predicted site. n = 3. (L) Expression levels of CGRP in the supernatant of DRGn with and without ES. n = 8. n.s., not significant.

To determine whether STAT3 regulates the transcriptional activity of NRP1, we first used the JASPAR database to predict potential STAT3 binding motifs within the NRP1 promoter region (Fig. 2H) and generated their sequence logos. To definitively validate the STAT3 binding site, we performed site-directed mutagenesis on the putative STAT3 response element. Functional assays demonstrated that STAT3 overexpression significantly enhanced wild-type promoter activity by approximately threefold (P < 0.0001). In contrast, this activation effect was markedly attenuated in the mutant construct (P = 0.0103). The marked attenuation of promoter activation in the mutant construct confirms that the mutated region constitutes a critical functional STAT3 response element (fig. S10). Next, we mapped the NRP1 promoter region, indicating the predicted STAT3 binding site (blue box) and primer positions [forward (F)/reverse (R)] for chromatin immunoprecipitation (ChIP)–PCR (Fig. 2J). ChIP-PCR analysis confirmed that STAT3 specifically binds to the NRP1 promoter under endogenous conditions (Fig. 2K). Collectively, these findings demonstrate that STAT3 promotes NRP1 transcription through direct binding to its promoter. We focused on a representative neuropeptide secreted by DRGn—CGRP—to assess the effects of ES on neuropeptide release. ES markedly increased CGRP protein levels in DRGn cultures (Fig. 2L).

In DRGn, ES can induce the nuclear translocation of pSTAT3, activate YAP1, up-regulate the expression of NRP1, and increase the secretion of the representative neuropeptide CGRP. Prior studies have established STAT3 as a central regulator of axonal regeneration following DRG injury: The cyclin-dependent kinase 5 (Cdk5)–STAT3 pathway promotes regenerative responses (30), whereas STAT3 deficiency substantially impairs axonal regeneration in the peripheral nervous system (31). In parallel, YAP/transcriptional coactivator with PDZ-binding motif (TAZ) has been shown to play an essential role in Schwann cell–mediated myelin regeneration and nerve repair (32). Together with our findings, these results support a model in which ES activates the pSTAT3/YAP1/NRP1 signaling axis in DRGn, thereby facilitating the ingrowth of CGRP⁺ nerve fibers and contributing to functional repair.

ES counteracts shSTAT3/YAP1 suppression to restore NRP1/CGRP

Subsequently, we verified the effect of ES on the STAT3/YAP1/NRP1 signaling axis, as well as the specific roles of ES and the STAT3/YAP1/NRP1 signaling axis in the secretion of CGRP by DRGn (Fig. 3A). We first examined the regulatory effects of short hairpin RNA against STAT3 (shSTAT3) and ES on STAT3 and NRP1 protein expression. As shown in Fig. 3 (B and C), ES partially reversed the shSTAT3-mediated knockdown of pSTAT3 protein, consequently markedly up-regulating the NRP1 expression. Subsequently, functional rescue experiments were performed. Under conditions of ES and shSTAT3 treatment, NRP1 was overexpressed in DRGn, and the differences in CGRP secretion were evaluated by enzyme-linked immunosorbent assay (ELISA). In addition, the effects of DRGn supernatant on human umbilical vein endothelial cell (HUVEC) migration were examined. ELISA results showed that NRP1 overexpression increased CGRP secretion to ~1.56-fold compared with the group treated only with ES and shSTAT3 (fig. S11A). Meanwhile, treatment of HUVECs with the supernatant from NRP1-overexpressing DRGn modestly accelerated cell migration, further confirming that NRP1 overexpression may promote CGRP secretion in DRGn (fig. S11B). Similarly, ES counteracted the shYAP1-induced YAP1 suppression, which also led to increased NRP1 levels (Fig. 3, D and E). In subsequent cotransduction experiments with shSTAT3 and shYAP1 in DRGn, ES was observed to alleviate the combinatorial suppression of NRP1 expression induced by dual knockdown. Quantitative analysis revealed that shSTAT3 exerted a stronger inhibitory effect on NRP1 compared to shYAP1 (Fig. 3, F to I). Last, we examined the effects of shSTAT3, shYAP1, and ES on the nuclear protein levels of pSTAT3 and YAP1. The results demonstrated that ES partially reversed the reduced nuclear expression of pSTAT3 and YAP1 induced by shSTAT3 and shYAP1, respectively (fig. S12). Together, these findings indicate that ES up-regulates both total and nuclear protein levels of pSTAT3 and YAP1.

To investigate paracrine effects, we evaluated the conditioned media from differentially treated DRGn on HUVEC behavior (Fig. 3J). ELISA quantification showed that both shRNA treatments effectively reduced CGRP secretion, with shSTAT3 producing more pronounced reductions (Fig. 3, K and L). We subsequently performed a scratch wound healing assay (Fig. 3M) and quantified the data following methodologies established in previous literature. The results indicated that conditioned media from shRNA-treated cells impaired endothelial cell migration and morphogenesis, with shSTAT3 showing a stronger inhibitory effect. Moreover, ES partially reversed the impairment of cell migration capacity induced by either shSTAT3 or shYAP1 (fig. S13). Given the crucial role of neurite outgrowth in tendon regeneration, these findings suggest that both shSTAT3 and shYAP1 may impair tendon repair processes, while ES could potentially counteract these detrimental effects.

Synthesis and structural characterization of P(VDF-TrFE)/RSF@P:P

Leveraging these mechanistic discoveries, we developed a piezoelectric self-lubricating composite material [P(VDF-TrFE)/RSF@P:P] that synergistically enhances peritendinous CGRP⁺ nerve fiber growth through piezoelectric stimulation–activated neurotrophic signaling pathways to drive endogenous tissue repair, while its dynamically tunable lubrication interface generated via piezoelectric hydration effects effectively suppresses fibrotic scarring associated with exogenous healing. The fabrication procedures for the P(VDF-TrFE) membrane and P(VDF-TrFE)/RSF@P:P are illustrated in fig. S14. Scanning electron microscopy (SEM) analysis revealed that the P(VDF-TrFE) membrane exhibited a typical nanofiber structure, with uniform nanofiber diameters of ~700 nm. The P(VDF-TrFE)/RSF@P:P sample, due to the surface hydrogel coating, displayed the characteristic porous structure of the hydrogel (fig. S15A). Further examination of the cross-sectional morphology of P(VDF-TrFE)/RSF@P:P showed that the P(VDF-TrFE) nanofibers were uniformly encapsulated within the hydrogel matrix (fig. S15B). Energy-dispersive x-ray spectroscopy (EDS) results (fig. S16) revealed the elemental composition of P(VDF-TrFE), including C and F, while the surface of P(VDF-TrFE)/RSF@P:P showed the presence of Ca, N, and S elements, confirming the existence of RSF and P:P. These findings are consistent with the results obtained from x-ray photoelectron spectroscopy (XPS) (full spectrum analysis). Furthermore, the mechanical properties of the materials were characterized, including stress-strain curves and elastic modulus (fig. S17). The results showed that the elastic modulus of the P(VDF-TrFE) group and the P(VDF-TrFE)/RSF@P:P group were 0.317 ± 0.008 MPa and 0.327 ± 0.013 MPa, respectively, with no significant difference between them. This suggests that the RSF@P:P hydrogel did not affect the mechanical properties of P(VDF-TrFE).

