Bifunctional adECM bioscaffold with STIM1-ASCs and IGF-2 promotes functional masseter VML repair via myogenesis and fibrosis suppression

Wei Liang; Rigele Ao; Mengli Xu; Mengying Jin; Meng Han; Zimo Wang; Wanwen Dang; Hongxu Wu; Weibo Lin; Yonghuan Zhen; Tao Xu; Yang An

doi:10.1016/j.bioactmat.2025.08.019

. 2025 Aug 29;54:466–491. doi: 10.1016/j.bioactmat.2025.08.019

Bifunctional adECM bioscaffold with STIM1-ASCs and IGF-2 promotes functional masseter VML repair via myogenesis and fibrosis suppression

Wei Liang ^a,¹, Rigele Ao ^a,¹, Mengli Xu ^a,¹, Mengying Jin ^a,¹, Meng Han ^a, Zimo Wang ^a, Wanwen Dang ^a, Hongxu Wu ^a, Weibo Lin ^a, Yonghuan Zhen ^a, Tao Xu ^b, Yang An ^a,^c,^⁎

PMCID: PMC12410469 PMID: 40918736

Abstract

Craniofacial muscles are essential for a variety of functions, including fine facial expressions. Severe injuries to these muscles often lead to more devastating consequences than limb muscle injuries, resulting in the loss of critical functions such as mastication and eyelid closure, as well as facial aesthetic impairment. Therefore, the development of targeted repair strategies for craniofacial muscle injuries is crucial. In this study, we engineered an adipose-derived decellularized extracellular matrix (adECM) bioscaffold co-loaded with seed cells and bioactive factors. The seed cells were STIM1-overexpressing adipose-derived stem cells (STIM1-ASCs), which exhibit directed and highly efficient myogenic differentiation, addressing the low differentiation efficiency of conventional ASCs that limits muscle regeneration. The bioactive factor used was insulin-like growth factor-2 (IGF-2), which modulates the immune microenvironment by reprogramming mitochondrial energy metabolism to promote M2 macrophage polarization. These M2 macrophages further suppress fibroblast collagen deposition, alleviating muscle fibrosis, while simultaneously enhancing the myogenic differentiation of STIM1-ASCs and myotube formation. Together, the recellularized adECM bioscaffold harnesses these dual mechanisms (promoting functional muscle regeneration and anti-fibrotic repair) to significantly improve the recovery of volumetric muscle loss (VML) in the masseter. The development of this bifunctional bioscaffold offers a novel therapeutic strategy and theoretical foundation for treating severe craniofacial muscle injuries.

Keywords: Volumetric muscle loss, Adipose decellularized extracellular matrix, Myogenic differentiation, Muscle regeneration, Fibrosis

Graphical abstract

We developed a large-scale decellularized adipose tissue scaffold (adECM) recellularized with STIM1-ASCs and loaded with IGF-2 cytokines. The STIM1-ASCs were engineered using lentiviral transfection technology and exhibit high myogenic differentiation potential, addressing the absolute deficiency of muscle satellite cells caused by VML. IGF-2, acting early, induces macrophage metabolic reprogramming, altering their energy metabolism pathways, then promotes the activation of M1 macrophages into M2 macrophages. The activation of M2 macrophages significantly reduces muscle fibrosis and facilitates the further fusion of myofibers into mature myofibers.

Highlights

•
Insufficient muscle regeneration and severe fibrotic scarring are pathological characteristics of craniofacial volumetric muscle loss (VML).
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The absolute deficiency of muscle satellite cells (SCs) and the imbalance in M1-to-M2 macrophage polarization are two fundamental causes of severe functional impairment in VML.
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A novel muscle tissue engineering construct was developed using adipose decellularized extracellular matrix (adECM), which integrates seed cells (STIM1-ASCs) and the cytokine IGF-2. Among these components, STIM1-ASCs address the issue of the absolute deficiency of SCs, while IGF-2 mitigates the occurrence of fibrotic scarring.
•
IGF-2 promotes macrophage polarization toward the M2 phenotype by inducing metabolic reprogramming and through activating PI3K-Akt signal pathway in macrophages, thereby facilitating muscle fiber maturation and reducing the extent of muscle fibrosis.

1. Introduction

Craniofacial volumetric muscle loss (VML), resulting from trauma, cleft lip/palate, and other causes, presents unique clinical challenges, leading to both functional impairments (e.g., mastication, deglutition, ocular closure) and significant aesthetic deformities [1]. These combined deficits substantially increase the risk of psychological comorbidities in affected patients [2]. The clinical burden is also considerable: data from 2020 indicate approximately 200,000 craniofacial reconstruction procedures annually in the U.S. alone, making it the fourth most common surgical intervention [3].

Current clinical management relies on composite tissue flaps. However, existing tissue flap transplantation techniques often lead to postoperative complications at the donor site, such as infection, hematoma, and scarring. Simultaneously, the structural and functional recovery of the damaged muscle at the recipient site after flap transplantation is suboptimal. The failure of revascularization and nerve reconstruction at both the donor and recipient sites, as well as the progressive fibrotic scar formation at the injury site, hinder the effective integration of muscle tissue at both sites, which are the main reasons for the poor therapeutic outcomes [4,5]. More importantly, tissue flap transplantation techniques fail to fully restore native sensation, dynamic function, and natural facial expressivity. This unmet clinical need underscores the urgency for developing novel therapies that can simultaneously reconstruct craniofacial architecture, restore physiological muscle function, and mitigate the psychological sequelae of VML.

The goal of muscle tissue engineering strategies is to achieve the regeneration of functional muscle tissue, and the main factors influencing functional muscle regeneration include the reinnervation of muscle by nerves and the adequate regeneration of blood vessels at the site of injury. As a result, a large number of studies have focused on using various tissue engineering materials to regenerate nerves and blood vessels at the VML site, with significant progress already made, as detailed in the reviews and studies by Grayson et al. and others [6,7]. In recent years, researchers have increasingly recognized that inflammation and fibrosis after VML injury are also key factors limiting functional muscle regeneration [8], as chronic inflammation and muscle fibrosis hinder muscle tissue regeneration, with their effects amplifying one another and further exacerbating muscle dysfunction.

Understanding the pathophysiology of muscle injury and repair is crucial for addressing the challenges associated with these conditions. On the one hand, the regenerative process primarily relies on muscle satellite cells (SCs), the sole myogenic precursors in skeletal muscle tissue. Following injury, SCs are activated, proliferate, and ultimately fuse to form mature myofibers [9]. However, VML is characterized by severe depletion of SCs, resulting in an insufficient pool of muscle progenitor cells to support regeneration [10,11]. Craniofacial muscles exhibit significant differences from limb and trunk muscles in terms of their embryonic origin, satellite cell phenotypes, extracellular matrix composition, genetic and molecular regulation during development, and the capacity and extent of regeneration following injury. The satellite cells of craniofacial muscles originate from the neural crest (pharyngeal arch mesoderm), whereas those of limb and trunk muscles derive from the somite mesoderm [12,13]. Craniofacial muscles have fewer SCs, leading to weaker repair abilities, more severe muscle fibrosis after injury, and poorer muscle regeneration compared to limb and trunk muscles [14,15]. Although various trunk and limb VML animal models have been developed, and the functional recovery of these VML models using tissue-engineered muscle grafts has been explored, there is still a need for tissue-engineered grafts better suited for craniofacial muscle injuries due to the inherent differences between craniofacial and limb/trunk muscles [3].

On the other hand, the immune microenvironment, particularly macrophages, plays an equally critical role in determining regenerative outcomes [16]. During the acute injury phase (days 1–2), pro-inflammatory M1 macrophages dominate the environment. By day 3, a phenotypic shift toward anti-inflammatory M2 macrophages occurs, which are essential for promoting the myogenic differentiation of muscle progenitors [17]. Disruption of this transition, resulting in persistent M1 dominance, leads to excessive extracellular matrix deposition and fibrotic scarring [18]. In summary, these pathophysiological insights should guide the development of next-generation tissue engineering strategies. An optimal approach would therefore involve: 1) replenishing the depleted muscle progenitor cell population, and 2) precisely modulating macrophage polarization dynamics. Such dual-targeting interventions could synergistically enhance functional muscle regeneration while minimizing fibrotic complications.

While satellite cells (SCs) represent the sole endogenous myogenic progenitors in skeletal muscle, their limited abundance (comprising only 1–5 % of total muscle cells), restricted in vitro expansion capacity, and phenotypic instability pose significant challenges for tissue engineering applications [19]. As an alternative, mesenchymal stem cells (MSCs) derived from various sources, including bone marrow, adipose tissue (adipose-derived stem cells, ASCs), umbilical cord mesenchymal stem cells, and embryonic lineages, have demonstrated myogenic differentiation potential under appropriate induction conditions [[20], [21], [22]]. Among these MSC types, ASCs have emerged as particularly promising candidates due to their accessibility, robust proliferative capacity, and favorable immunogenic profile [23]. However, their clinical translation is hindered by suboptimal myogenic conversion rates (∼15 % efficiency), which are considerably lower than those of native SCs [24]. Our previous work, which utilized ASCs for tibialis anterior VML repair in rat models, confirmed their myogenic potential through histological evidence but highlighted limitations in both fiber quantity and maturation [25]. To address these limitations, we employed single-cell RNA sequencing to uncover the molecular determinants of ASC myogenesis. We identified stromal interaction molecule (STIM), a key regulator of calcium homeostasis with two isoforms (STIM1/STIM2) [26,27], as a critical mediator of directed myogenic differentiation. While the role of STIM in C2C12 myoblast fusion is well established [28], its function in ASC myogenesis remains unexplored. Therefore, we hypothesize that overexpression of STIM1 in ASCs (resulting in STIM1-ASCs) may overcome current limitations in differentiation efficiency, offering enhanced therapeutic potential for muscle regeneration.

