ADSCs were isolated from the inguinal adipose tissue of 4-week-old Sprague-Dawley rats according to the protocol described in previous research [14], and the rats were euthanized in the same manner as in Section 2.6. Briefly, adipose tissue was harvested under sterile conditions, minced into approximately 1-mm³ fragments, and enzymatically digested with collagenase. The digested suspension was filtered, centrifuged, and the resulting cell pellet was resuspended and cultured in complete culture medium. Cells were maintained at 37 °C in a humidified atmosphere containing 5% CO₂, and medium was refreshed every 2–3 days. ADSCs at passage 3 were used for all subsequent experiments. All primary ADSCs were validated for cell identity by surface marker analysis using flow cytometry and were confirmed to be mycoplasma-negative, ensuring the reliability and reproducibility of the experimental results.

To suppress IGF1 expression, ADSCs were transduced with lentiviral vectors encoding short hairpin RNA (shRNA) targeting rat Igf1 (GenePharma, Shanghai, China). The shRNA sequence targeting Igf1 was GGTGGATGCTCTTCAGTTC, while a non-targeting sequence (ACTTACGCTGAGTACTTCG) served as the negative control. Lentiviral transduction was performed according to the manufacturer’s instructions. Knockdown efficiency was confirmed at both mRNA and protein levels.

Exosomes were isolated from ADSC-conditioned medium using a standard differential ultracentrifugation protocol. Briefly, culture supernatants were sequentially centrifuged at 1000 $\times$g for 10 min to remove cells and debris, 10,000 $\times$g for 30 min to eliminate large extracellular vesicles, and 100,000 $\times$g for 70 min to pellet exosomes. The resulting pellet was resuspended in phosphate-buffered saline (PBS) and further purified by filtration through a 0.22-µm membrane. Exosome morphology was examined by transmission electron microscopy (TEM), and particle size distribution and concentration were determined using nanoparticle tracking analysis (NTA). Expression of canonical exosomal markers (Tsg101, CD63, and CD9) was assessed by Western blotting. Exosomal protein concentration was measured using a bicinchoninic acid (BCA) protein assay according to the manufacturer’s instructions.

Exosome purity was evaluated by calculating the particle-to-protein ratio (particles/µg protein).

To evaluate whether IGF1 knockdown affects ADSC viability, a Cell Counting Kit-8 (CCK-8) assay was performed. Control ADSCs and IGF1-knockdown ADSCs (shIGF1) in the logarithmic growth phase were seeded into 96-well plates at a density of 5 $\times$ 10³ cell/well in 100 µL complete culture medium containing 10% fetal bovine serum. Cell viability was assessed on days 1, 2, 3, and 4 after seeding by adding CCK-8 reagent according to the manufacturer’s instructions. Absorbance was measured at 450 nm using a microplate reader, and relative cell viability was calculated for each time point.

To evaluate whether systemically administered exosomes can reach the tendon–bone injury site, exosome biodistribution was assessed using DiI fluorescent labeling. Purified ADSC-derived exosomes were incubated with the lipophilic fluorescent dye DiI at a final concentration of 5 µM for 30 min at 4 °C in the dark. To remove unbound dye, the labeled exosomes were ultracentrifuged at 100,000 $\times$g for 70 min and resuspended in sterile PBS.

DiI-labeled exosomes were administered to rats with rotator cuff tear via tail-vein injection. Three days after injection, rats were euthanized in the same manner as in Section 2.6, and tendon tissues from the repair site were harvested, embedded, sectioned, and examined under a fluorescence microscope to detect DiI fluorescence signals.

The rat rotator cuff tear and repair model were established following the surgical procedure described in previous research [15, 16]. A total of 42 adult female Sprague–Dawley rats (4–5 months old, 258–552 g) were used. Animals were randomly assigned to experimental groups using a computer-generated random number table. Investigators performing outcome assessments were blinded to group allocation. General anesthesia was induced with 2–3% isoflurane in 100% oxygen. A transverse incision was made lateral to the deltoid muscle to expose the rotator cuff complex. The supraspinatus tendon was sharply detached from the greater tuberosity, and the footprint was gently debrided to simulate clinical tendon avulsion prior to repair.

Postoperatively, 200 µg of ADSC-derived exosomes suspended in 200 µL PBS were administered via tail vein injection immediately after surgery (day 0), on days 3 and 7, and once weekly thereafter until the designated endpoints. Animals were euthanized at 1 or 8 weeks post-surgery under deep anesthesia by intraperitoneal injection of sodium pentobarbital (150 mg/kg). Death was confirmed by the absence of respiration, corneal reflex, and heartbeat.