Fourier transform infrared spectroscopy (FTIR) analysis confirmed the presence of P(VDF-TrFE) in the P(VDF-TrFE)/RSF@P:P composite material, with characteristic peaks observed at 842 and 881 cm⁻¹ (33). Furthermore, the analysis identified β-sheet domains within the silk fibroin, characterized by peaks at 1515 and 1637 cm⁻¹ (34), alongside poly(3,4-ethylenedioxythiophene) (PEDOT), which exhibited a peak at 982 cm⁻¹ (fig. S18). The prominent bands observed at 1515 and 1637 cm⁻¹ were respectively assigned to C─N─H bending vibrations and C═O stretching vibrations (35). The peak at 982 cm⁻¹ is indicative of the symmetric stretching vibration of C─O─C in PEDOT (36), whereas the peaks at 842 and 881 cm⁻¹ were respectively attributed to the symmetric stretching vibration of CF₂ and the asymmetric bending vibration of CF₂ in P(VDF-TrFE) (37). The x-ray diffraction (XRD) pattern of the composite material, as depicted in fig. S19, revealed a pronounced peak at approximately 2θ = 19.7°, signifying the β-crystal structure inherent to the silk fibroin gel. In the XPS analysis, the O 1s peak appeared at ~530 to 533 eV, with chemical shifts corresponding to different oxygen-containing functional groups. The component at ~531.0 eV was assigned to the carbonyl (C═O) chemical state, whereas the component at ~532.6 eV was attributed to ether/hydroxyl (C─O) groups. The O 1s peak positions and relative area ratios were consistent between the two material groups, indicating no substantial differences in their surface oxygen-containing chemical environments (fig. S20).

The physicochemical characterization of P(VDF-TrFE)/RSF@P:P

We evaluated the piezoelectric properties of the P(VDF-TrFE)/RSF@P:P by monitoring the changes in voltage and current generated under different ultrasound intensities (Fig. 4A). The results demonstrated that both voltage and current increased with the rise in ultrasound intensity, confirming the material’s piezoelectric effect (Fig. 4, B and C). Under ultrasound stimulation at 1 W/cm², the piezoelectric patch generated an open-circuit voltage of 0.37 V and a short-circuit current of 0.93 μA. A 14-day repeated ultrasound-activation durability test was then performed, during which the piezoelectric output was monitored daily. After 14 days of cyclic stimulation, the patch maintained a stable performance, exhibiting an open-circuit voltage of 0.34 V and a short-circuit current of 0.96 μA. These results indicate no statistically significant difference in piezoelectric output over time, confirming the long-term fatigue durability and stability of the patch under repeated ultrasound activation (fig. S21).

Fig. 4. — (A) Schematic diagram of material piezoelectric property testing. (B) Output short-circuit current waveforms of P(VDF-TrFE) membrane under different ultrasound intensities. (C) Output open-circuit voltage waveforms of P(VDF-TrFE) membrane under different ultrasound intensities. (D) The rheological properties reveal the angular frequency behavior of RSF@P:P hydrogel. (E) The rheological properties reveal the oscillation strain γ behavior of RSF@P:P hydrogel. (F and G) Contact angles of P(VDF-TrFE) membrane and P(VDF-TrFE)/RSF@P:P. (H to K) Surface roughness measurement. (L and M) Quantitative measurement of scratch heights for two groups. (N) Comparison of friction coefficients between PLA/RSF@P:P group and P(VDF-TrFE)/RSF@P:P group. (O) Comparison of wear volume between PLA/RSF@P:P group and P(VDF-TrFE)/RSF@P:P group. n = 6.

Rheological analysis was conducted using time-scan tests, and the results of angular frequency and oscillatory strain indicated that the storage modulus (G′) was higher than the loss modulus (G″), suggesting that the material was in a gel state (Fig. 4, D and E). We also measured the contact angle properties of P(VDF-TrFE) and P(VDF-TrFE)/RSF@P:P (Fig. 4, F and G). The results show that P(VDF-TrFE) exhibits strong hydrophobicity, while P(VDF-TrFE)/RSF@P:P demonstrates strong hydrophilicity. The lubricating properties of the material were tested using a friction and wear testing machine. Under the same mechanical loading conditions, compared to the nonpiezoelectric material polylactic acid (PLA)/RSF@P:P, the P(VDF-TrFE)/RSF@P:P material exhibited visibly lighter lubrication tracks, indicating superior lubricating performance (Fig. 4, H and I). Subsequently, the surface roughness of the PLA/RSF@P:P and P(VDF-TrFE)/RSF@P:P groups was measured using laser confocal microscopy (Fig. 4, J and K). The results showed that the roughness of the PLA/RSF@P:P group was significantly higher than that of the P(VDF-TrFE)/RSF@P:P group. Quantitative analysis of the cross-sections of the wear scars was performed, and the scar height was plotted (Fig. 4, L and M). The results revealed that the depth of the friction surface in the P(VDF-TrFE)/RSF@P:P group was significantly lower than that in the PLA/RSF@P:P group, indicating reduced friction resistance and confirming the lubricating effect under piezoelectric stimulation. This further substantiated that the piezoelectric effect of P(VDF-TrFE) contributes to enhanced lubrication. A detailed analysis of the lubricating effect demonstrated that the coefficient of friction (COF) of P(VDF-TrFE)/RSF@P:P remained consistently around 0.07 ± 0.002, significantly lower than that of PLA/RSF@P:P (0.11 ± 0.003) (Fig. 4N). In addition, the wear volume of the P(VDF-TrFE)/RSF@P:P sample (1.29 × 10⁵ ± 11,910 μm³) was significantly lower than that of the PLA/RSF@P:P sample (3.02 × 10⁵ ± 12,982 μm³) (Fig. 4O). This is attributed to the piezoelectric properties of P(VDF-TrFE), which generate electrical signals under mechanical loading, inducing a gel-sol transition in the silk fibroin and forming a liquid layer on the hydrogel surface, thereby enhancing lubrication.

The assessment of biocompatibility and cell-cell cross-talk in vitro

The live/dead staining of PC12 cells was initially assessed, as shown in fig. S22. Across all groups, the proportion of dead cells remained consistently low at 1, 3, and 5 days postculture (with representative images shown for day 5 only), indicating the high biocompatibility of the P(VDF-TrFE)/RSF@P:P composite. In addition, cell proliferation was evaluated using the Cell Counting Kit-8 (CCK-8) assay at the same time points. Figure S23 shows no significant differences in proliferation among the control, P(VDF-TrFE), RSF@P:P hydrogel, and P(VDF-TrFE)/RSF@P:P groups, suggesting that PC12 cells maintained their proliferative capacity throughout the culture period. These results collectively highlight the excellent biocompatibility of the P(VDF-TrFE)/RSF@P:P composite.

DRGn were cultured on different materials and subjected to ultrasound to induce piezoelectric activation, after which the conditioned media were collected and applied to HUVEC cultures to assess their growth responses (Fig. 5A). DRGn were seeded in the bottom of culture dishes containing different materials and subjected to ultrasound to induce piezoelectric effects. The conditioned medium from the different dishes was collected and used to culture HUVECs. Subsequently, the growth of HUVECs in each experimental group was observed. Furthermore, the impact of piezoelectric stimulation on PC12 cell proliferation was evaluated. PC12 cells were subjected to different treatments and stained with 5-ethynyl-2′-deoxyuridine (EdU; with PLA serving as a nonpiezoelectric control). The results indicated that piezoelectric stimulation significantly enhanced cell proliferation (fig. S24).

Fig. 5. — (A) Schematic showing DRGn cultured on P(VDF-TrFE)/RSF@P:P and subjected to ultrasound stimulation; the resulting conditioned medium was applied to HUVECs, and HUVECs were not directly subjected to ultrasound treatment. (B) Scratch wound healing assay performed on HUVECs treated with conditioned medium from DRGn cultures (DRGn were exposed to ultrasound; HUVECs were not directly subjected to ultrasound). n = 3. (C) EdU incorporation assays visualizing cellular proliferation across treatment conditions. n = 3. (D) Quantification of EdU-positive cell rate (%) in HUVECs under different treatment conditions. n = 3.

Subsequently, we evaluated the growth of HUVECs cocultured with different groups of DRGn. The results revealed that HUVECs cocultured with piezoelectrically stimulated DRGn exhibited significantly faster migration (Fig. 5C) and proliferation (Fig. 5D) compared to the other groups. This effect is likely due to the piezoelectric stimulation promoting DRGn growth, leading to increased secretion of growth factors, which in turn enhanced HUVEC growth. We also assessed the effect of each group on the tube formation ability of HUVECs (fig. S25A). The results showed that the P(VDF-TrFE)/RSF@P:P group exhibited superior branch points (fig. S25B) and capillary length (fig. S25C) compared to the other groups, further confirming that coculturing with DRGn treated with P(VDF-TrFE)/RSF@P:P promoted the growth of HUVECs.