The efficacy of tissue-engineered constructs in reducing muscle fibrosis fundamentally relies on precise spatiotemporal modulation of macrophage phenotypes within the injury microenvironment [29]. Among the three essential components of tissue engineering scaffolds, cytokine-based strategies emerge as the most promising approach for achieving this targeted immunomodulation [16]. Recent research has elucidated the pleiotropic role of insulin-like growth factor-2 (IGF-2) in orchestrating muscle repair through dynamic, phase-specific mechanisms [30]. During the early stages of injury, IGF-2 primarily stimulates the proliferation of muscle progenitor cells to replenish the depleted satellite cell pool. Subsequently, it induces a functional shift toward immunomodulation, promoting the polarization of pro-regenerative M2 macrophages. These alternatively activated macrophages perform dual therapeutic functions: 1) suppressing fibrotic tissue formation through paracrine signaling, and 2) enhancing myoblast fusion to facilitate the formation of mature myofibers [31]. This temporal versatility distinguishes IGF-2 from conventional M2-polarizing cytokines (e.g., IL-4, IL-13) [32]. While these interleukins demonstrate immunomodulatory potential, they lack the complementary myogenic properties of IGF-2, specifically the ability to activate muscle progenitor cells and promote terminal myogenic differentiation [33].

Futhermore, natural tissue-derived scaffolds demonstrate superior clinical potential compared to synthetic alternatives due to their reduced immunogenicity and enhanced biocompatibility [6,34]. Among these, decellularized extracellular matrix (dECM) materials have emerged as particularly promising, as they preserve native 3D microarchitectures and retain critical bioactive factors that support cellular proliferation and differentiation [35]. Several dECM products, including small intestinal submucosa (SIS), urinary bladder matrix (UBM), and skeletal muscle/dermal dECMs, have successfully achieved clinical translation, demonstrating efficacy in VML treatment across both preclinical and clinical studies [[36], [37], [38]]. However, these materials face notable limitations, including donor tissue scarcity, technically demanding preparation protocols, and substantial production costs [39]. In contrast, adipose-derived dECM (adECM) offers distinct advantages: 1) abundant tissue availability, 2) simplified processing requirements, and 3) cost-effectiveness, while maintaining the beneficial properties of other dECM materials [40,41]. Our recent investigations have specifically shown that volumetric adECM scaffolds significantly enhance functional muscle regeneration in VML models, while mitigating muscle dysfunction [25,42,43]. Notably, compared to alternative adECM formulations (e.g., hydrogels or 3D-printed bioinks) [44], volumetric adECM scaffolds better preserve: (i) native ECM composition, (ii) structural integrity, and (iii) microenvironmental cues critical for myofiber regeneration—avoiding the compositional and architectural compromises inherent to hydrogel processing [[45], [46], [47]]. This advantage is particularly relevant for large-volume defects, as tissue-derived dECM scaffolds have demonstrated superior performance over 3D-printed alternatives in volumetric tissue restoration [48].

In addition, given that craniofacial muscles have fewer satellite cells and more severe muscle fibrosis after injury, developing an animal model that accurately reflects the pathophysiological characteristics of craniofacial VML is important, but it also presents challenges. Researchers, such as Lucas and Brittany [3,49], have found that histological results from bite muscle VML or masseter muscle VML models exhibit significant variations even within the same experimental group. Even when maintaining a consistent VML volume, slight variations in the shape and location of the muscle excision can reduce group homogeneity. Despite these issues, precise anatomical localization and critical muscle defect excision can significantly reduce individual variability within experimental groups. Zhao et al. investigated the volume of critical muscle defects in the mouse masseter VML model and identified a 2 mm diameter muscle defect as the critical threshold, which provides important reference information for the development of craniofacial VML models [50]. Meanwhile, Sydney et al. and Lucas et al. published experimental papers in the same year, providing a detailed description of the preparation process for the SD rat masseter VML model, offering accurate anatomical localization and clear surgical areas for researchers [3,51].

In summary, this study developed a novel therapeutic strategy for masseter VML using a recellularized adECM bioscaffold incorporating STIM1-overexpressing adipose-derived stem cells (STIM1-ASCs) and insulin-like growth factor-2 (IGF-2). The approach addresses two critical aspects of muscle regeneration by: 1) genetically enhancing the myogenic potential of the cells, and 2) precisely modulating the regenerative microenvironment through immunomodulation. As illustrated in the Graphical Abstract, this combinatorial strategy synergistically tackles the dual challenges of insufficient myogenesis and excessive fibrosis in VML repair. The study establishes both a theoretical framework and practical methodology for developing next-generation therapies for craniofacial muscle reconstruction, with particular relevance to functional and aesthetic restoration.

2. Materials and methods

2.1. Preparation of the adECM with vascular pedicle

First, following the protocol from previous studies, we harvested vascularized adipose tissues from the inguinal region of SD rats [42,52]. During the procedure, we carefully dissected and separated the blood-supplying vessels. A blunt hollow needle, matching the vessel diameter, was inserted into the supplying vessel toward the distal end and fixed at the vascular pedicle using sutures. Through this hollow needle, heparin solution was injected into the adipose tissues to prevent thrombosis formation. Next, the distal end of the hollow needle was connected to a bioreactor, where a continuous perfusion of 0.5 % SDS (Sodium Dodecyl Sulfate) solution (S1010, Beijing Solarbio Science & Technology Co., Ltd. China) was applied for 48 h, followed by deionized water for 24 h to remove residual SDS, then 1 % Triton-X 100 (Cat No.1271889, Leyan, Shanghai, China) for 24 h, and another 24 h of deionized water to clear remaining Triton-X 100. Finally, a 72-h perfusion of isopropanol solution was conducted to completely remove lipid components from the tissue, followed by deionized water for 24 h to eliminate any residual isopropanol. The temperature used in this experiment was maintained at 37 °C, with a flow rate of 2–3 mL/min. Additionally, we ensured that the perfusion pressure in the bioreactor was set at 80 mm Hg, which is crucial for preventing the collapse of native vasculature within the adECM scaffold. When the tissue achieved a transparent appearance, the preparation of adECM was deemed complete, and the sample was stored in PBS containing 1 % penicillin-streptomycin.

2.2. Characterization of adECM

2.2.1. cDNA content

The residual nucleic acid concentration in adECM was measured using a commercial tissue DNA extraction kit (E.Z.N.A.® Tissue DNA Kit, Omega, USA). Lyophilized adECM (with an equivalent amount of lyophilized native adipose tissue as the control) was digested in OB proteinase solution at 55 °C for 2 h, followed by centrifugation at 10,000g for 5 min at 4C. DNA concentration in the supernatant was then measured using a Nanodrop (BioRad Laboratories, Hercules, CA) device. Each sample group had n = 3.

2.2.2. Glycosaminoglycan (GAG) content

Native adipose tissue and adECM samples (n = 3) were weighed and digested overnight at 65 °C using a papain extraction reagent. Sulfated glycosaminoglycans (GAGs) were quantified with a Sulfated Glycosaminoglycan Assay Kit (GENMED, China) following the manufacturer's instructions. Absorbance was measured at 656 nm using a microplate reader (SpectraMax M5, Molecular Devices, Sunnyvale, CA, USA).

2.2.3. Swelling rate and degradation rate

Native adipose tissue and adECM samples (100 mg each) were weighed separately and incubated in PBS at 37 °C. The weight of each sample was recorded every 30 min, and a swelling rate curve was plotted after 6 h. This experiment was repeated three times. The swelling rate was calculated using the formula: Swelling rate = (Ww - Wd)/Wd, where Ww is the swollen weight, and Wd is the dry weight.

Additionally, samples of native adipose tissue and adECM with the same initial weight were incubated in a 0.05 mg/mL collagenase type I solution (BioFroxx, Germany) at the same temperature. The residual dry weight of each sample was measured every 3 days, and degradation rate curves for native adipose tissue and adECM were plotted after 15 days. This experiment was also repeated three times. The degradation ratio (%) was calculated as: Degradation ratio = (m0 - mt)/m0 × 100, where m0 is the initial weight, and mt is the weight on the specified day of degradation.

2.2.4. Histological staining

Native adipose tissue and adECM were fixed in 4 % paraformaldehyde (EE0001, Shandong Sparkjade Biotechnology Co., Ltd., China), embedded in paraffin or OCT, and sliced into 5 μm sections. The slides were stained with hematoxylin and eosin (H&E), Masson trichrome, oil Red O, Alcian blue and DAPI staining, respectively.

2.2.5. Scanning electron microscopy (SEM)

Scanning electron microscopy (FE-SEM, S-4800, Hitachi) was used to observe the microstructure of native adipose tissue and adECM. Samples were fixed in a 2.5 % glutaraldehyde solution at room temperature, followed by dehydration in a graded ethanol series, immersion in isoamyl acetate, vacuum drying, gold sputter coating, and SEM imaging.

2.2.6. Mechanical properties assessment

The native adipose tissue and adECM were trimmed into cylinders with a diameter of 5 mm and a thickness of 5 mm. Their biomechanical properties were tested using a dynamic mechanical analyzer (E3000, UK). Samples were compressed under a 0.1 N load at a speed of 15 mm/min. A stress-strain curve was generated, and the compression modulus of each sample was calculated based on this curve’ slope. Similarly, the tensile properties of the adECM and native adipose tissue were evaluated. An initial tensile force of 0.5 N was applied at a speed of 10 mm/min. A stress-strain curve was plotted, and the tensile modulus of the samples was calculated.

2.2.7. SDS-polyacrylamide gel electrophoresis (SDS-PAGE)

SDS-PAGE was performed to analyze the protein composition of adECM. The adECM and collagen I hydrogel solution (serving as the control group) were electrophoresed on 10 % polyacrylamide gels under reducing conditions (with 5 % 2-mercaptoethanol). Rainbow broad spectrum protein marker (11–245 kD, PR1920-20T, Solarbio, China) was used as a protein ladder. Protein bands were visualized with Gel-Code Blue, and images were captured using a camera (Canon, Japan).

2.3. IGF-2 loaded adECM regulates macrophage polarization

2.3.1. The efficiency of adECM loading IGF-2

According to the method described in previous literature [53], PBS solutions of rat IGF-2 (MCE, HY-P700185AF) at concentrations of 0 ng/mL, 5 ng/mL, 10 ng/mL, and 20 ng/mL were first prepared. Then, 1 mL of each PBS solution and 10 mg of lyophilized adECM scaffold were transferred together into a 1.5 mL EP tube. The EP tube was then placed upright in a cell incubator (37 °C, humidified 5 % CO₂ atmosphere) for co-incubation. After 24 h, we obtained adECM scaffolds loaded with different concentrations of IGF-2. Additionally, 500 μL of the IGF-2 solution was collected from each EP tube. The remaining IGF-2 content in the solution was measured using ELISA to estimate the amount of IGF-2 loaded onto the adECM scaffold, which represents the drug loading capacity of IGF-2. This experiment was repeated three times.