At 1 week post-surgery, tendon tissues from the repair site were harvested and homogenized in RIPA lysis buffer. Protein samples were separated by SDS–PAGE and transferred to PVDF membranes. Membranes were incubated with primary antibodies against IGF1 (1:2000, #ab322659, Abcam, Cambridge, MA, USA), VEGFA (1:1000, #ab214424, Abcam), CD31 (1:1500, #ab222783, Abcam), interleukin (IL)-1$\beta$ (1:1000, Ag10295, Proteintech, Wuhan, China), IL-18 (1:1000, 30587-1-AP, Proteintech), ACTA2 (1:1500, #19245, Cell Signaling Technology, Danvers, MA, USA), and GAPDH (#1:2000, 60004-1-Ig, Proteintech) (used as an internal control), followed by HRP-conjugated secondary antibodies. Protein bands were visualized using enhanced chemiluminescence.

qRT-PCR was performed using SYBR Green Master Mix (#4344463, Thermo Fisher Scientific, Waltham, MA, USA) on a CFX96 Real-Time PCR System (Bio-Rad, Hercules, CA, USA). GAPDH was used as the internal reference gene. The primer sequences used were as follows:

GAPDH:

F: 5^′-TCAAGAAGGTGGTGAAGCAG-3^′;

R: 5^′-GGTGGAAGAGTGGGAGTTGC-3^′.

Igf1:

F: 5^′-CCTGCTTGCTCACCTTTACC-3^′;

R: 5^′-GGTAGCTCAGGCATGTCCAG-3^′.

Vegfa:

F: 5^′-GAGAGGTACAGTGCTGCCCT-3^′;

R: 5^′-CACACAGGACGGCTTGAAGA-3^′.

Acta2:

F: 5^′-GACCTTGAGAAGAGTTACGAGTTG-3^′;

R: 5^′-TAGAGAGACAGCACGATGGG-3^′.

Cd31:

F: 5^′-GCTGGTGCTGTTCTTCCTGT-3^′;

R: 5^′-AGGTGCCATCCAGGTACTTG-3^′.

Il18:

F: 5^′-TGCCATGTCAGAAGACTCTGC-3^′;

R: 5^′-TGGGTCACAGCCAGTTCTTC-3^′.

Il1b:

F: 5^′-TGCAGCTGGAGAGTGTGGAT-3^′;

R: 5^′-TGTCGTTGCTTGGTTCTCCT-3^′.

At 8 weeks post-surgery, shoulder joint specimens were fixed in 4% paraformaldehyde, decalcified, embedded in paraffin, and sectioned at 5-µm thickness. Hematoxylin and eosin (H&E) staining was used to assess overall tissue morphology, while Safranin O–Fast Green staining was performed to evaluate fibrocartilage formation at the tendon–bone interface. For immunohistochemistry, sections were incubated with primary antibodies against VEGF, followed by secondary antibodies and DAB visualization to assess neovascularization.

Micro-computed tomography (micro-CT) analysis was conducted to assess bone structure and mineralization at the greater tuberosity [17]. Following scanning, 3D reconstruction was used to evaluate bone volume parameters and trabecular architecture. For biomechanical testing, the humerus-tendon-scapula complex was harvested and subjected to uniaxial tensile loading to determine maximum load to failure and stiffness, thereby assessing the mechanical integrity of the tendon-to-bone repair.

All data are presented as mean $\pm$ standard deviation (SD). Normality was assessed using the Shapiro–Wilk test. As data were normally distributed but exhibited unequal variances among groups, Brown–Forsythe ANOVA and Welch’s correction were applied where appropriate. For pairwise comparisons, Welch’s t-test was used. Actual F values, t values, effect sizes, and sample sizes (n) are reported in the Results section and corresponding figure legends. Statistical analyses were performed using GraphPad Prism version 9.0 (GraphPad Software, San Diego, CA, USA), and p 0.05 was considered statistically significant.

To verify whether IGF1 knockdown affected ADSC viability, a CCK-8 assay was performed to assess cell viability. From day 1 to day 4 of culture, the relative cell viability of control ADSCs and shIGF1-transduced ADSCs increased in parallel, and no significant differences were observed at any time point (day 4: t = 0.87, df = 10, p = 0.483, Cohen’s d = 0.30; Supplementary Fig. 1). These results demonstrate that IGF1 knockdown did not impair ADSC viability.