Last, we evaluated the lubricating effect of P(VDF-TrFE)/RSF@P:P in vitro by applying P(VDF-TrFE) to RSF@P:P and gelatin methacryloyl (GelMA) hydrogels and assessing its impact on cell adhesion. CCK-8 assay results indicated that P(VDF-TrFE)/RSF@P:P effectively inhibited cell adhesion, with the optical density (OD) value being approximately half that of the other groups (fig. S26). These findings demonstrate that under piezoelectric conditions, RSF@P:P effectively reduces fibroblast adhesion in vitro, and its impact on peritendinous adhesions in vivo will be presented in subsequent results.

In vivo assessment of P(VDF-TrFE)/RSF@P:P in tendon repair

This study systematically validated the therapeutic efficacy of P(VDF-TrFE)/RSF@P:P material in Achilles tendon repair through a multimodal evaluation framework. As illustrated in Fig. 6A, standardized Achilles TI models were established with uniform assessments conducted at designated time points. The postoperative evaluation system encompassed three dimensions: microstructural characterization (SEM), functional assessment (gait analysis), and histopathological examination (38).

Fig. 6. — (A) Timeline of the animal experiments. (B) Representative images and quantitative data from SEM, with the diameters of 20 randomly selected tendon fibers measured in each image. Calculations were performed using ImageJ. (C and D) A rose plot of tendon fibers in TI and P(VDF-TrFE)/RSF@P:P groups. (E) Bar chart results of fiber diameter for each group. (F and G) Footprints and gaits were analyzed using AFI analysis at 4 and 8 weeks after surgery. n = 6. (H and I) Representative images of H&E staining and the density of cell fibers stained with eosin. Calculations were performed using ImageJ (n = 4, biologically independent samples). (J to L) Representative images and quantitative data from Masson staining, including CVF and collagen fiber area. n = 6. (M to P) Immunohistochemical staining for Col I and Col III in the regenerated tendon at 8 weeks postoperatively. n = 6. S, skin; T, Achilles tendon.

At the microstructural level, SEM revealed markedly improved collagen fiber alignment in the treatment group at 8 weeks postoperation. Compared to other groups, the P(VDF-TrFE)/RSF@P:P group exhibited thicker collagen fibers with superior spatial organization (Fig. 6B). Quantitative analyses using fiber diameter histograms and orientation angle rose plots confirmed a 22.9% increase in mean collagen fiber diameter (168 ± 34 nm) compared to the injury group (Fig. 6E), accompanied by more concentrated fiber orientation distributions (Fig. 6, C and D), indicative of enhanced fiber remodeling. Functional evaluation through gait analysis (Fig. 6F) demonstrated distinct kinematic patterns: Healthy preoperative rats maintained minimal heel contact during ambulation, whereas postinjury rats at 4 weeks exhibited compensatory heel-ground contact. By 8 weeks posttreatment, the P(VDF-TrFE)/RSF@P:P group showed substantial functional recovery, evidenced by normalized heel contact area that approached physiological patterns compared to the TI group (Fig. 6G). Histopathological analyses of repair sites through hematoxylin and eosin (H&E) staining (Fig. 6H) and Masson’s trichrome staining (Fig. 6J) provided structural validation. Quantitative evaluations revealed statistically significant increases in eosin-stained cellular fiber density (Fig. 6I), collagen fiber density (Fig. 6K), and fiber cross-sectional area (Fig. 6L) in the treatment group compared to controls, corroborating the material’s proregenerative effects. Immunohistochemical quantification further substantiated these findings, with the P(VDF-TrFE)/RSF@P:P group exhibiting elevated expression levels of collagen I (Col I) and collagen III (Col III) compared to both TI and PLA/RSF@P:P groups (Fig. 6, M to P). Notably, immunohistochemical staining (fig. S27A) and quantitative analyses (fig. S27, B and C) demonstrated significantly higher scleraxis (SCX) expression in the treatment group, consistent with established tendon regeneration markers. The results of immunofluorescence staining showed that P(VDF-TrFE)/RSF@P:P promoted the ingrowth of peritendinous blood vessels and SeNFs (fig. S28). Last, we evaluated the impact of NRP1 on tendon repair. The results demonstrated that shNRP1-mediated knockdown markedly impaired tendon healing, as evidenced by histopathological analysis showing disorganized tissue architecture, pronounced inflammatory infiltration, and excessive scar tissue deposition in H&E-stained sections (fig. S29A). Furthermore, the SCX staining intensity was reduced compared with the TI group (fig. S29B). Collectively, these findings confirm the beneficial role of NRP1 in promoting tendon regeneration.

Subsequently, we evaluated the anti-adhesion efficacy of P(VDF-TrFE)/RSF@P:P in peritendinous tissue repair. Macroscopic observations demonstrated distinct differences among treatment groups: Both TI and PLA/RSF@P:P groups exhibited substantial adhesion formation, as evidenced by resistance encountered during forceps penetration between the repaired tendon and surrounding tissues (fig. S30A). In notable contrast, the P(VDF-TrFE)/RSF@P:P group showed minimal tissue adherence, permitting effortless blunt dissection of the tendon interface. These qualitative findings were quantitatively validated through gross adhesion scoring, which revealed statistically significant reduction in adhesion severity for P(VDF-TrFE)/RSF@P:P compared to both control groups (fig. S30B). Histopathological analyses provided complementary evidence at the microstructural level. H&E (fig. S31A) and Masson’s trichrome staining (fig. S31B) consistently demonstrated preserved tissue planes between the repair site and adjacent structures in the P(VDF-TrFE)/RSF@P:P group, whereas TI and PLA/RSF@P:P specimens showed complete obliteration of anatomical boundaries with dense fibrotic adhesions. Semiquantitative histological scoring systematically confirmed these morphological observations (fig. S31C), with P(VDF-TrFE)/RSF@P:P achieving significantly lower adhesion scores than comparator groups.

Collectively, these multimodal assessments establish that P(VDF-TrFE)/RSF@P:P effectively mitigates posttraumatic peritendinous adhesion formation. This antifibrotic property is particularly advantageous for tendon healing, as preservation of anatomical gliding surfaces is critical for restoring normal biomechanical function during rehabilitation.

P(VDF-TrFE)/RSF@P:P promotes tendon regeneration in Bama minipigs

To better mimic the reparative effects of P(VDF-TrFE)/RSF@P:P on injured human tendons, this study investigated their impact on tendon regeneration in a large animal model, the Bama minipig (Fig. 7, A and B). SEM revealed that the Achilles tendon fibers in the P(VDF-TrFE)/RSF@P:P group exhibited a more regular arrangement (Fig. 7C). Quantitative analysis further showed that the tendon fiber diameter in the P(VDF-TrFE)/RSF@P:P group was significantly greater than that in the TI and PLA/RSF@P:P groups, demonstrating the promotive effect of piezoelectric materials on tendon healing in Bama minipig (Fig. 7, D to F). In addition, H&E staining indicated that the density of cell fibers stained with eosin was markedly higher in the piezoelectric material group compared to the other two groups (Fig. 7, G and H). Furthermore, Masson’s trichrome staining and subsequent quantitative analysis revealed that the collagen fiber area and collagen volume fraction (CVF) were markedly superior in the piezoelectric group relative to the TI and PLA/RSF@P:P groups (Fig. 7, I to K). Last, immunofluorescence staining of tissue sections revealed that the P(VDF-TrFE)/RSF@P:P group exhibited clearly increased levels of SCX protein expression within the tendon (Fig. 7, L to N) compared to other experimental groups. These findings collectively confirm the promotive effects of piezoelectric materials on the healing of injured tendons in Bama minipigs.