2.3.2. IGF-2 release analysis of adECM

The IGF-2-loaded adECM scaffolds (5 ng/mL, 10 ng/mL, and 20 ng/mL) were immersed in 1 mL of PBS and incubated at 37 °C. At different time points (30 min, 1, 2, 3, 4, 5, 6, and 7 days), 500 μL of solution was collected from each group, and an equal volume of PBS was added to maintain the solution volume at 1 mL. The IGF-2 solutions collected at different time points were temporarily stored at −80 °C. Finally, ELISA (MM-0196R2; MEIMIAN, JIANGSU, CHINA) was used to measure the amount of IGF-2 released from the adECM scaffold at each time point, and a time-release curve was plotted.

2.3.3. Bone marrow-derived macrophages (BMDMs) cell isolation and culture

Male SD rats (4–6 weeks old) were sacrificed, and their femurs were dissected [33]. The bone marrow was flushed out using α-minimum essential medium (α-MEM; Hyclone Laboratories, Logan, UT, USA). The cell suspension was then passed through 40 μm microporous filter membranes. After centrifugation at 1000 rpm for 3 min, the supernatant was discarded, and 10 mL of red blood cell lysis bufer was added, with the mixture shaken every 5 min for a total of 10 min. After centrifuging again at 1000 rpm for 3 min, the cell pellet was resuspended in medium containing 10 % inactivated fetal bovine serum (FBS01, Sino Biological Inc, China) and 1 % Mycoplasma Elimination Reagent Kit (FM421, TransGen Biotech, China), then, cells were seeded in a T25 cm² culture flask. After 3–4 days of incubation, suspended cells were collected, centrifuged, and then inoculated into a T25 cm² flask with 20 ng/mL GM-CSF (granulocyte-macrophage colony-stimulating factor) to stimulate macrophage differentiation. The culture medium was refreshed every 2 days, and BMDMs were obtained after 7 days. Then, BMDMs were identifed by immunofuorescence staining to detect cells expressing CD11b and F4/80.

2.3.4. IGF-2-loaded adECM scaffolds regulates macrophage polarization in vitro

The IGF-2-loaded adECM scaffolds were co-cultured with macrophage using Transwell inserts (KTA5010, Abbkine, Wuhan, China). Macrophages were seeded at a density of 10 × 10⁵ cells/well in the lower chamber of 6-well plates. Various IGF-2-loaded adECM scaffolds (0 ng/mL, 5 ng/mL, 10 ng/mL, and 20 ng/mL) in 2 mL of α-MEM medium were added to the upper chamber of the Transwells. For the control group, 2 mL of α-MEM medium was added to the upper chamber, while the positive group received 2 mL of α-MEM medium supplemented with 100 ng/mL of IL-4. After 72 h of co-culture, the collected cells were incubated with CD80-PE antibody (1:200, 12-0800-82, eBioscience), CD206-APC (MRC1/APC) antibody (1:100, bs-4727R, Bioss), and F4/80-FITC antibody (1:200, AER-051-F, ThermoFisher) for 30 min. The cells were then washed twice with staining buffer and resuspended in 100 μL of staining buffer. Finally, the stained cells were analyzed using a flow cytometer (BD Accuri C6, USA).

2.3.5. Quantitative real-time polymerase chain reaction (RT-qPCR)

Total RNA of macrophages or fibroblasts was extracted using TRIzol RNA isolation reagent (AC0205-B, Shandong Sparkjade Biotechnology Co., Ltd). cDNA was reverse-transcribed using a ReverTra Ace qPCR kit (Toyobo, Osaka, Japan). Real-time quantitative polymerase chain reaction (RT-qPCR) was performed using Evo M-MLV One Step RT-qPCR Kit (SYBR, AG11732, ACCURATE BIOTECHNOLOGY (HUNAN)CO., LTD, ChangSha, China) for the following genes:

GAPDH, forward 5′-GTCCATGCCATCACTGCCACTC-3′ and reverse 5′-GATGACCTTGCCCACAGCCTTG-3’;

IL-1β, forward 5′-CAACTGTCCCTGAACTCAACTGT-3′ and reverse 5′-GAGATGCTGCTGTGAGATTTGAA-3’;

Arg-1, forward 5′-CAGATCCAACCCGATTATAAGGGA-3′ and reverse 5′-GTCTCTCACATTGTACTCTGTTTCT-3’;

IL-10, forward 5′-CAGAAATCAAGGAGCATTTG-3′ and reverse 5′-CTGCTCCACTGCCTTGCTTT-3’;

TGF-α, forward 5′-CCAACAAGGAGGAGAAGTTCC-3′ and reverse 5′-TCTGCTTGGTGGTTTGCTAC-3’;

β-actin, forward 5′-GCCGCCAGCTCACCA-3′ and reverse 5′-CCCACGATGGAGGGGAAGA-3’;

COL1A1, forward 5′-CGAGGCTCTGAAGGTCCCC-3′ and reverse 5′-CCAGGAGCACCATTGGCA-3’;

COL3A1, forward 5′-TCGAGGCAGTGATGGTCAAC-3′ and reverse 5′-GGTCCAACTTCACCCTTAGCA-3’; Relative gene expression was normalized to the housekeeping gene GAPDH by the 2^−ΔΔCt method.

2.3.6. Seahorse analysis

The extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), indicative of glycolysis and respiration, respectively, were monitored using an XF96 Extracellular Flux Analyzer (Agilent Technologies, Santa Clara, CA, USA) in conjunction with the Seahorse XF Glycolysis Stress Test Kit and Seahorse XF Cell Mito Stress Test Kit. Briefly, macrophages were cultured in transwell six-well plates using high-glucose DMEM medium containing 10 % fetal bovine serum with or without 100 ng/mL IL-4. After that, the macrophages were co-incubated with adECM scaffolds loaded with different IGF-2 (0 ng/mL, 5 ng/mL, 10 ng/mL, and 20 ng/mL) for 72 h. Then, the cells were seeded into a 96-well FluxPak at a density of 1 × 10⁴ cells per well and cultured for 12 h with the original medium. Mitochondrial respiratory parameters and glycolytic activity were assessed via OCR (pmol/min/10,000 cells) and ECAR (mpH/min/10,000 cells), respectively, using injection solutions that included Oligomycin, FCCP, rotenone/antimycin A,glucose, 2-deoxy-D-glucose. To investigate the mechanism by which IGF-2 regulates macrophage energy metabolism, we also established the adECM + IGF-2+LY294002 group. In this group, the concentration of IGF-2 is 10 ng/mL, and the concentration of LY294002 is 25 μM [54].

2.3.7. Western blotting

Macrophages in the adECM + IGF-2 (10 ng/mL) group, adECM + IGF-2+LY294002 group, and IL-4 group were lysed using RIPA lysis buffer supplemented with phosphatase and protease inhibitor cocktail and detected the concentration by BCA Protein Assay Kit (C503061, Beijing Boxbio Science & Technology Co.,Ltd.). After mixing with SDS loading buffer and heating at 95 °C for 5 min, 15 μL samples were loaded onto an One-Step PAGE Gel (E303-01, Vazyme, Biotech Co., Ltd, China) and subsequently electrotransferred onto PVDF membranes (LS-PVDF-045-Z, LANSO, China). The membranes were blocked with TBST containing 5 % nonfat dry milk at room temperature for 1.5 h and incubated with primary antibodies at the indicated dilutions overnight at 4 °C. The primary antibodies used were as follows: PI3K (#4257T, CST, 1:1000), p-PI3K (#4249T, CST, 1:1000), AKT (#9272, CST, 1:1000), p-AKT (#4060T, CST, 1:1000), and GAPDH (#181602, Abcam, UK, 1:1000). Following incubation with a goat anti-rabbit secondary antibody for 1.5 h at room temperature, the PVDF membranes were visualized using an ECL chemiluminescence kit (E1050, LabLead, China) and imaged with the TANON 5200 Multi imaging system (TANON, Beijing, China).

2.3.8. RNA sequencing (RNA-seq)

RNA samples from macrophage suspension in the Control group and adECM + IGF-2 (10 ng/mL) group, were prepared for general transcriptome RNA sequencing data. The RNA samples were extracted by the Trizol method. RNA-seq was performed by Oe Biotech (Shanghai, China). The edge package in R package DESeq2 was used to screen out the differently expressed genes with the threshold: p-value <0.05 and Log² Fold Change (FC) ≥ 1.

2.4. IGF-2-loaded adECM scaffolds alter the function of human normal skin fibroblasts by promoting M2 macrophage polarization

After co-culturing the IGF-2-loaded adECM scaffold with macrophages, the supernatant was mixed with the fibroblast culture medium at a 1:1 vol ratio to prepare the fibroblast-conditioned medium. Fibroblasts were seeded at a density of 1 × 10⁵ cells/well on the bottom of the Transwell (pre-coated by Plurigel Matrix, GL101, Vazyme Biotech Co.,Ltd) chamber in a 24-well plate. Then, 1 mL of fibroblast-conditioned medium was added, and the plate was placed in a cell incubator for 24 h. Afterward, the Transwell (KTA5010, Abbkine, Wuhan, China) chamber was removed, washed with PBS (6062712, Shenzhen Dakewe Bioengineering Co., Ltd.), and the bottom of the chamber was immersed in crystal violet staining solution for 10 min. The number of fibroblasts migrating through the Transwell membrane in each group was observed under a microscope, and images were taken. The results were analyzed and quantified using Image J software.

For the scratch assay, human normal skin fibroblasts were first seeded in a 96-well plate and cultured in different fibroblast-conditioned medium. Once the cells were fully adherent, the 96-well plate was transferred to the Incucyte system for the scratch assay. Images were taken every 3 h to record the closure of the scratch, and the scratch closure rate for each group was analyzed using Image J software.

2.5. Myogenic differentiation of STIM1-ASCs inside IGF-2 loaded adECM

2.5.1. Preparation of STIM1-ASCs and myogenic differentiation

First, according to our previous method, adipose-derived stem cells (ASCs) were isolated from the inguinal fat tissue of rats and their cell markers (CD11b, CD29, CD34, CD45, CD44, CD73, CD90) were detected by flow cytometry [25]. Next, green fluorescent protein (GFP) and STIM1 lentiviral vectors were constructed by a commercial company. ASCs at passages 2–3 were cultured in 6-well plates and transduced with lentiviruses at a multiplicity of infection (MOI) of 10. 3 days post-transduction, GFP-ASCs were observed using a confocal laser scanning microscope, and GFP-ASCs and STIM1-ASCs were selected using puromycin. Finally, 1 × 10⁶ STIM1-ASCs were separately seeded into confocal dishes and subjected to myogenic differentiation using commercially available myogenic induction medium (CTCC-Y006, CHINA), with half-medium changes every other day. After 10 days, both GFP-ASCs and STIM1-ASCs underwent myosin heavy chain (MHC) immunofluorescence staining, and the expression levels of myogenesis-related proteins (MYH2, MHC, and Desmin) were analyzed using Western blot.