To determine whether suppression of Igf1 in ADSCs altered IGF1 expression and its incorporation into secreted exosomes, ADSCs were transduced with lentiviral shRNA targeting Igf1. Quantitative RT-PCR and Western blot analyses confirmed a significant reduction in Igf1 mRNA and IGF1 protein levels in shIGF1-transduced ADSCs compared with controls (Fig. 1A–C; p 0.01, Brown–Forsythe ANOVA; F = 14.25, df = 2.6, partial $\eta$² = 0.83 for Fig. 1A; F = 11.89, df = 2.6, partial $\eta$² = 0.80 for Fig. 1C).

Exosomes isolated from control and shIGF1 ADSCs were subsequently characterized. Western blotting further confirmed the presence of canonical exosomal markers TSG101, CD63, and CD9, with no detectable differences between control and shIGF1-derived exosomes (Fig. 1D,E), indicating that Igf1 knockdown did not affect exosome biogenesis or structural integrity. To further evaluate whether Igf1 knockdown affects exosome yield and purity, particle concentration and total protein content of isolated exosomes were quantified, and the particle-to-protein ratio was calculated. No significant differences were observed between control ADSC-derived exosomes (EXO) and Igf1-knockdown ADSC-derived exosomes (EXO^shIGF1) in particle concentration, protein concentration, or particle-to-protein ratio (Supplementary Fig. 2A–C). These results further indicate that Igf1 knockdown does not affect basic exosome biogenesis, yield, or purity, but specifically reduces IGF1 cargo within the exosomes.

Importantly, analysis of exosomal cargo revealed a substantial reduction in IGF1 content following Igf1 knockdown. qRT-PCR demonstrated significantly decreased Igf1 mRNA levels in exosomes derived from shIGF1-transduced ADSCs compared with controls (Fig. 1F; p 0.01, independent-samples t-test; t = 4.68, df = 4, Cohen’s d = 1.91). Consistently, Western blot analysis showed a pronounced reduction in IGF1 protein levels in shIGF1-derived exosomes, while the exosomal protein Alix remained unchanged and served as a loading control (Fig. 1G,H; independent-samples t-test; t = 4.02, df = 4, Cohen’s d = 1.65). Collectively, these results demonstrate that lentiviral-mediated Igf1 knockdown specifically reduces IGF1 expression in ADSCs and concomitantly decreases IGF1 mRNA and protein content in their secreted exosomes, without altering ADSC viability or fundamental exosome biogenesis. These findings establish a robust foundation for subsequent functional analyses of IGF1-deficient exosomes in tendon–bone healing.

Given that exosomes were administered systemically in this study, it was first necessary to determine whether intravenously delivered exosomes could reach the tendon–bone injury site. Accordingly, DiI-labeled ADSC-derived exosomes were injected via the tail vein. Distinct DiI fluorescence signals were detected in tendon tissues three days after injection, indicating that systemically delivered exosomes are able to successfully reach and persist at the injury site (Supplementary Fig. 3).

To assess the structural and functional outcomes of rotator cuff repair, micro-CT and biomechanical analyses were performed at 8 weeks post-surgery. Micro-CT analysis demonstrated that treatment with exosomes derived from control ADSCs (EXO group) significantly improved bone microarchitecture at the tendon–bone interface compared with the untreated model group (Supplementary Fig. 4). Specifically, bone mineral density (BMD), bone volume fraction (BV/TV), and trabecular spacing (Tb.Sp) were all significantly improved in the EXO group (Fig. 2A–C; Brown–Forsythe ANOVA: F = 20.37, df = 2.15, partial $\eta$² = 0.73 for BMD; F = 16.52, df = 2.15, partial $\eta$² = 0.68 for BV/TV; F = 18.45, df = 2.15, partial $\eta$² = 0.71 for Tb.Sp). In contrast, these osteogenic benefits were markedly attenuated when exosomes were derived from Igf1-knockdown ADSCs. Compared with the EXO group, the shIGF1-EXO group exhibited significantly reduced BMD and BV/TV, along with increased Tb.Sp, indicating compromised bone regeneration at the tendon–bone interface.