Fig. 7. — (A) Timeline of Bama minipig in vivo experimental procedures. (B) Repair of damaged Achilles tendon using P(VDF-TrFE)/RSF@P:P. (C, D, E and G) Representative images and quantitative data from SEM, with the diameters of 20 randomly selected tendon fibers measured in each image. Calculations were performed using ImageJ. (F) Representative images of H&E staining. (H) The density of cell fibers stained with eosin. Calculations were performed using ImageJ (n = 4, biologically independent samples). (I) Representative images of Masson staining. (J and K) Quantitative data of CVF and collagen fiber area from Masson staining. n = 4. (L) Immunofluorescence staining for SCX (red) after TI at 4 months postoperatively (DAPI: blue). (M and N) Quantitative analysis of immunofluorescence staining of SCX. n = 3. (O) Blood routine and biochemical analysis results for evaluating the biosafety of P(VDF-TrFE)/RSF@P:P. (P) Representative H&E staining images of major organs.

The biocompatibility of P(VDF-TrFE)/RSF@P:P was evaluated through a comprehensive in vivo safety assessment in Bama minipigs. Hematological and biochemical analyses revealed no significant abnormalities, with all parameters remaining within normal physiological ranges, indicating unaffected liver and kidney function as well as metabolic status (Fig. 7O). Histopathological examination of the heart, liver, spleen, lungs, and kidneys via H&E staining showed intact tissue architecture, normal cellular morphology, and no evident pathological changes such as inflammation, edema, degeneration, or necrosis (Fig. 7P). These collective findings demonstrate the good biosafety of P(VDF-TrFE)/RSF@P:P in vivo, supporting its potential for clinical translation.

DISCUSSION

Our study revealed pronounced neurovascular remodeling during tendon repair and demonstrated its close association with activation of the YAP1-STAT3 axis. Further mechanistic investigations showed that ES activates the YAP1/pSTAT3/NRP1 pathway. CGRP⁺ SeNFs exhibited parallel growth with peritendinous angiogenesis, while transcriptomic analysis, protein interaction networks, and coimmunoprecipitation collectively supported a mechanistic model in which ES promotes YAP1-mediated nuclear translocation of pSTAT3, thereby transcriptionally up-regulating NRP1 expression. This pathway was further validated in primary DRGn, suggesting that ES enhances tendon repair by activating sensory neurons and driving angiogenesis, thus providing a clear molecular basis for its therapeutic effects (Fig. 8).

Fig. 8. — (I) ES enhances the interaction between YAP1 and STAT3, promotes STAT3 nuclear translocation and transcriptional activity, and thereby up-regulates the STAT3 target gene NRP1. (II) Preparation of P(VDF-TrFE)/RSF@P:P and its gel-sol transition. (**III**) In vivo application of P(VDF-TrFE)/RSF@P:P.

At the translational level, this mechanistic insight informed the design of a piezoelectric-lubricating patch [P(VDF-TrFE)/RSF@P:P]. The inner layer comprises P(VDF-TrFE) nanofibers that, when ultrasonically activated, generate localized bioelectric signals to sustain activation of the YAP1/pSTAT3/NRP1 axis. The outer layer, an RSF@P:P hydrogel, undergoes reversible gel-sol transitions under piezoelectric stimulation, thereby reducing peritendinous adhesions. In an in vivo Achilles TI model, external ultrasound elicited robust piezoelectric activation of the patch, effectively restoring collagen alignment and accelerating gait recovery.

In this study, although the PEDOT:poly(styrenesulfonate) (PSS) component itself is not biodegradable, it is highly biocompatible and is fully encapsulated within a biodegradable RSF matrix (39). As the RSF matrix gradually degrades in vivo, the encapsulated PEDOT:PSS nanostructures are released in minute quantities—a process whose biosafety is supported by previous long-term biological studies on PEDOT:PSS (40, 41). Moreover, our biosafety data from the Bama minipig model demonstrated no significant inflammation or organ toxicity after 4 months of implantation, providing strong experimental evidence supporting this safe clearance pathway.

Despite these promising outcomes, our study has certain limitations. Mechanistic validation was primarily conducted at the transcriptomic level, and further investigation using single-cell spatial transcriptomics, particularly in dorsal root ganglia, would enable a more precise understanding of the cell-specific responses to ES. This “dual-target” strategy—enhancing regenerative signals while inhibiting adhesions—provides a feasible translational pathway for improving tendon surgery outcomes and holds promise for expansion into other soft tissue repair fields.

MATERIALS AND METHODS

Fabrication of P(VDF-TrFE)/RSF@P:P

In this study, we used a fully encapsulated configuration rather than a simple surface coating. The RSF@P:P hydrogel was applied through a swelling-infiltration and embedding approach, resulting in complete envelopment of the P(VDF-TrFE) electrospun membrane.

For the P(VDF-TrFE) electrospun membrane, the P(VDF-TrFE) copolymer (Piezotech FC30, Piezotech S.A.S.) was dissolved in a mixed solvent of N,N-dimethylformamide (DMF; lot no. 33120, Sigma-Aldrich) and acetone (6:4 ratio) with stirring until the polymer was completely dissolved. The P(VDF-TrFE) solution (15 wt %) was loaded into a syringe, a metal needle was attached, and the electrospinning process was initiated with a feeding rate of ~1.0 ml h⁻¹, ensuring the formation of a Taylor cone as a criterion for successful spinning. After electrospinning, the P(VDF-TrFE) membrane was placed in a 60°C environment for ~20 min to remove residual solvent.

RSF@P:P hydrogel:silk fibroin was prepared as previously described (10) and dissolved with CaCl₂ in deionized water. PEDOT:PSS (lot no. 739324, Sigma-Aldrich) was mixed with glutaraldehyde and gradually added to the silk fibroin solution under stirring. In this system, PEDOT provides conductivity, PSS serves as a dopant/hydrophilic group, and Ca²⁺ coordinates with PSS sulfonic groups to form ionic cross-links. Meanwhile, glutaraldehyde reacts with free amino groups of silk fibroin to form Schiff bases, enhancing molecular interactions and promoting β-sheet formation. The electrospun P(VDF-TrFE) membrane was then coated with RSF@P:P hydrogel to obtain the composite material.

Fabrication of PLA electrospun membrane

For the PLA electrospun membrane, PLA (Jinan Daigang Co., Jinan, China) was dissolved in a mixed solvent of DMF (lot no. 33120, Sigma-Aldrich) and chloroform at a ratio of 3:7, stirring until the polymer was completely dissolved. The 10 wt % PLA solution was loaded into a syringe, and the electrospinning process was initiated with a feeding rate of ~0.5 to 1.0 ml h⁻¹, using the formation of a Taylor cone as the criterion for successful spinning.

Functionalization of P(VDF-TrFE)/RSF@P:P

To facilitate initial cell attachment, all membranes were subjected to brief oxygen plasma activation [100-W power, 20-SCCM (standard cubic centimeter per minute) O₂ flow, 20-mtorr pressure, and a 30-s exposure] prior to cell seeding (42, 43). This step was applied uniformly to all groups and was not part of the mechanistic intervention. No additional chemical grafting or ligand immobilization was performed.

Characterization of P(VDF-TrFE)/RSF@P:P

The microstructures of P(VDF-TrFE)/RSF@P:P were examined by field-emission SEM (400 Model, USA) after Au/Pd coating, and elemental composition was analyzed by EDS. FTIR (Nicolet iS20, Thermo Fisher Scientific), XRD (Ultima IV, Rigaku; 10° to 80° 2θ), and XPS (ESCALAB 250XI, Thermo Fisher Scientific) were used for structural characterization. Electrical output was assessed under ultrasound stimulation (probe area: 5 cm²; UT1041, Contemporary) by recording current and voltage. Rheological properties were measured at 37° ± 1°C using a HAAKE MARS3 rheometer (cone-plate, 1°, 60 mm).

Mechanical and lubrication performance testing

Rectangular specimens with dimensions of 30 mm by 5 mm were fabricated for tensile testing using a universal testing machine (Instron 5969). Prior to mechanical testing, all samples were presoaked in phosphate-buffered saline (PBS; pH 7.4) at 37°C for 24 hours to simulate physiological wet conditions. The gauge length was set to 10 mm, and the crosshead speed was 5 mm/min. Stress-strain curves were continuously recorded, and the Young’s modulus was determined from the linear elastic region of each curve.