2.5.2. SCs isolation and culture

According to the previous study [33], male SD (Sprague-Dawley) rats, aged 5 days, were sacrificed, and all leg muscles were dissected into small pieces. The tissue was then treated with 0.2 % type I collagenase for 30 min, followed by centrifugation at 500×g for 3 min. The supernatant was discarded, and the pellet was incubated with trypsin containing 0.2 % EDTA at 37 °C for 30 min. The digestion was terminated by the addition of complete medium. The remaining impurities were removed by filtration through 40 μm filters. The cells were subsequently centrifuged at 200×g for 3 min and resuspended in 5 mL of medium. A stepwise density gradient was created by carefully adding 2 mL of 80 % Percoll (P8370, Solarbio, China), 8 mL of 20 % Percoll, and 3 mL of cell suspension into the centrifuge tube. The sample was then centrifuged at 5000×g for 5 min. Cells from the interphase between the 20 % and 80 % Percoll layers were collected for inoculation. The suspended cells were subjected to differential adhesion for 1 h and 2 h, after which the remaining suspended cells were removed and re-inoculated. When the cells reached approximately 70 %–80 % confluence, they were passaged to prevent differentiation of the muscle SCs.

2.5.3. STIM1-ASCs and SCs myogenic differentiation

ASCs, STIM1-ASCs and SCs were separately seeded into confocal dishes and subjected to myogenic differentiation using commercially available myogenic induction medium (CTCC-Y006, CHINA), with half-medium changes every other day. After 10 days, we performed MHC immunofluorescence staining on the cells from each group and calculated the myotube fusion rate, following the method described by PEDRO et al., which is the ratio of the number of nuclei within fused myotubes to the total number of nuclei [55].

2.5.4. Cytotoxicity test of adECM

The IGF-2-loaded adECM scaffold was immersed in DMEM at 37 °C for 72 h to prepare the extract as a conditioned medium. GFP-ASCs were seeded in 96-well plates at a density of 5000 cells/well and incubated overnight in growth medium (DMEM containing 10 % FBS) at 37 °C with 5 % humidified CO₂. The growth medium was then replaced with the conditioned medium. On days 1, 3, and 5, fresh DMEM medium containing 1/10 (v/v) CCK-8 solution was added to each well and incubated at 37 °C for 1 h. Absorbance was subsequently measured at 450 nm using a microplate reader.

2.5.5. Distribution and proliferation of ASCs inside adECM

The IGF-2-loaded adECM scaffold was placed in a 6-well plate. ASCs were seeded into the adECM by injecting 50 μL of the cell suspension (5 × 10³ cells) into the vascular pedicle. After 4–6 h of pre-cultivation, 2 mL of DMEM medium containing 10 % FBS was gently added to the 6-well plate. The culture medium was refreshed every other day. At 1, 3, and 7 days post-cell inoculation (n = 3 per time point), the cells were stained by with Calcein AM/PI (Keygen, BioTECH, China) and scanned by layer using a confocal laser scanning microscope (TCS-SP8 DIVE, Leica, Germany) to construct three-dimensional (3D) view graphs and observe the 3D distribution and proliferation of ASCs within the IGF-2-loaded adECM scaffold. Additionally, the number of cells in each 3D view graph was quantified using Image J software.

2.5.6. Myogenic differentiation of STIM1-ASCs inside IGF-2 loaded adECM

First, supernatants collected from the co-culture of different IGF-2-loaded adECM scaffolds with macrophages were mixed with commercially available myogenic induction medium at a 1:1 vol ratio to create various myogenic induction media (adECM + IGF-2 (0 ng/mL), adECM + IGF-2 (5 ng/mL), adECM + IGF-2 (10 ng/mL), and adECM + IGF-2 (20 ng/mL)). Next, 2 × 10⁵ STIM1 ASCs were injected into adECM scaffolds through the vascular pedicle and cultured in a 6-well plate containing 2 mL DMEM medium. After 24 h, the medium was replaced with the myogenic induction medium for myogenic differentiation, with half the medium refreshed every other day. On day 10, the cells within the adECM scaffolds were stained with a mouse monoclonal antibody against MHC (1:10,000, #ab37484, Abcam, USA), followed by incubation with goat anti-mouse DyLight 647 (1:500, #ab150115, Abcam, USA). After staining with Hoechst33342 (D0030, Beijing Solarbio Science & Technology Co., Ltd. China) for 30 min, the cells were visualized using a confocal laser scanning microscope.

Additionally, STIM1-ASCs in the adECM were lysed using RIPA lysis buffer supplemented with a protease inhibitor cocktail and detected the concentration by BCA Protein Assay Kit (C503061, Beijing Boxbio Science & Technology Co.,Ltd.). After mixing with SDS loading buffer and heating at 95 °C for 5 min, 15 μL samples were loaded onto an One-Step PAGE Gel (E303-01, Vazyme, Biotech Co., Ltd, China) and subsequently electrotransferred onto PVDF membranes (LS-PVDF-045-Z, LANSO, China). The membranes were blocked with TBST containing 5 % nonfat dry milk at room temperature for 1.5 h and incubated with primary antibodies at the indicated dilutions overnight at 4 °C. The primary antibodies used were as follows: Desmin (#ab32362, Abcam, UK, 1:100,000), MHC (#ab37484, Abcam, USA, 1:10,000), GAPDH (#181602, Abcam, UK, 1:1000), and MYH2 (#ab124973, Abcam, USA, 1:5000). Following incubation with a goat anti-rabbit secondary antibody for 1.5 h at room temperature, the PVDF membranes were visualized using an ECL chemiluminescence kit (E1050, LabLead, China) and imaged with the TANON 5200 Multi imaging system (TANON, Beijing, China).

2.6. Masseter VML injury reconstruction with STIM1-ASCs recellularized IGF-2-loaded adECM

2.6.1. Masseter VML injury model

Healthy 8-week-old male Sprague Dawley rats (Vital River, Beijing, China) were used to establish a masseter volumetric muscle loss (VML) model. The animal protocols were approved by the Peking University Biomedical Ethics Committee (No. A2024081). All in vivo experiments were conducted in accordance with the National Institutes of Health guidelines for the care and use of laboratory animals. Isoflurane anesthesia was used to sedate the animals.

Before surgery, the SD rats were placed in a sealed container connected to a gas anesthesia system and anesthetized with 3 % isoflurane. After ensuring proper anesthesia, the animals were positioned laterally, with the neck extended to a neutral position, and the nostrils placed in the anesthetic cone. During the procedure, the concentration of isoflurane was adjusted to 2 % to maintain anesthesia. Hair at the surgical site, including the areas around the oral commissure, upper right ear, and mandibular angle, was removed using hair removal cream, and the area was disinfected with iodine tincture. A longitudinal incision of 3–4 cm was made along the right side of the face from the lower lip pad to the right ear line. The skin was then carefully separated and lifted, exposing the right masseter muscle and the two facial nerve branches (buccal and mandibular branches) [51,56]. A 1 cm transverse incision was made on the fascia covering the masseter, and the space between the fascia and masseter was bluntly dissected, ensuring the integrity of the remaining fascia. A sterile biopsy needle was used to create a 5 mm diameter, 5 mm deep circular muscle defect (the excised muscle was weighed and recorded), ensuring the defect was located between the facial nerve buccal and mandibular branches without damaging the nerves. AdECM scaffolds were then implanted into the circular defect site of the masseter. Finally, the deep fascia and skin incisions were sutured with 5-0 absorbable sutures. Within 5 days post-surgery, ibuprofen was added to the drinking water to alleviate pain and inflammation at the surgical site [50]. To minimize selection bias, group allocation was concealed by an independent researcher who was not involved in the experiment until the interventions were administered.

The animals were divided into 6 groups (n = 10 per group):

1)
Native group: No masseter VML defect.
2)
Untreated group: Masseter VML defect without adECM implantation.
3)
adECM group: The adECM scaffold was implanted into the masseter VML defect.
4)
adECM loading with IGF-2 group (AI): IGF-2-loaded adECM was implanted into the masseter VML defect.
5)
STIM1-ASCs recellularized adECM group (AS): The adECM scaffold recellularized with 2 × 10⁵ STIM1-ASCs was implanted into the masseter VML defect.
6)
STIM1-ASCs recellularized IGF-2-loaded adECM group (ASI): IGF-2-loaded adECM scaffold recellularized with 2 × 10⁵ STIM1-ASCs was implanted into the masseter VML defect. The rats were weighed weekly, starting from the day of modeling (0, 7, 14, 21, 28, 35, and 42 days).

Following the transplantation of the various adECM scaffolds for 42 days, the masseter muscles from both sides of each rat were excised. The right side represented the modeling side. The muscle mass of the modeling side was weighed, and the masseter volume was calculated using the following equation: Volume (V) = (tissue length) × (tissue width)²/2 [50].

2.6.2. Histologic evaluation of masseter

At days 7 and 42 post-implantation, the rats were euthanized using an overdose of CO₂, and the mid-belly section of the right-side masseter muscle was excised to prepare paraffin sections. The sections were then stained with H&E, Sirius Red, Masson, and immunofluorescence. The myofiber diameter, number of centronucleated myofibers, and fiber cross-sectional area (FCSA) in the skeletal muscle defect were analyzed using Image J software.

Additionally, the sections embedded by OCT (AS82015, AMEKO, China) were incubated overnight at 4 °C with rabbit polyclonal antibodies against CD31 (#ab182981, Abcam, USA), MHC (#ab37484, Abcam, UK), Ki67 (#ab16667, Abcam, USA), Laminin (#PA1-16730, Thermo Scientific, USA), AChR (#RM2953, Biodragon, China), iNOS (#GB11119, Servicebio), CD163 (#GB15340, Servicebio), CD68 (#GB113109, Servicebio), and Pax3 (#ab180754, Abcam, USA). The following day, they were incubated with species-matched secondary antibodies conjugated to Alexa Fluor 488 (#ab150077, Abcam, USA) and Alexa Fluor 647 (#ab150115, Abcam, USA). Nuclei were counterstained with DAPI. Photometric analyses, including the positive areas of Sirius Red staining, CD31⁺, MHC⁺, CD68⁺/CD163⁺, CD68⁺/iNOS⁺, and Pax3⁺/Ki67⁺ markers, were conducted using Image J software on randomly selected fields from three slides per sample.