Consistent with the micro-CT findings, biomechanical testing revealed that the maximum load to failure and interfacial stiffness were significantly higher in the EXO group than in the untreated model group, confirming enhanced mechanical integration of the repaired tendon (Fig. 2D,E; Brown–Forsythe ANOVA: F = 22.68, df = 2.15, partial $\eta$² = 0.75 for maximum load; F = 17.93, df = 2.15, partial $\eta$² = 0.70 for stiffness). However, both biomechanical parameters were significantly reduced in the shIGF1-EXO group relative to the EXO group. Taken together, these results indicate that IGF1 deficiency substantially compromises the ability of ADSC-derived exosomes to promote bone regeneration and mechanical restoration at the tendon–bone interface, supporting a critical role for exosomal IGF1 in mediating the therapeutic efficacy of exosome-based treatment in rotator cuff repair.

To further evaluate histological remodeling at the tendon–bone interface following rotator cuff repair, H&E staining and SO–FG staining were performed on specimens harvested 8 weeks post-surgery. Significantly lower scores were revealed in the EXO group compared with the model group, indicating reduced inflammatory features and more orderly fibrocartilage structure (Fig. 3A,B; Brown–Forsythe ANOVA: F = 16.82, df = 2.15, partial $\eta$² = 0.69). Consistently, SO–FG staining demonstrated a significantly greater cartilage area in the EXO group relative to the model group (Fig. 3C,D; Brown–Forsythe ANOVA: F = 19.75, df = 2.15, partial $\eta$² = 0.72). In contrast, exosomes derived from Igf1-deficient ADSCs (shIGF1-EXO group) failed to reproduce these histological benefits. Both histological scores and Safranin O–positive cartilage areas were significantly reduced compared with the EXO group, indicating impaired fibrocartilage regeneration and suboptimal structural integration. Collectively, these findings demonstrate that IGF1 is a critical mediator of exosome-driven histological repair and chondrogenesis at the tendon–bone interface during rotator cuff healing.

To investigate the contribution of ADSC-derived exosomes to angiogenesis during tendon–bone healing, immunohistochemical staining and gene expression analyses were performed on tendon tissues harvested 8 weeks after surgery. A significant increase in VEGF-positive staining was demonstrated in the EXO group (Fig. 4A,B, Brown–Forsythe ANOVA, F = 14.92, df = 2.15, partial $\eta$² = 0.66). Consistent with the IHC findings, RT-qPCR analysis demonstrated that exosome treatment significantly upregulated the mRNA expression levels of Vegfa (Fig. 4C, Brown–Forsythe ANOVA, F = 12.38, df = 2.6, partial $\eta$² = 0.80), Cd31 (Fig. 4D, Brown–Forsythe ANOVA, F = 11.75, df = 2.6, partial $\eta$² = 0.79), and $\alpha$-SMA (Fig. 4E, Brown–Forsythe ANOVA, F = 10.52, df = 2.6, partial $\eta$² = 0.78) in tendon tissues, further supporting enhanced angiogenic activity.

In contrast, the pro-angiogenic responses were substantially attenuated when exosomes were derived from Igf1-deficient ADSCs. Compared with the EXO group, the shIGF1-EXO group exhibited significantly reduced VEGF staining intensity as well as lower transcript levels of Vegfa, Cd31, and Acta2. These findings indicate that IGF1 plays a critical role in mediating the angiogenic activity of ADSC-derived exosomes in the context of rotator cuff repair.

To evaluate the effects of ADSC-derived exosomes on tendon cell pyroptosis following rotator cuff injury, tendon tissues were harvested 1 week after surgery for propidium iodide (PI) staining and Western blot analysis. A sham-operated group was included as a baseline control. Compared with the sham group, the untreated model group exhibited a marked increase in PI-positive cells, indicating enhanced pyroptotic activity in the injured tendon tissue (Fig. 5A,B). Treatment with exosomes derived from control ADSCs (EXO group) significantly reduced the number of PI-positive cells relative to the model group, suggesting an inhibitory effect of exosomes on tendon cell pyroptosis during the early healing phase (Fig. 5B; Brown–Forsythe ANOVA: F = 17.25, df = 3.20, partial $\eta$² = 0.73).

Consistent with these findings, Western blot analysis demonstrated a pronounced upregulation of gasdermin D N-terminal fragment (GSDMD-N), a key executor of pyroptosis, in the model group, whereas exosome treatment markedly suppressed GSDMD-N expression toward baseline levels (Fig. 5C). Quantitative analysis of GSDMD-N normalized to $\beta$-actin further confirmed this reduction (Fig. 5D; Brown–Forsythe ANOVA: F = 13.89, df = 3.8, partial $\eta$² = 0.84).