To further assess surface tribological properties, ball-on-disk friction tests were conducted using a tribometer (UMT-2, Bruker). Samples were punched into 20-mm-diameter disks and prehydrated in PBS at 37°C for 24 hours prior to testing. A stainless steel ball was used as the counterface under a normal load of 1 N, with a sliding speed of 10 mm/s, stroke length of 5 mm, and a total test duration of 10 min. The COF was continuously recorded throughout the test, and the average COF during the steady-state stage was calculated. The morphology of wear tracks was examined using a laser scanning confocal microscope (Keyence VK-X1000), and the wear volume was quantified via three-dimensional surface profilometry analysis.

Cell lines and culture conditions

The cell lines used in this study [HUVEC, PC12, and human embryonic kidney (HEK) 293T cells] were obtained from the National Collection of Authenticated Cell Lines (Chinese Academy of Sciences). The specific identifiers are as follows: HUVEC (catalog no. SCSP-5330; CSTR: 19375.09.3101HUMSCSP5330), PC12 (catalog no. TCR 9; CSTR: 19375.09.3101RATTCR9), and HEK293T (catalog no. SCSP-502; CSTR: 19375.09.3101HUMSCSP502). The cells were maintained in Dulbecco’s modified Eagle’s medium (Gibco) supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% penicillin/streptomycin [penicillin (100 U/ml) and streptomycin (100 μg/ml)].

The pLV3-U6-Stat3(rat)-shRNA1-CopGFP-Puro and pLV3-U6-Yap1(rat)-shRNA3-CopGFP-Puro plasmids were purchased from Miaoling Bioscience (Wuhan, China) and used to generate the corresponding lentiviral constructs for transducing DRGn. Both lentiviruses were applied at a multiplicity of infection of 25, which is consistent with previously reported studies (44).

Biocompatibility evaluation in vitro

Cell viability was assessed using CCK-8 (Beyotime, Shanghai, China) and live/dead cell assays (45). The CCK-8 assay was conducted in 96-well plates over 1, 3, and 5 days according to the manufacturer’s instructions. OD at 450 nm was measured using a microplate reader (Leica Microsystems, Wetzlar, Germany). In addition, a live/dead assay was performed in 12-well plates using a working solution containing calcein-AM and propidium iodide (46).

Cell migration assay

In brief, cells were seeded in a six-well plate and allowed to reach 80 to 90% confluence. A sterile pipette tip was used to gently create a uniform straight line in the cell monolayer. Initial (0-hour) and 24-hour images of the scratch were captured under a microscope, and the changes in the width of the wound area at different time points were compared to assess the cell migration ability.

EdU staining

Cells were seeded in a six-well plate and cultured until 70 to 80% confluence. EdU (lot no. ST067, Beyotime) labeling was then performed, and cells were further incubated for 12 hours. The cells were subsequently fixed and permeabilized, and EdU detection was carried out. After staining the nuclei with Hoechst 33342, the cells were observed under a microscope.

qRT-PCR

Total RNA was extracted using TRIzol reagent (lot no. R0016, Beyotime), and cDNA was synthesized through reverse transcription with the Prime Script RT Reagent Kit (lot no. RR047A, TaKaRa, Japan). The primers used were as follows: Map3k13, forward: 5′-ACACACCGGGAGAGGTTTTG-3′ and reverse: 5′-GTCTTCAGGTCCGTTCGTGT-3′; Nrp1, forward: 5′-CACCCGGTCTTCCATAAGGG-3′ and reverse: 5′-GAGAACATTCGGGCCCTCTC -3′; Gsk3b, forward: 5′-CCACAGCAGCCTCAGATACT-3′ and reverse: 5′-AACGTGACCAGTGTTGCTGA-3′; Ptprf, forward: 5′-CTCCCAGCGCTTTGAGGTAA-3′ and reverse: 5′-GAACGCAGCTGCTTGATACG -3′; SCX, forward: 5′-AACCCTGTCGTGTTCATGCT-3′ and reverse: 5′- GTCACGGTCTTTGCTCAACTT-3′; Col I, forward: 5′-CATGTTCAGCTTTGTGGACCTC-3′ and reverse: 5′-CAGCTGACTTCAGGGATGTCT-3′; Col III, forward: 5′-CCTCCCGGCAACAATGGTAA-3′ and reverse: 5′-GGGACCTGGATGTCCACTTG-3′; and glyceraldehyde-3-phosphate dehydrogenase (GAPDH), forward: 5′-GAAGCTGGTCATCAACGGGA-3′ and reverse: 5′-CGACATACTCAGCACCAGCA-3′.

Western blot analysis

Proteins were extracted, separated by SDS–polyacrylamide gel electrophoresis (SDS-PAGE), and transferred onto membranes. After blocking (Quick Block, Beyotime), membranes were incubated overnight at 4°C with primary antibodies against β-actin, STAT3, pSTAT3 (Tyr⁷⁰⁵), YAP1, NRP1, and histone H3 (Abmart or Cell Signaling Technology, CST), followed by horseradish peroxidase–conjugated secondary antibodies for 1 hour at room temperature. Signals were visualized by chemiluminescence.

IP assay

The IP experiment was conducted in primary DRGn. Protein A/G magnetic beads were incubated with anti-STAT3 antibody and anti–immunoglobulin G (IgG) antibody at 4°C for 2 hours. Protein lysates from DRGn were then incubated with the antibody-conjugated beads overnight at 4°C. The resulting complexes were washed four times with phosphate-buffered saline with Tween-20 (PBST), after which 1× SDS-PAGE sample buffer was added to the IP complexes and boiled. The prepared protein samples were subjected to SDS-PAGE for separation, followed by Western blot analysis using standard protocols.

Dual-luciferase assays

Because HEK293T cells exhibit high transfection efficiency and low background noise, this study also used the HEK293T cell line, which is commonly used for luciferase reporter assays (47). HEK293T cells were seeded in 24-well plates and transfected at 50 to 70% confluence. The NRP1 promoter region [−2000 to 0 base pairs (bp)] was synthesized and cloned into pGL3-Enhancer (pGL3-NRP1-WT). The expression plasmid pCDNA-3.1-Stat3(rat)-3×Myc, pCDNA-3.1-3×Myc (Vector) was purchased from Miaolingbio (Wuhan, China), and pRL-SV40-C (Beyotime, Jiangsu, China; no. D2768) was used as the internal control. Plasmids were cotransfected into HEK293T cells for 24 hours, with a total DNA amount of ~500 ng per well. We further generated a mutant construct of the NRP1 promoter region, pGL3-NRP1-Mut (−1936 to −1928 bp), in which the core STAT3-binding sequence was completely replaced with a poly-adenine tract. The specific sequence information for the target site is as follows: wild-type sequence, TTCCCAACA; mutant sequence, TAAAAAAAA. This substitution was designed to specifically disrupt the binding capacity of the transcription factor STAT3. After 24 hours of transfection, the cells were lysed using lysis buffer. The lysates were then transferred to a detection plate, and Firefly luciferase activity was measured by adding the Firefly luciferase substrate. Subsequently, Renilla luciferase activity was measured by adding the Renilla luciferase substrate. The luminescence signals were detected using a luminometer (Synergy, BioTek Instruments).

Chromatin immunoprecipitation

First, formaldehyde cross-linking was used to bind proteins to DNA in the cells. After cross-linking, the reaction was terminated with glycine (final concentration of 0.125 M), and the cells were washed with PBS. Next, the cells were lysed with lysis buffer for 15 min, and chromatin was fragmented by sonication. Then, the chromatin was precleared with protein A/G agarose beads, typically incubated for 1 hour. The chromatin was incubated with specific antibodies against the target protein STAT3 (lot no. 12640S), usually overnight at 4°C. Protein A/G beads were added, and the mixture was incubated for 1 to 2 hours to precipitate the antibody-bound chromatin. After IP, the samples were washed multiple times (including low salt, high salt, LiCl, TE buffers, etc.) to remove nonspecifically bound materials. Reverse cross-linking is then performed using an elution buffer containing SDS and NaCl to release the DNA from the protein-DNA complexes. The DNA was further purified through phenol-chloroform extraction or using a purification kit. Last, the enrichment of the target DNA regions was analyzed by PCR, ensuring the reliability of the results. IgG controls and input DNA were used to validate the specificity and accuracy of the experiment.