2.6.3. Imaging evaluation of masseter

Ultrasound imaging of the masseter muscle was performed 42 days post-implantation in the cross-sectional plane. After anesthetizing the rats, ultrasonic gel was applied to the hairless masseter area, with hair removal achieved using a depilatory cream. Images were captured bilaterally under identical acquisition settings in B-mode using the ultrasonic photoacoustic multi-mode imaging system (Vevo F2 LAZR-X, USA), equipped with a 30 MHz linear transducer.

2.6.4. Mastication rate measurement and eating behaviour analyses

Mastication behavior was analyzed based on previous studies [50]. Briefly, food was removed from the rats overnight, and 500 g of food was reintroduced the following day. The mastication behavior of each rat was recorded from a lateral view using a video camera (CANON, Tokyo, Japan) at a frame rate of 60 frames per second. Each chewing episode was defined as starting with maximum jaw closure. The number of consecutive chewing cycles (maximum jaw opening followed by complete closure) was counted over the next 59 consecutive frames to calculate the chewing rate (number of chewing cycles per second). Mastication rates were determined by analyzing three independent, consecutive chewing clips for each animal within 60 frames of video. The data represent the average chewing rate (chews per second) for each rat.

Additionally, on the day food was reintroduced, the rats were kept on the experimental platform for 12 h, from 7:00 a.m. to 7:00 p.m. A video camera was used to record their movements and activities. As the animals consumed food, the amount of food intake was monitored. At 7:00 p.m., the remaining food was weighed to calculate the amount of food consumed by each rat.

2.6.5. Contraction strength evaluation of masseter muscle

The contractile properties of the masseter muscle were measured using a Muscle Strip Myograph System (DMT820MS, Aarhus, Denmark) as described in previous studies [25,57]. Briefly, on day 42 post-implantation, the isolated masseter muscle was suspended in Krebs-Henseleit solution at 25 °C and oxygenated with 20 % O₂ and 5 % CO₂. After setting the initial length of the isolated muscle, twitch responses were recorded at 20 V and 100 Hz with a 5 ms pulse width using LabChart 7.2 (ADInstruments). To verify muscle viability, electrical stimulation was applied in the form of three tetanic waves at 20 V with a pulse width of 20 ms and an approximate runtime of 1 min. A rest period of 3 min was allowed after each stimulation to prevent muscle fatigue. The maximal force for each group was normalized to the weight of the isolated masseter muscle.

2.6.6. Bite force measurement

According to the previous protocol for maximal bite force testing. Briefly, we adapted the animals by placing them in individual cages 2 h prior to the test. Bite force was measured using the FlexiForce sensor (B201-L-8) and the Load and Force system from SICHIRAY (WUXI, CHINA). The animals were tested for 1 min without any painful stimuli, and the raw data was continuously recorded for subsequent analysis. Each rat underwent the test three times, with a 3-min interval between each trial. The maximum bite force (in grams) for each animal was recorded for each test, and the mean of all trials was then calculated.

2.7. Statistical analysis

All data are presented as the mean ± standard deviation (s.d.). Statistical analyses were performed using Prism 9.0 software (GraphPad Prism 9.5.1, USA). Comparisons between two groups were conducted using the t-test, while comparisons among multiple groups were performed using one-way ANOVA. A p-value of <0.05 was considered statistically significant (∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001).

3. Results

3.1. Preparation and characterization of adECM with vascular pedicle

The preparation method of vascularized adECM is illustrated in Fig. 1A. Adipose tissue with a vascular pedicle was harvested from the inguinal region of rats. Using a hollow needle inserted into the blood vessel, the adipose tissue was connected to a bioreactor and sequentially perfused with solutions of SDS, isopropanol, and deionized water. This process resulted in the final adECM scaffolds. The gross morphology of the adipose tissue before and after decellularization is shown in Fig. 1B. After decellularization, the adECM appeared white and translucent, with its overall shape largely preserved. H&E staining, Oil Red O staining, and DAPI staining (Fig. 1C–E) confirmed the absence of visible lipids, cells, or intracellular components in the adECM. Additionally, DNA concentration measurements revealed that the residual DNA content in the adECM was 2.7 ± 0.13 ng/mg (Fig. 1F), which is significantly lower than the international standard of 50 ng/mg [58]. These results indicate that our method effectively removes cells and intracellular components from adipose tissue, ensuring the production of high-quality adECM.

To verify whether the extracellular matrix (ECM) components of adipose tissue were fully preserved after decellularization, we first evaluated the retention of glycosaminoglycans (GAGs), a hydrophilic proteoglycan molecule critical for facilitating interactions between cells and between cells and the ECM [59]. Quantitative analysis and specific staining (Alcian blue staining) were utilized for this purpose. As shown in Fig. 1G, the GAGs content in adECM (0.72 ± 0.08 μg/mg) was slightly higher than that in adipose tissue (0.66 ± 0.07 μg/mg), although the difference was not statistically significant. Alcian blue staining indicated the presence of prominent GAGs in adECM (Fig. 1H). We also used Masson staining to observe other protein components of the ECM. As shown in Fig. 1I, the proportion of collagen in adECM was significantly higher than that in adipose tissue (per field). Further SDS-PAGE analysis confirmed that the collagen in adECM was predominantly type I collagen, with protein bands nearly identical in position to those of commercially available rat tail type I collagen (Fig. 1J). This finding aligns with previous reports in the literature, which state that type I collagen accounts for over 90 % of the collagen in adipose tissue ECM [34,60]. Additionally, scanning electron microscopy (SEM) revealed that the microstructure of the adECM closely resembled the 3D microstructure of native adipose tissue (Fig. 1K). However, the pores in adECM were relatively smaller, likely due to the removal of cells and the collapse or compression of the ECM structure during the decellularization process.

The swelling ratio of adECM determines its ability to facilitate material exchange within the bioscaffold. During a 6-h swelling performance test, we observed that the wet weight of adECM at each time point was consistently higher than that of native adipose tissue (Fig. 1L). This suggests that the prepared adECM exhibits superior water retention, likely due to its extensive internal porosity. To evaluate the stability of adECM, equal masses of adECM and native adipose tissue were immersed in a type I collagenase solution (0.05 mg/mL). As shown in Fig. 1M, the degradation patterns of adECM and native adipose tissue were similar at each time point, indirectly confirming that type I collagen, a critical ECM component, was fully preserved during the preparation process.

The mechanical properties of the adECM scaffold are crucial for tissue regeneration [61]. We therefore assessed the compressive and tensile properties of adECM. From the stress-strain curves of compressive performance (Fig. 1N), the slope indicates that the compressive modulus of adECM (0.183 ± 0.016 10 kPa) was significantly lower than that of native adipose tissue (0.524 ± 0.008 10 kPa, p < 0.001), which is likely due to the removal of cells within the adECM structure. Conversely, analysis of the stress-strain curves for tensile performance and their quantitative slope measurements (Fig. 1O) revealed that the tensile modulus of adECM (0.008 ± 0.002 10 kPa) was significantly higher than that of native adipose tissue (0.005 ± 0.001 10 kPa). According to the literature [62], this result can be attributed to the higher concentration of type I collagen in adECM per unit volume. In conclusion, we successfully prepared vascularized adECM using a bioreactor. The internal cells and their contents were thoroughly removed, while key extracellular matrix components, such as GAGs and type I collagen, were fully retained. Additionally, adECM exhibited excellent swelling capacity, stability, and mechanical properties, all of which are critical for supporting the metabolism and differentiation of cells within the adECM scaffold.

We also evaluated the biosafety of the adECM scaffold using live/dead cell staining assays. Specifically, we seeded 1 × 10⁴ ASCs into the adECM scaffold via the vascular pedicle and performed Calcein-AM and PI staining on the ASCs within the scaffold on days 3 and 7. The results, shown in Fig. S1, indicate that on both day 3 and day 7, the ASCs in the scaffold exhibited good growth (green), with no significant PI (red) positive cells observed. These results suggest that the adECM scaffold has no significant impact on the cell viability of ASCs and demonstrates good biosafety.

3.2. The IGF-2 loaded adECM induce the polarization of macrophage into M2 macrophage phenotype

We used a transwell co-culture system to study the regulatory effects of IGF-2-loaded adECM on the polarization of macrophages (Fig. 2A). First, following methods reported in the literatures [63,64], different concentrations of IGF-2 (0 ng/mL, 5 ng/mL, 10 ng/mL, and 20 ng/mL) were incubated with 10 mg of adECM in PBS for 24 h to prepare various IGF-2-loaded adECM constructs, including adECM alone and adECM + IGF-2 at 5 ng/mL, 10 ng/mL, and 20 ng/mL. We initially quantified the IGF-2 loading efficiency of adECM at different IGF-2 concentrations using ELISA, and the results are presented in Table 1. After 24 h of incubation, the loading efficiencies of IGF-2 for adECM in the 5 ng/mL, 10 ng/mL, and 20 ng/mL groups were 59.83 ± 2.87 %, 79.27 ± 0.64 %, and 89.71 ± 0.35 %, respectively. Next, the IGF-2 release profiles were assessed by immersing the IGF-2-loaded adECM constructs in PBS and measuring the IGF-2 concentration in the solution at various time points (30 min, 1, 2, 3, 4, 5, 6, and 7 days). As shown in Fig. 2B, all groups exhibited a burst release of IGF-2 during the first 30 min, with a significantly higher release rate compared to other time points. Specifically, in the adECM + IGF-2 (5 ng/mL) group, the amount of IGF-2 released during this period was nearly 45 % of the total loaded amount. Additionally, the adECM + IGF-2 (5 ng/mL) group completely released IGF-2 by day 5, while the adECM + IGF-2 (10 ng/mL) group released nearly all IGF-2 (99.06 %) by day 7. In contrast, the adECM + IGF-2 (20 ng/mL) group released only 74.36 % of its total IGF-2 by day 7. These results suggest that higher IGF-2 loading concentrations result in a more sustained release profile of IGF-2.

Table 1.

IGF-2 loading on adECM scaffold.

adECM	IGF-2	Drug loading (%)
10 mg	5 ng/mL	59.83 ± 2.87
10 mg	10 ng/mL	79.27 ± 0.64
10 mg	20 ng/mL	89.71 ± 0.35

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Data are shown as the mean ± s.d. (n = 3).