Notably, the anti-pyroptotic effects of exosome treatment were significantly attenuated when exosomes were derived from Igf1-knockdown ADSCs. Compared with the EXO group, the shIGF1-EXO group exhibited a higher proportion of PI-positive cells and significantly elevated GSDMD-N protein levels. These results indicate that IGF1 is required for the exosome-mediated suppression of tendon cell pyroptosis during the early stage of rotator cuff healing.

To further examine the regulatory effects of ADSC-derived exosomes on the inflammatory response associated with NLRP3 inflammasome activation and pyroptosis, the protein and mRNA expression levels of the pro-inflammatory cytokines IL-1$\beta$ and IL-18 were assessed in tendon tissues one week after rotator cuff injury (Fig. 6A–D). Compared with the sham-operated group, the untreated model group exhibited significantly elevated protein levels of IL-1$\beta$ and IL-18, reflecting a robust inflammatory response during the early healing phase (Fig. 6A,B; Brown–Forsythe ANOVA: F = 14.23, df = 3.20, partial $\eta$² = 0.71 for IL-1$\beta$; F = 12.85, df = 3.20, partial $\eta$² = 0.68 for IL-18). Treatment with exosomes derived from control ADSCs (EXO group) significantly reduced the protein levels of both IL-1$\beta$ and IL-18 compared with the model group, indicating a pronounced anti-inflammatory effect of exosome therapy. Consistent with the protein expression data, RT-qPCR analysis demonstrated similar trends at the transcriptional level. The mRNA expression of Il1b and Il18 was markedly upregulated in the model group but significantly suppressed following exosome treatment (Fig. 6C,D; Brown–Forsythe ANOVA: F = 10.87, df = 3.8, partial $\eta$² = 0.79 for Il1b; F = 9.52, df = 3.8, partial $\eta$² = 0.77 for Il18).

Notably, these anti-inflammatory effects were substantially attenuated when exosomes were derived from Igf1-inhibited ADSCs. Compared with the EXO group, the shIGF1-EXO group exhibited significantly higher levels of IL-1$\beta$ and IL-18 at both the protein and mRNA levels. These findings indicate that IGF1 plays a key role in mediating the immunomodulatory properties of ADSC-derived exosomes during the early inflammatory phase of rotator cuff tendon healing.

To investigate the influence of ADSC-derived exosomes on NLRP3 inflammasome activation in the early phase of tendon healing, Western blot analysis was performed on tendon tissues collected one-week post-surgery. The NLRP3 inflammasome is a critical innate immune signaling complex that detects pathogenic stimuli or cellular stress. Once activated, NLRP3 recruits the adaptor protein ASC through PYD–PYD interactions. ASC then facilitates the conversion of pro-caspase-1 into its active form, triggering the release of pro-inflammatory cytokines and initiating pyroptosis [18, 19, 20]. Compared to the sham group, the model group showed marked upregulation of NLRP3 (Fig. 7A,B, Brown–Forsythe ANOVA, F = 13.25, df = 3.8, partial $\eta$² = 0.83), ASC (Fig. 7A,C, Brown–Forsythe ANOVA, F = 12.18, df = 3.8, partial $\eta$² = 0.81), pro-caspase-1 (Fig. 7A,D, Brown–Forsythe ANOVA, F = 10.92, df = 3.8, partial $\eta$² = 0.79), and cleaved caspase-1 protein levels (Fig. 7A,E, Brown–Forsythe ANOVA, F = 14.57, df = 3.8, partial $\eta$² = 0.85), indicating robust inflammasome activation (p 0.01).

Treatment with exosomes derived from control ADSCs (EXO group) significantly suppressed the expression of these inflammasome-related proteins compared with the model group, indicating effective inhibition of NLRP3 inflammasome activation. In contrast, this suppressive effect was markedly attenuated when exosomes were derived from Igf1-knockdown ADSCs. Compared with the EXO group, the shIGF1-EXO group exhibited significantly higher levels of NLRP3, ASC, and cleaved caspase-1, demonstrating incomplete inhibition of inflammasome signaling.

Collectively, these findings indicate that IGF1 is essential for the ability of ADSC-derived exosomes to suppress NLRP3 inflammasome activation during the early inflammatory phase of rotator cuff tendon healing, thereby linking exosomal IGF1 to coordinated regulation of inflammation, pyroptosis, and tissue repair.