The complete NRP1 promoter sequence is shown in fig. S32, and the ChIP-PCR targeted regions are marked in red. Specific primers were designed for the ~2000-bp region upstream of the rat NRP1 promoter (including the STAT3 predicted binding sequence TTCCCAACA): forward primer, 5′-AGAGATGGCTCAGTGGTTAA-3′; reverse primer, 5′-TACACACAATTAAAATAATT-3′. The PCR amplification signal is standardized with input DNA to correct the amount of chromatin. Normal IgG was used as a negative control to verify the specificity of the binding and the reliability of the experiment.

Enzyme-linked immunosorbent assay

Culture supernatants from treated DRGn were collected, and concentrations of CGRP (lot no. E-EL-R0135, Elabscience, China) were quantified using ELISA kits according to the manufacturer’s protocols.

Immunofluorescence staining

The cells were fixed with paraformaldehyde and then incubated with a blocking solution containing Triton X-100 (lot no. P0096, Beyotime) for 1 hour. The samples were subsequently incubated overnight at 4°C with diluted primary antibodies targeting the proteins of interest (pSTAT3 Tyr⁷⁰⁵, YAP1, and NRP1). After three washes with PBS, the samples were incubated for 1 hour at room temperature in the dark with secondary antibody. Last, the samples were observed using laser scanning confocal microscopy (LSM800, Zeiss).

Tenocyte isolation, expansion, and culture

The Achilles tendons of the rats were dissected and immediately placed in a 75% alcohol solution. After the removal of the tendon epitenon, the tissues were washed three times with Hank’s solution. The entire tendon segment was then digested in a mixed enzyme solution containing 0.25% trypsin (lot no. C0201, Beyotime) and 0.1% collagenase I (lot no. ST2294, Beyotime) at 37°C for 30 min. After digestion was stopped with Hank’s solution, the tendon was cut into 1.0- to 2.0-mm³ pieces and further digested in the same enzyme solution at 37°C in a 5% CO₂ incubator for 1.0 to 2.0 hours. During digestion, the flask was gently shaken every 30 min to aid in digestion and tissue dispersion, and the process was monitored under an inverted microscope. When most tissue pieces were loosened and many cells had been released, digestion was terminated by adding F-12 medium containing 20% FBS. The digested mixture was filtered to remove tissue debris, and the filtrate was centrifuged at 1000 rpm for 10 min. The supernatant was discarded, and the pellet was washed once with F-12 medium containing 20% FBS (lot no. 04-001-1ACS, BI). The resulting cell suspension was prepared, and the cell count was adjusted to 5 × 10⁵ cells, which were seeded into 25-ml culture flasks for further cultivation.

DRGn cultures

Purified primary DRGn were enzymatically dissociated from L1 to L6 ganglia of adult male Sprague-Dawley rats (200 to 250 g body weight) following established isolation protocols (48). The neuronal cultures were subsequently maintained in serum-free medium supplemented with 1% (v/v) N2 formulation (lot no. 17502048, Gibco) and recombinant NGF (50 ng/ml; lot no. 450-34, PeproTech) for 7 days to achieve functional maturation prior to experimental interventions. Based on previously reported parameters in the literature for piezoelectric stimulation induced by ultrasound-activated materials in vitro, the cells were subjected to 10 s of ultrasound activation at 0.5 W/cm² (49).

Rat and Bama minipig TI model

All animal procedures were approved by the institutional review committee of Shanghai Jiao Tong University [SYXK (Hu)2021–0028], and those involving Bama minipig was approved by the Institutional Animal Care and Use Committee under approval number IACUC25-0204.

Male Sprague-Dawley rats aged 6 to 8 weeks were anesthetized. The Achilles tendon was transected, and a modified Kessler suture was applied. In the P(VDF-TrFE)/RSF@P:P group, the P(VDF-TrFE)/RSF@P:P material was wrapped around the TI site, while in the injury group, the tendon was sutured directly in layers.

For the Bama minipig, anesthesia was administered, and the hindlimb tendons were exposed. A partial transverse TI (~1 cm) was created, followed by a modified Kessler suture. The P(VDF-TrFE)/RSF@P:P material was applied to the injury site, and the wound was sutured and closed. After 4 months, the animals were euthanized, and the tendons were harvested for further analysis.

Previous studies have shown that peptidergic sensory neuropeptides (CGRP) peak at 2 to 3 weeks postinjury, remain elevated during 2 to 6 weeks, and then progressively decline over 4 to 16 weeks as remodeling proceeds. In parallel, autonomic neuropeptides [vasoactive intestinal peptide, (VIP); neuropeptide Y, (NPY)] begin to increase at approximately week 4, indicating the onset of the remodeling phase (50). Based on these findings, we restricted ultrasound intervention to the first 4 weeks postinjury, thereby targeting the sensory nerve–dominant proregenerative window and avoiding late-phase neurovascular overactivation. In vivo ultrasound treatment was administered in cycles of two consecutive treatment days followed by one rest day; each session lasted 20 min with parameters set at 1-MHz frequency, 0.5-W/cm² power output, and 50% duty cycle (51).

Achilles function index

Achilles function index (AFI) analysis was performed at 4 and 8 weeks postoperatively as previously described (n = 6) (52). Briefly, the hindpaws were dipped in black ink, and the rats were allowed to walk down a narrow channel, making footprints on white paper. AFI was calculated according to the classic formula: AFI = 74 × PLF + 161 × TSF + 48 × ITF – 5 (53). PLF denotes the print length factor, the relative difference in footprint length (heel–to–longest-toe distance) between the injured and healthy sides; TSF denotes the toe spread factor, the relative difference in the maximal distance between the first and fifth toes; and ITF denotes the intermediary toe spread factor, the relative difference in the distance between the second and fourth toes. They are calculated as PLF = (EPL − NPL)/NPL, TSF = (ETS − NTS)/NTS, and ITF = (EIT − NIT)/NIT, where E and N refer to the injured (experimental) and healthy (sham) sides, respectively.

Histology and immunohistochemistry

After gross evaluation, the specimens were preserved in 4% paraformaldehyde. The samples underwent gradient dehydration and decalcification before being embedded in paraffin and sectioned at a thickness of ~5 μm along the sagittal plane, ensuring that the repair site was centered. Sections were stained with H&E and Masson’s trichrome following the supplier’s protocols to evaluate tissue morphology (10). Immunohistochemical staining for Col I and Col III was performed in accordance with the manufacturer’s instructions.

Biosafety evaluation in vivo

Blood routine and biochemical analyses were performed to evaluate the biological safety of the material in Bama minipig (n = 3). Samples were collected 4 months postoperatively and analyzed by Jiangsu Kebiao Medical Technology Group Co., Ltd. In parallel, H&E staining of major organs, including the heart, liver, spleen, lungs, and kidneys, was conducted following the aforementioned protocol (10).

Statistical analyses

Statistical analyses were performed using GraphPad Prism 10.0 (GraphPad Software, CA, USA). Normality of data distribution was assessed by the Shapiro-Wilk test, and homogeneity of variance was evaluated by Levene’s test. For comparisons between two groups, a two-tailed unpaired Student’s t test was used when data met normality assumptions; otherwise, the Mann-Whitney U test was applied. For comparisons among multiple groups, one-way analysis of variance (ANOVA) was performed followed by Tukey’s or Bonferroni’s post hoc test as appropriate.

Differential expression analysis of RNA-seq data was conducted using the DESeq2 package. Multiple testing was corrected by the Benjamini-Hochberg method, with an adjusted P < 0.05 considered significant. GSEA was performed using the Broad Institute pipeline, with the significance threshold set at a false discovery rate of <0.25 according to recommended standards.

The number of replicates for each experiment was as follows: animal experiments were performed with n = 6 biological replicates (independent animal samples per group), in vitro cell experiments with n = 3 biological replicates (independent cell cultures), and RNA-seq analysis with n = 3 biological replicates. The number of replicates for all immunofluorescence, molecular, and functional assays is indicated in the corresponding figure legends. All statistical tests were two-sided, and P < 0.05 was considered statistically significant.