The macrophages used in the experiment were induced from rat femoral bone marrow mononuclear cells. We evaluated the classical markers of macrophages (CD11b and F4/80) using immunofluorescence staining, as shown in Fig. S2. The mature macrophages expressed both CD11b and F4/80 cell markers, indicating the successful isolation of macrophages. Next, we co-cultured different IGF-2-loaded adECM scaffolds with macrophages. Untreated macrophages were used as the Control group, while macrophages treated with the M2 macrophage-promoting agent IL-4 served as the Positive control. After 3 days of co-culture, flow cytometry was used to analyze the proportions of M1 and M2 macrophages in each group. Macrophages were gated using the F4/80 marker, and the proportions of M1 (CD80) and M2 (CD206) macrophages were subsequently assessed (Fig. 2C and Fig. S3). As shown in Fig. S4A and S4B, compared to the Control group, the proportion of CD80 (M1) macrophages significantly decreased only in the IL-4-treated group, with no notable increase or decrease in CD80 levels observed in the other groups. In contrast, treatment with adECM + IGF-2 (10 ng/mL) and adECM + IGF-2 (20 ng/mL) significantly increased the proportion of M2 (CD206) macrophages to 54.46 ± 1.25 % and 51.57 ± 0.49 %, respectively, compared to the Control group (p < 0.01, Fig. 2D). Additionally, we performed RT-qPCR to measure the expression levels of M1-related genes (IL-1β, TGF-α) and M2-related genes (IL-10, Arg-1). The results, shown in Fig. S4C and S4D, revealed that only the adECM + IGF-2 (10 ng/mL) group significantly reduced the expression levels of IL-1β and TGF-α, with effects comparable to those in the IL-4-treated group. Moreover, the adECM + IGF-2 (10 ng/mL) group exhibited significantly higher expression levels of IL-10 and Arg-1 compared to the Control group (p < 0.05), whereas other treatment groups (except IL-4) showed no statistically significant differences (Fig. 2E and F). These results suggest that IGF-2-loaded adECM regulates macrophage polarization toward the M2 phenotype through IGF-2 release, with adECM + IGF-2 (10 ng/mL) and adECM + IGF-2 (20 ng/mL) demonstrating the strongest effects in promoting M2 macrophage activation. However, bare adECM alone neither promoted M1 nor M2 macrophage polarization, contrary to reports from other studies suggesting that adECM itself can alter macrophage phenotypes [65].

Study have shown that specific metabolic pathways in macrophages are closely associated with their phenotypes and functions [66]. Generally, M1 macrophages primarily rely on aerobic glycolysis and exhibit impaired mitochondrial oxidative phosphorylation (OXPHOS), while M2 macrophages depend on mitochondrial OXPHOS for energy metabolism. Based on this, we investigated the effects of adECM + IGF-2 on the glucose metabolism of macrophages. Seahorse analysis (Fig. 2G) revealed that compared to the Control group, adECM + IGF-2 (5 ng/mL), adECM + IGF-2 (10 ng/mL), and adECM + IGF-2 (20 ng/mL) significantly downregulated Extra Cellular Acidification Rate (ECAR) included Glycolysis, Glycolytic capacity, Glycolytic reserve of macrophages. These trends were consistent with the IL-4 group, which also reduced macrophage's Glycolytic function (Fig. 2H). Conversely, adECM + IGF-2 (5 ng/mL), adECM + IGF-2 (10 ng/mL), and adECM + IGF-2 (20 ng/mL) upregulated mitochondrial oxygen consumption rate (OCR) in macrophages, with significant improvements observed in basal respiration, maximal respiration, spare respiratory capacity (Fig. 2I). Notably, adECM + IGF-2 (10 ng/mL) demonstrated the strongest promotion of Mitochondrial respiration, reaching levels comparable to those observed in the IL-4 group (Fig. 2J). In summary, according to the results of flow cytometry and Seahorse analysis, we confirmed that IGF-2-loaded adECM regulates macrophage polarization via IGF-2 release. More importantly, IGF-2 appears to facilitate the mitochondrial respiration of macrophages through metabolic reprogramming, ultimately driving macrophages toward the M2 phenotype.

To further clarify the mechanism by which IGF-2 alters mitochondrial metabolism in macrophage, we performed RNA sequencing on macrophages treated with adECM + IGF-2 (10 ng/mL) and compared the results with the RNA-seq data of macrophages from the adECM group, as shown in Fig. 3. From the principal component analysis (PCA) plot, we can observe that both samples cluster well within their respective groups, and the distance between the two samples is relatively large, indicating a significant difference between the adECM + IGF-2 (10 ng/mL) group and adECM group (Fig. 3A). Compared to the adECM group, the volcano plot and heatmap of differentially expressed genes show that the number of up-regulated and down-regulated genes in the adECM + IGF-2 (10 ng/mL) group were 436 and 654, respectively (|Log2FC| >1 and p < 0.05, Fig. 3B and C). Next, GO analysis of the up-regulated genes revealed that the macrophages were mainly involved in biological processes such as “regulation of immune effector process”, “cytokine-mediated signaling pathway”, and “chemotaxis among other immune regulatory processes” (Fig. 3D). Interestingly, we noted glucose metabolism processes among the top 5 biological processes, which appears to be related to glucose metabolism in macrophage mitochondria, suggesting that adECM + IGF-2 may alter mitochondrial metabolism to regulate macrophage polarization. Furthermore, to identify the signaling pathways through which adECM + IGF-2 regulates M2 polarization in macrophages, we performed KEGG analysis on the upregulated differentially expressed genes (Fig. 3E). The results showed that signaling pathways such as “cytokine-cytokine receptor interaction”, “PI3K-Akt signaling pathway” and “calcium signaling pathway” were activated, with the PI3K-Akt signaling pathway showing the greatest enrichment and most significant difference in differentially expressed genes.

Fig. 3 — **RNA-Seq to investigate the mechanism by which IGF-2 loaded adECM regulates M2 macrophage activation.** (A) Principal component analysis (PCA). (B) Volcano plot of differentially expressed genes (DEGs). (C) Heatmap of DEGs. (D) Top 10 up-regulated biological processes revealed through GO enrichment analysis. (E) The top-ranked signaling pathway analyzed by the KEGG platform in the adECM + IGF-2 (10 ng/mL) group. (F) The extracellular acidification rate (ECAR) in Control group, adECM + IGF-2 (10 ng/mL) group, adECM + IGF-2+LY294002 group and IL-4 group was measured using the Seahorse XF96 Extracellular Flux Analyzer, and statistical results for Glycolysis, Glycolytic capacity and Glycolytic reserve are shown in (G). Abbreviations: 2-DG, 2-deoxy-D-glucose; Rot/AA, rotenone/antimycin A. n = 5 per group. (H) The oxygen consumption rate (OCR) in Control group, adECM + IGF-2 (10 ng/mL) group, adECM + IGF-2+LY294002 group and IL-4 group was detected by the Seahorse XF96 Analyzer and statistical results for basal respiration, maximal respiration, and spare respiratory capacity are shown in (I). Abbreviations: FCCP, carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone; Rot/AA, rotenone/antimycin A. n = 5 per group. (J) The protein levels of PI3K-AKT pathway were evaluated by western blotting after cells were treated by adECM + IGF-2 (10 ng/mL) and adECM + IGF-2+LY294002. (K) Evaluation of mRNA expression of Oxidative phosphorylation enzymes after cells were treated by adECM + IGF-2 (10 ng/mL) and adECM + IGF-2+LY294002, n = 5 per group. All data are presented as mean ± s.d. Statistical significance was determined using one-way ANOVA: ∗∗∗P < 0.001, ∗∗P < 0.01, ∗P < 0.05, compared with the Control group.

After that, we intervened with macrophages using a PI3K-Akt pathway inhibitor (LY294002) to create an adECM + IGF2+LY294002 group. The results showed that under LY294002 intervention, the elevated levels of phosphorylated PI3K (p-PI3K) and phosphorylated AKT (p-AKT) in macrophages (observed in the adECM + IGF2 group) were significantly reduced (Fig. 3J). Additionally, the cellular acidification rate (ECAR) of macrophage mitochondria in the adECM + IGF2+LY294002 group was significantly increased, with statistical significance compared to the adECM + IGF2 group (P < 0.001, Fig. 3F–G). However, the mitochondrial oxygen consumption rate (OCR) of macrophages in the adECM + IGF2+LY294002 group was markedly decreased, showing a significant difference compared to the adECM + IGF2 group (Fig. 3H–I). Furthermore, under LY294002 treatment, the reduction in macrophage OCR/ECAR ratio was accompanied by decreased mRNA expression levels of ATP synthase F1 subunit alpha (ATP5A1), NADH dehydrogenase (Ndufb5), and Cytochrome c oxidase subunit IV isoform 1 (COX4I1), all of which are key enzymes involved in mitochondrial oxidative phosphorylation (Fig. 3K). These results suggest that adECM + IGF2 regulates macrophage metabolic reprogramming through the PI3K-Akt signaling pathway, enhancing mitochondrial oxidative phosphorylation and ultimately promoting M2 macrophage polarization.

3.3. The IGF-2 loaded adECM regulates fibroblast function by promoting M2 macrophage activation

To determine whether the IGF-2 loaded adECM regulates fibroblast function by modulating M2 macrophage activation, we collected the supernatant from different adECM + IGF-2 co-cultures with macrophages and mixed it with fibroblast culture medium at a 1:1 ratio to create a conditioned medium for studying fibroblast functions such as proliferation and migration. First, we performed a scratch assay on fibroblasts. As shown in Fig. 4A and B, in the representative images of fibroblasts at 6 h and 9 h, the fibroblasts in the adECM + IGF-2 (5 ng/mL) group proliferated significantly faster than those in the adECM + IGF-2 (0 ng/mL) group. On the other hand, the fibroblasts in the adECM + IGF-2 (10 ng/mL) and adECM + IGF-2 (20 ng/mL) groups proliferated more slowly than those in the adECM + IGF-2 (0 ng/mL) group. We used the scratch closure rate as an indicator of fibroblast proliferation rate and quantitatively analyzed the proliferation rates of each group at different time points (6 h and 9 h). The results showed that the proliferation rate in the adECM + IGF-2 (10 ng/mL) group was significantly lower than that in the adECM + IGF-2 (0 ng/mL) group at both time points, with a statistically significant difference (p < 0.05). Although the proliferation rate in the adECM + IGF-2 (20 ng/mL) group was also lower than that in the adECM + IGF-2 (0 ng/mL) group, no statistically significant difference was observed between them (Fig. 4C).