Acknowledgments

We thank J. Jin for assistance with the mechanical performance testing.

Funding:

This work received financial support from the following: the National Key Research and Development Program of China (2024YFC2418105), the National Natural Science Foundation of China (grant nos. 82172408, 82572709, 82425035, and 9236810338), the Shanghai Jiao Tong University Medical College “Two-Hundred Talent” Program (no. 20191829), the Program of Shanghai Academic/Technology Research Leader (no. 22XD1422600), Shanghai Municipal Health Commission (no. 2022YQ073), the Shanghai “Medical Star” Young Medical Talent Training Funding Program (Outstanding Youth), the Shanghai Health Commission Health Industry Research Project (Excellent Project), the Shanghai Municipal Health Commission Research Project (no. 20234Z0008)c Fundamental Research Funds for the Central Universities (YG2024ZD18), the Shanghai Science and Technology Industry High-Quality Development Program (25S11901600), the Shanghai Key Technology Research and Development Program (25CL2900400), the China Postdoctoral Science Foundation (2024M762062), and the Shanghai Municipal Natural Science Foundation Program for Basic Research (25ZR1402425).

Author contributions:

Conceptualization: J.W., Fan Wang, J.L., and W.C. Methodology: Y.X., Fan Wang, and Z.L. Investigation: J.W. and X.L. Formal analysis: P.Z. Data curation: J.L. Visualization: Z.L. and J.W. Validation: Fan Wang. Project administration: Fei Wang. Funding acquisition: Fei Wang, W.C., and S.L. Supervision: Fei Wang, W.C., and S.L. Writing—original draft: J.W. and Fan Wang. Writing—review and editing: Fei Wang, J.W., and W.C.

Competing interests:

The authors declare that they have no competing interests.

Data, code, and materials availability:

All data and code needed to evaluate and reproduce the results in the paper are present in the paper and/or the Supplementary Materials. This study did not generate new materials.

Supplementary Materials

This PDF file includes:

Figs. S1 to S32

Uncropped Western blots

sciadv.aec1272_sm.pdf^{(11.8MB, pdf)}

REFERENCES

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figs. S1 to S32

Uncropped Western blots

sciadv.aec1272_sm.pdf^{(11.8MB, pdf)}

Data Availability Statement

All data and code needed to evaluate and reproduce the results in the paper are present in the paper and/or the Supplementary Materials. This study did not generate new materials.

[R1] 1.Rajpar I., Tomlinson R. E., Function of peripheral nerves in the development and healing of tendon and bone. Semin. Cell Dev. Biol. 123, 48–56 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R2] 2.Liu Y., Xie W., Tang Z., Tan Z., He Y., Luo J., Wang X., A reconfigurable integrated smart device for real-time monitoring and synergistic treatment of rheumatoid arthritis. Sci. Adv. 10, eadj0604 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R3] 3.Traweger A., Scott A., Kjaer M., Wezenbeek E., Silva R. S., Kennedy J. G., Butler J. J., Gomez-Florit M., Gomes M. E., Snedeker J. G., Dakin S. G., Wildemann B., Achilles tendinopathy. Nat. Rev. Dis. Primers 11, 20 (2025). [DOI] [PubMed] [Google Scholar]

[R4] 4.Maffulli N., Peretti G. M., Treatment decisions for acute Achilles tendon ruptures. Lancet 395, 397–398 (2020). [DOI] [PubMed] [Google Scholar]

[R5] 5.Kracun D., Gorlach A., Snedeker J. G., Buschmann J., Reactive oxygen species in tendon injury and repair. Redox Biol. 81, 103568 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R6] 6.Darrieutort-Laffite C., Blanchard F., Soslowsky L. J., Le Goff B., Biology and physiology of tendon healing. Joint Bone Spine 91, 105696 (2024). [DOI] [PubMed] [Google Scholar]

[R7] 7.Cherief M., Xu J., Li Z., Tower R. J., Ramesh S., Qin Q., Gomez-Salazar M., Yea J.-H., Lee S., Negri S., Xu M., Price T., Kendal A. R., Fan C.-M., Clemens T. L., Levi B., James A. W., TrkA-mediated sensory innervation of injured mouse tendon supports tendon sheath progenitor cell expansion and tendon repair. Sci. Transl. Med. 15, eade4619 (2023). [DOI] [PubMed] [Google Scholar]

[R8] 8.Silva M., Ferreira F. N., Alves N. M., Paiva M. C., Biodegradable polymer nanocomposites for ligament/tendon tissue engineering. J. Nanobiotechnology 18, 23 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R9] 9.Ackermann P. W., Li J., Lundeberg T., Kreicbergs A., Neuronal plasticity in relation to nociception and healing of rat Achilles tendon. J. Orthop. Res. 21, 432–441 (2003). [DOI] [PubMed] [Google Scholar]

[R10] 10.Wang J.-Y., Yuan Y., Zhang S.-Y., Lu S.-Y., Han G.-J., Bian M.-X., Huang L., Meng D.-H., Su D.-H., Xiao L., Xiao Y., Zhang J., Gong N.-J., Jiang L.-B., Remodeling of the intra-conduit inflammatory microenvironment to improve peripheral nerve regeneration with a neuromechanical matching protein-based conduit. Adv. Sci. 11, e2302988 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R11] 11.Kennedy V. Jr., Long M. D., Walters J., Adewuyi A. A., Franz C. K., Applications of advances in therapeutic electrical stimulation techniques and technologies in precision peripheral nerve repair: A narrative review. Adv. Technol. Neurosci. 2, 97–101 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R12] 12.Geremia N. M., Gordon T., Brushart T. M., Al-Majed A. A., Verge V. M., Electrical stimulation promotes sensory neuron regeneration and growth-associated gene expression. Exp. Neurol. 205, 347–359 (2007). [DOI] [PubMed] [Google Scholar]

[R13] 13.Brushart T. M., Jari R., Verge V., Rohde C., Gordon T., Electrical stimulation restores the specificity of sensory axon regeneration. Exp. Neurol. 194, 221–229 (2005). [DOI] [PubMed] [Google Scholar]

[R14] 14.Hennings K., Kamavuako E. N., Farina D., The recruitment order of electrically activated motor neurons investigated with a novel collision technique. Clin. Neurophysiol. 118, 283–291 (2007). [DOI] [PubMed] [Google Scholar]

[R15] 15.Schneider T., Filip J., Soares S., Sohns K., Carr R., Rukwied R., Schmelz M., Optimized electrical stimulation of C-nociceptors in humans based on the chronaxie of porcine C-fibers. J. Pain 24, 957–969 (2023). [DOI] [PubMed] [Google Scholar]

[R16] 16.Kolimechkov S., Seijo M., Swaine I., Thirkell J., Colado J. C., Naclerio F., Physiological effects of microcurrent and its application for maximising acute responses and chronic adaptations to exercise. Eur. J. Appl. Physiol. 123, 451–465 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R17] 17.Al-Majed A. A., Brushart T. M., Gordon T., Electrical stimulation accelerates and increases expression of BDNF and trkB mRNA in regenerating rat femoral motoneurons. Eur. J. Neurosci. 12, 4381–4390 (2000). [PubMed] [Google Scholar]