Fig. 4 — **IGF-2-loaded adECM scaffolds alter the function of human normal skin fibroblasts by promoting M2 macrophage polarization.** (A) Fibroblasts scratch assay: Fibroblasts were treated with different conditioned media, and images of the scratch were taken every 3 h. The remaining area of the scratch was indicated by the yellow-green color, Scale bar = 400 μm. (B) Time-scratch closure rate curves of fibroblasts under different conditioned medium interventions, showing the time taken for complete scratch closure. (C) Quantitative analysis of the proliferation rate of fibroblasts in different groups at 6 h and 9 h. (D) Representative images of fibroblast migration ability assessed by Transwell assay under different conditioned medium interventions, Scale bar = 100 μm. (E) Quantitative analysis of the effect of different conditioned media on fibroblast migration ability. (F) RT-qPCR was used to detect the expression levels of COL1A1 and COL3A1 genes in fibroblasts of each group. Data are presented as mean ± s.d. (n ≥ 3), Statistical significance was determined using one-way ANOVA: ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001, compared with the adECM + IGF-2 (0 ng/mL) group; while ^#P < 0.05, compared with the adECM + IGF-2 (20 ng/mL) group.

Next, we performed a transwell experiment to evaluate the migration ability of fibroblasts. After 24 h of culture, crystal violet staining showed that the fibroblast migration through the transwell membrane was the least in the adECM + IGF-2 (10 ng/mL) group, followed by the adECM + IGF-2 (20 ng/mL) group (Fig. 4D). Further quantitative statistical analysis revealed that, compared to the adECM + IGF-2 (0 ng/mL) group, the cell migration rate was significantly reduced in both the adECM + IGF-2 (10 ng/mL) and adECM + IGF-2 (20 ng/mL) groups. Among them, the fibroblasts in the adECM + IGF-2 (10 ng/mL) group showed the least migration, significantly fewer than those in the adECM + IGF-2 (20 ng/mL) group, and the difference was statistically significant (Fig. 4E). Additionally, we conducted RT-qPCR experiments to evaluate the collagen secretion function of fibroblasts. As shown in Fig. 4F, fibroblasts in the adECM + IGF-2 (5 ng/mL) group expressed significantly higher levels of COL1A1 and COL3A1 genes compared to the adECM + IGF-2 (0 ng/mL) group (p < 0.05). However, fibroblasts in the adECM + IGF-2 (10 ng/mL) and adECM + IGF-2 (20 ng/mL) groups expressed significantly lower levels of COL1A1 and COL3A1 genes than those in the adECM + IGF-2 (0 ng/mL) group (p < 0.001), with no statistical difference between the two groups. Based on the above experimental results, we conclude that the IGF-2 loaded adECM can regulate the activation of M2 macrophages, thereby affecting the proliferation, migration, and collagen secretion functions of fibroblasts. Among them, the fibroblast functions in the adECM + IGF-2 (10 ng/mL) and adECM + IGF-2 (20 ng/mL) groups were significantly inhibited.

3.4. Myogenic differentiation of STIM1-ASCs when in IGF-2 loaded adECM

To investigate whether STIM1-ASCs can undergo myogenic differentiation within the IGF-2-loaded adECM bioscaffold, and whether IGF-2 in the scaffold can enhance the myogenic differentiation efficiency of STIM1-ASCs through macrophage polarization regulation, we performed the following experiments. First, We used flow cytometry to detect the expression of surface markers (CD11b, CD29, CD34, CD45, CD44, CD73, CD90) on ASCs to evaluate whether the obtained ASCs comply with the international standards. The results, as shown in Fig. S5, revealed that the expression of CD11b (0.1 %), CD34 (0.12 %), and CD45 (0.1 %) was negative, while the expression of CD29 (95.75 %), CD44 (96.87 %), CD73 (99.92 %), and CD90 (99.82 %) was positive. These results suggest that the cultured ASCs have high purity and comply with the international standards for mesenchymal stem cell identification [67]. Next, we constructed the STIM1-ASCs cell model using lentiviral transfection technology. Then, supernatants from co-cultures of different adECM + IGF-2 scaffolds and macrophages were collected and mixed with myogenic induction medium at a 1:1 vol ratio to prepare the conditional myogenic induction medium. Finally, STIM1-ASCs were seeded into the adECM scaffold and cultured in the conditional myogenic induction medium, as illustrated in the schematic diagram (Fig. 5A).

After lentiviral transfection, we validated the expression of STIM1 protein in STIM1-ASCs using Western blot. The results (Fig. 5C) showed a significant upregulation of STIM1 protein in STIM1-ASCs, confirming the successful construction of STIM1-ASCs. Furthermore, the myogenic differentiation potential of STIM1-ASCs was assessed. After 10 days of myogenic induction under 2D in vitro culture conditions, immunofluorescence staining for MHC (a marker of mature myogenic differentiation [6]) revealed a substantial increase in multinucleated myotubes in the STIM1-ASCs group compared to the control group (GFP-ASCs) (Fig. 5B). Western blot analysis further confirmed that STIM1-ASCs exhibited significantly higher MHC protein expression (Fig. 5C). In conclusion, we successfully constructed STIM1-ASCs, which exhibit high myogenic differentiation potential.

In addition, we compared the myogenic differentiation efficiency of ASCs, STIM1-ASCs, and SCs in vitro. After 10 days of myogenic induction, we performed MHC immunofluorescence staining on the cells from each group and calculated the myotube fusion rate, following the method described by PEDRO et al., which is the ratio of the number of nuclei within fused myotubes to the total number of nuclei [55]. The results, shown in Fig. S6, indicated that ASCs exhibited very low levels of MHC expression and maintained a mesenchymal stem cell-like polygonal shape, with no mature myotubes formed. This finding is consistent with conclusions reported in the literature, where the myogenic differentiation efficiency of ASCs is very low, even less than 1 % [68,69]. After lentiviral transfection of STIM1, the myogenic differentiation efficiency of ASCs significantly improved. The myotube fusion rate of STIM1-ASCs reached 43.55 %. However, compared to SCs, which had a myotube fusion rate of 80.40 %, the myotube fusion rate of STIM1-ASCs remains relatively low, with a statistically significant difference between the two groups (P < 0.001).

Futhermore, we also conducted Transmission Electron Microscopy (TEM) observations on STIM1-ASCs induced by myogenic differentiation media. As shown in Fig. S7A, after 10 days of myogenic induction, the cells gradually adopted an elongated shape and aligned in a specific direction (indicated by red double arrows). Additionally, bundles of fibers with varying thickness were observed in the cytoplasm, with the direction of these bundles roughly aligning with the cell's long axis (Fig. S7B, indicated by red single arrow). Moreover, TEM analysis revealed occasional “myosacs” structures in the STIM1-ASCs (Fig. S7C, indicated by a red ∗). These structures occupied the main area of the cytoplasm, leading to the aggregation of mitochondria and other organelles at the cell edges [70]. Interestingly, as shown in Fig. S7D, typical nuclear indentation and fissure structures (indicated by a yellow ∗) were observed in the cells. These features are only present in mature myotubes or muscle fibers, and it is believed that their appearance may be related to potential contractile functions [71].

Before seeding ASCs into the adECM scaffold, we collected the adECM extract and co-cultured it with ASCs to evaluate its cytocompatibility. The CCK-8 assay showed that after 5 days of culture, the viability of ASCs remained unaffected, with OD values displaying an upward trend (Fig. 5D). Subsequently, ASCs were seeded into the adECM via its vascular pedicle. 3D fluorescence imaging revealed (Fig. 5E and F) that the number of ASCs (stained by Calcein-AM, green) significantly increased over time (from day 1 to day 7), proliferating from 664.2 ± 81.0 cells/mm² to 2372.3 ± 373.6 cells/mm², and with no significant PI (red) positive cells observed (Fig. S1). Moreover, the cells were evenly distributed throughout the scaffold. These results indicate that the adECM bioscaffold demonstrates excellent cytocompatibility. After vascular pedicle cell seeding, STIM1-ASCs distributed uniformly within the adECM and exhibited robust proliferation.

After confirming that ASCs can proliferate normally within the scaffold, we inoculated STIM1-ASCs into the scaffold using the same method, and induced STIM1-ASCs myogenic differentiation using the previously prepared conditional myogenic induction medium. The myogenic differentiation efficiency of the STIM1-ASCs within the recellularized adECM was evaluated via MHC immunofluorescence staining and Western blot analysis. As shown in Fig. 5G, MHC immunofluorescence staining indicated that STIM1-ASCs in the adECM + IGF-2 (0 ng/mL) group expressed MHC protein, but at a relatively low level. Compared to the adECM + IGF-2 (0 ng/mL) group, the fluorescence intensity of MHC protein significantly increased in the adECM + IGF-2 (5 ng/mL) and adECM + IGF-2 (10 ng/mL) groups. However, the expression level of MHC protein showed no notable increase in the adECM + IGF-2 (20 ng/mL) group. Western blot analysis further quantified the expression levels of myogenesis-related proteins (MHC, MYH2, and Desmin) across the groups. Consistent with the immunofluorescence results, the adECM + IGF-2 (5 ng/mL) and adECM + IGF-2 (10 ng/mL) groups exhibited the highest expression levels of MHC, MYH2, and Desmin proteins compared to the adECM + IGF-2 (0 ng/mL) group, while the adECM + IGF-2 (20 ng/mL) group showed no significant differences (Fig. 5H and I). In summary, adECM + IGF-2 (5 ng/mL) and adECM + IGF-2 (10 ng/mL) significantly enhanced the myogenic differentiation efficiency of STIM1-ASCs. Combining these findings with the results from flow cytometry and Seahorse analyses, we conclude that adECM + IGF-2 (10 ng/mL) effectively promotes macrophage polarization toward the M2 phenotype and then enhances the myogenic differentiation of STIM1-ASCs. Therefore, in subsequent animal experiments, we utilized the recellularized adECM + IGF-2 (10 ng/mL) scaffold seeded with STIM1-ASCs as the tissue engineering bioscaffolds.