[R18] 18.Zhong H., Xing C., Zhou M., Jia Z., Liu S., Zhu S., Li B., Yang H., Ma H., Wang L., Zhu R., Qu Z., Ning G., Alternating current stimulation promotes neurite outgrowth and plasticity in neurons through activation of the PI3K/AKT signaling pathway. Acta Biochim. Biophys. Sin. 55, 1718–1729 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R19] 19.Johnson M., Transcutaneous electrical nerve stimulation: Mechanisms, clinical application and evidence. Rev. Pain 1, 7–11 (2007). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R20] 20.Zareen N., Yung H., Kaczetow W., Glattstein A., Mazalkova E., Alexander H., Chen L., Parra L. C., Martin J. H., Molecular signaling predicts corticospinal axon growth state and muscle response plasticity induced by neuromodulation. Proc. Natl. Acad. Sci. U.S.A. 121, e2408508121 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R21] 21.Qiu J., Cafferty W. B., McMahon S. B., Thompson S. W., Conditioning injury-induced spinal axon regeneration requires signal transducer and activator of transcription 3 activation. J. Neurosci. 25, 1645–1653 (2005). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R22] 22.Burton A., Wang Z., Song D., Tran S., Hanna J., Ahmad D., Bakall J., Clausen D., Anderson J., Peralta R., Sandepudi K., Benedetto A., Yang E., Basrai D., Miller L. E., Tresch M. C., Gutruf P., Fully implanted battery-free high power platform for chronic spinal and muscular functional electrical stimulation. Nat. Commun. 14, 7887 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R23] 23.Marquez-Chin C., Popovic M. R., Functional electrical stimulation therapy for restoration of motor function after spinal cord injury and stroke: A review. Biomed. Eng. Online 19, 34 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R24] 24.He F., Ge W., Martinowich K., Becker-Catania S., Coskun V., Zhu W., Wu H., Castro D., Guillemot F., Fan G., de Vellis J., Sun Y. E., A positive autoregulatory loop of Jak-STAT signaling controls the onset of astrogliogenesis. Nat. Neurosci. 8, 616–625 (2005). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R25] 25.Sun F., Park K. K., Belin S., Wang D., Lu T., Chen G., Zhang K., Yeung C., Feng G., Yankner B. A., He Z., Sustained axon regeneration induced by co-deletion of PTEN and SOCS3. Nature 480, 372–375 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R26] 26.He L., Pratt H., Gao M., Wei F., Weng Z., Struhl K., YAP and TAZ are transcriptional co-activators of AP-1 proteins and STAT3 during breast cellular transformation. eLife 10, (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R27] 27.Igarashi R., Oda M., Okada R., Yano T., Takahashi S., Pastuhov S., Matano M., Masuda N., Togasaki K., Ohta Y., Sato S., Hishiki T., Suematsu M., Itoh M., Fujii M., Sato T., Generation of human adult hepatocyte organoids with metabolic functions. Nature 641, 1248–1257 (2025). [DOI] [PubMed] [Google Scholar]

[R28] 28.Menacho I. P., Koster R., van der Sloot A. M., Quax W. J., Osinga J., van der Sluis T., Hollema H., Burzynski G. M., Gimm O., Buys C. H. C. M., Eggen B. J. L., Hofstra R. M. W., RET-familial medullary thyroid carcinoma mutants Y791F and S891A activate a Src/JAK/STAT3 pathway, independent of glial cell line-derived neurotrophic factor. Cancer Res. 65, 1729–1737 (2005). [DOI] [PubMed] [Google Scholar]

[R29] 29.Fan W., Yang X., Hu X., Huang R., Shi H., Liu G., A novel conductive microtubule hydrogel for electrical stimulation of chronic wounds based on biological electrical wires. J. Nanobiotechnology 22, 258 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R30] 30.Hwang J., Namgung U., Cdk5 phosphorylation of STAT3 in dorsal root ganglion neurons is involved in promoting axonal regeneration after peripheral nerve injury. Int. Neurourol. J. 24, S19–S27 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R31] 31.Bareyre F. M., Garzorz N., Lang C., Misgeld T., Büning H., Kerschensteiner M., In vivo imaging reveals a phase-specific role of STAT3 during central and peripheral nervous system axon regeneration. Proc. Natl. Acad. Sci. U.S.A. 108, 6282–6287 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R32] 32.Grove M., Lee H., Zhao H., Son Y.-J., Axon-dependent expression of YAP/TAZ mediates Schwann cell remyelination but not proliferation after nerve injury. eLife 9, e50138 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R33] 33.Jiang Y., Gong L., Hu X., Zhao Y., Chen H., Feng L., Zhang D., Aligned P(VDF-TrFE) nanofibers for enhanced piezoelectric directional strain sensing. Polymers 10, 364 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R34] 34.Salama A., Cellulose/silk fibroin assisted calcium phosphate growth: Novel biocomposite for dye adsorption. Int. J. Biol. Macromol. 165, 1970–1977 (2020). [DOI] [PubMed] [Google Scholar]

[R35] 35.Li R., Li Y., Wu Y., Zhao Y., Chen H., Yuan Y., Xu K., Zhang H., Lu Y., Wang J., Li X., Jia X., Xiao J., Heparin-poloxamer thermosensitive hydrogel loaded with bFGF and NGF enhances peripheral nerve regeneration in diabetic rats. Biomaterials 168, 24–37 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R36] 36.Sun B., Wu T., Wang J., Li D., Wang J., Gao Q., Bhutto M. A., El-Hamshary H., Al-Deyab S. S., Mo X., Polypyrrole-coated poly(l-lactic acid-co-epsilon-caprolactone)/silk fibroin nanofibrous membranes promoting neural cell proliferation and differentiation with electrical stimulation. J. Mater. Chem. B 4, 6670–6679 (2016). [DOI] [PubMed] [Google Scholar]

[R37] 37.Yaseen H. M. A., Park S., Enhanced power generation by piezoelectric P(VDF-TrFE)/rGO nanocomposite thin film. Nanomaterials 13, 860 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

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PERMALINK

Activation of the YAP1/pSTAT3/NRP1 axis in peritendinous sensory nerves promotes tendon healing

Jiayi Wang

Fan Wang

Jingwen Liu

Yao Xiao

Zhaoyang Li

Xiaonan Liu

Peilin Zhang

Fei Wang

Wenguo Cui

Shen Liu

Roles

Abstract

INTRODUCTION

RESULTS

Neurovascular remodeling and STAT3-YAP1 axis activation in tendon repair

Fig. 1. Neurovascular remodeling and STAT3-YAP1 axis activation in tendon repair.

ES drives YAP1/pSTAT3 translocation and NRP1 up-regulation

Fig. 2. ES enhances nuclear translocation of YAP1/pSTAT3 and subsequently up-regulates NRP1 transcription.

ES counteracts shSTAT3/YAP1 suppression to restore NRP1/CGRP

Fig. 3. Validation of ES regulating NRP1 levels via YAP1/pSTAT3 nuclear translocation to influence neurite outgrowth.

Synthesis and structural characterization of P(VDF-TrFE)/RSF@P:P

The physicochemical characterization of P(VDF-TrFE)/RSF@P:P

Fig. 4. Electrical and tribological characterization data of P(VDF-TrFE)/RSF@P:P.

The assessment of biocompatibility and cell-cell cross-talk in vitro

Fig. 5. Ultrasound-activated P(VDF-TrFE)/RSF@P:P promotes HUVECs migration and proliferation via DRGn-conditioned medium.

In vivo assessment of P(VDF-TrFE)/RSF@P:P in tendon repair

Fig. 6. Promotive effects of P(VDF-TrFE)/RSF@P:P on tendon recovery in rats.

P(VDF-TrFE)/RSF@P:P promotes tendon regeneration in Bama minipigs

Fig. 7. P(VDF-TrFE)/RSF@P:P enhance tendon healing in Bama minipig.

DISCUSSION

Fig. 8. Schematic illustration of the P(VDF-TrFE)/RSF@P:P promoting tendon regeneration.

MATERIALS AND METHODS

Fabrication of P(VDF-TrFE)/RSF@P:P

Fabrication of PLA electrospun membrane

Functionalization of P(VDF-TrFE)/RSF@P:P

Characterization of P(VDF-TrFE)/RSF@P:P

Mechanical and lubrication performance testing

Cell lines and culture conditions

Biocompatibility evaluation in vitro

Cell migration assay

EdU staining

qRT-PCR

Western blot analysis

IP assay

Dual-luciferase assays

Chromatin immunoprecipitation

Enzyme-linked immunosorbent assay

Immunofluorescence staining

Tenocyte isolation, expansion, and culture

DRGn cultures

Rat and Bama minipig TI model

Achilles function index

Histology and immunohistochemistry

Biosafety evaluation in vivo

Statistical analyses

Acknowledgments

Funding:

Author contributions:

Competing interests:

Data, code, and materials availability:

Supplementary Materials

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REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

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