3.5. STIM1-ASCs recellularized IGF-2 loaded adECM treatment for masseter VML

To evaluate the muscle regeneration effects of the aforementioned tissue-engineered bioscaffolds on craniofacial VML, we established a muscle defect (5 mm in diameter and 5 mm in height) in the right masseter muscle of rats, following protocols described in previous studies [2]. Different adECM scaffolds were transplanted into the injury site, including the alone adECM scaffold (adECM group), IGF-2-loaded adECM scaffold (AI group), STIM1-ASCs recellularized adECM scaffold (AS group), and STIM1-ASCs recellularized IGF-2-loaded adECM scaffold (ASI group). Animals without any treatment for the VML defect served as the Control group, while animals with no VML defect were designated as the Native group. Starting from day 1 post-surgery, the average body weight of each group was recorded at 7-day intervals. On day 42, the regenerated masseter muscles were assessed through histological staining, imaging (ultrasound), and functional indicators (muscle strength). Additionally, to investigate macrophage polarization and satellite cell proliferation within the defect site, dual immunofluorescence staining was performed on the masseter muscles of all groups on day 7. The complete workflow for the animal experiment is illustrated in Fig. 6A.

The weekly body weight growth curves of the experimental groups (Fig. 6C) revealed differences in weight gain rates among the groups. The Control group and adECM group exhibited slower weight gain, while the Native group showed the fastest growth. Nonetheless, throughout the entire experimental period, no animals died or displayed persistent signs of illness. All animals demonstrated continuous weight gain, indicating that the tissue-engineered scaffolds used in this study had no adverse effects on animal health. Additionally, all adECM scaffolds exhibited good biocompatibility. At the end of the experiment (day 42), bilateral masseter muscle specimens were collected from each group for analysis. Gross morphological observations of the masseter muscles (Fig. 6B) showed that the right masseter muscles of the Control group and adECM group were visibly smaller than the normal left masseter muscles. Using the volume calculation formula: Volume (V) = (tissue length) × (tissue width)²/2 [50], the volumes of the right masseter muscles were quantitatively assessed. The results showed no significant differences in muscle volume between the adECM group, AI group, and AS group compared to the Control group. However, the masseter muscle volume in the ASI group was significantly increased compared to the Control group, with statistical significance (p < 0.05, Fig. 6D). Similarly, the weights of the right masseter muscles followed a pattern consistent with their volumes. The ASI group demonstrated a significant increase in muscle weight compared to the Control group, while the other treatment groups did not show significant increases (Fig. 6E).

To further investigate muscle regeneration within the adECM scaffolds, H&E staining was performed on the masseter muscles of each group, as shown in Fig. 6F. After 42 days of treatment, no distinct muscle fiber structures were observed in the injury site of the Control group, although vascular-like cavity structures were present. Similarly, the adECM group exhibited no detectable muscle fiber structures, consistent with previous study demonstrating that alone adECM transplantation alone cannot regenerate muscle fibers [72]. In the AI group and AS group scaffolds, a small number of regenerated muscle fibers and nascent vascular structures were observed. In contrast, the ASI group exhibited a substantial presence of muscle fiber signals within the scaffold. Among these newly formed muscle fibers, some remained immature, characterized by centrally located nuclei. Quantitative analysis of the regenerated muscle fibers, including their diameter, the number of mature muscle fibers (nuclei located at the periphery), and the cross-sectional area (FCSA) of the fibers, was performed based on established methods [73]. The diameter, number of mature muscle fibers, and FCSA of regenerated fibers in the AS group and ASI group scaffolds were significantly higher than those in the Control group. Notably, the ASI group contained more mature muscle fibers than the AS group, but no significant differences in fiber diameter or FCSA were observed between these two groups (Fig. 6G). Since vascularization is critical for the regeneration of muscle fibers [6], CD31 immunofluorescence staining was used to evaluate vascular structures in the scaffolds, and the average fluorescence intensity was quantified (Fig. 6H and I). Strong CD31 fluorescence signals were detected in the scaffolds of the AI group, AS group, and ASI group, with no statistically significant differences among these groups. This result aligns with the findings from H&E staining, suggesting that enhanced vascularization supports the regeneration and metabolism of muscle fibers.

The results of MHC and Laminin dual immunofluorescence staining (Fig. 6J and K) revealed extensive muscle fiber formation (MHC, green fluorescence) and substantial Laminin deposition (red fluorescence) within the scaffolds of the ASI group. The MHC fluorescence intensity in the AI group and AS group was noticeably weaker than in the ASI group, although both groups exhibited statistically significant differences compared to the Control group. Similarly, no significant MHC fluorescence signal was detected in the adECM group, consistent with the H&E staining results. Neural regeneration in the muscle injury area is an important prerequisite for improving muscle function and preventing muscle atrophy. Therefore, we performed immunofluorescence staining of acetylcholine receptors (AChR) at the neuromuscular junctions in the scaffold and muscle. The results showed that the AChR positive rate in the AI group and ASI group was significantly higher than that in the Control group, and the AChR positive rate in the AS group was also higher than that in the Control group (Fig. 6L). Quantitative analysis revealed that the fluorescence intensity of AChR in the AI group and ASI group was significantly higher than in the Control group, with no statistical difference between the AI and ASI groups. Although the fluorescence intensity of AChR in the AS group was higher than that in the Control group, no significant statistical difference was observed between them (Fig. 6M).

To assess the ability of different adECM scaffolds to alleviate late-stage fibrotic scar formation following muscle injury, Sirius Red staining was performed on the masseter muscles of each group (Fig. 6N and O, Figures S8). Compared to the Control group, the Sirius Red-positive area in the AI group, AS group, and ASI group was significantly reduced, indicating a marked reduction in muscle fibrosis. Importantly, the ASI group exhibited the smallest Sirius Red-positive area, with a statistically significant difference compared to the AI group and AS group (p < 0.01). Although the AI group showed less fibrosis than the AS group, the difference was not statistically significant.

3.6. STIM1-ASCs recellularized IGF-2 loaded adECM promotes Pax3⁺ proliferation and M2 polarization in vivo

According to previous studies, approximately 1 week after muscle injury marks a critical time window for satellite cell activation and macrophage polarization in the muscle microenvironment [65,74,75]. To investigate the potential mechanisms by which the tissue-engineered bioscaffolds promotes muscle regeneration and reduces fibrosis in vivo, we performed histological sectioning and immunofluorescence staining on day 7 post-scaffold transplantation. As shown in Fig. 7A, signals of newly formed muscle fibers were already observed within the ASI group scaffold, whereas no newly formed muscle tissue was detected in the scaffolds of other treatment groups. Based on the mechanism by which IGF-2 facilitates muscle repair, it mobilizes residual satellite cells (SCs) in the early stages of injury, reactivating them from their quiescent state and promoting their proliferation to replenish the local satellite cell population [31,34]. To verify this, we analyzed and quantified Pax3⁺/Ki67⁺ cells in the injury regions of each group (Pax3 being a marker for satellite cells and Ki67 a marker for cell proliferation [76]). As shown in Fig. 7B, the number of Pax3⁺/Ki67⁺ double-positive cells (indicated by yellow arrows) was significantly increased in the AI group, AS group, and ASI group compared to the Control group, with statistical significance. Among these, the AI group had a higher number of double-positive cells than the AS group, but fewer than the ASI group, with a statistically significant difference between the latter two (p < 0.01, Fig. 7C). These results suggest that the observed effects cannot be solely attributed to IGF-2, as some studies have reported that ASCs can differentiate into Pax3⁺ muscle progenitor cells under specific conditions, contributing to the observed increase in Pax3⁺/Ki67⁺ cells [77,78].

Fig. 7 — **STIM1-ASCs recellularized IGF-2-loaded adECM promotes Pax3⁺ proliferation and M2 macrophage polarization in vivo.** (A) H&E staining of regenerated muscle fibers at the VML site on day 7. Scale bar = 50 μm. Black dashed lines indicate the approximate interface between regenerated and remaining muscle tissue. (B) Infiltration and proliferation of Pax3⁺/Ki67⁺ cells at the VML site in each group (red: Pax3, green: Ki67, blue: DAPI, n = 3). Scale bar = 50 μm. (C) Quantitative analysis of Pax3⁺/Ki67⁺ cell counts per field. (D) Dual immunofluorescence staining images: First row: iNOS (red) and CD68 (green) in different groups 7 days after treatment. Nuclei are shown in blue, and yellow arrows indicate CD68⁺/iNOS⁺ (M1) cells. Second row: CD163 (red) and CD68 (green) in different groups 7 days after treatment. Nuclei are shown in blue, and yellow arrows indicate CD68⁺/CD163⁺ (M2) cells. Scale bar = 50 μm. (E) Statistical analysis of the percentage of CD68⁺/iNOS⁺ and CD68⁺/CD163⁺ cells. Data are presented as mean ± s.d. (n = 3). Statistical significance was determined using one-way ANOVA: ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001 compared with the Control group; ^##p < 0.01 compared with the ASI group.

The phenotype and function of macrophages in the early injury microenvironment play a crucial role in determining the degree of later-stage muscle fibrosis and the formation of mature muscle fibers. During the first 3 days, M1 macrophages dominate, primarily responsible for clearing bacteria and necrotic tissue. Between days 3 and 8, M1 macrophages transition to the M2 phenotype, which suppresses fibroblast proliferation, mitigates fibrosis, and promotes the fusion of immature myofibers into thick, mature muscle fibers [16,34]. therefore, to investigate this transition, we performed immunofluorescence staining for CD68/iNOS and CD68/CD163 in the injury regions of each group on day 7. CD68 marks M0 macrophages, iNOS marks M1 macrophages, and CD163 marks M2 macrophages. In the Control group and adECM group, there were significantly more CD68⁺/iNOS⁺ (M1) macrophages per field of view, with very few CD68⁺/CD163⁺ (M2) macrophages (yellow arrows). Conversely, in the AI group, AS group, and ASI group, the number of CD68⁺/iNOS⁺ (M1) macrophages was substantially reduced, while the number of CD68⁺/CD163⁺ (M2) macrophages was significantly increased (Fig. 7D). Among these, the AI group and ASI group exhibited the highest numbers of M2 macrophages, with no statistically significant difference between them (Fig. 7E).

In addition, to assess whether ASI and AI plays a role in the metabolic reprogramming of macrophages in vivo, we used rotenone (an inhibitor of mitochondrial oxidative phosphorylation [79]) to block the effect of IGF-2 on glucose metabolism. We then observed the level of macrophage polarization to M2 macrophages in muscle tissue at day 7. Considering that other treatment groups did not involve IGF-2 intervention, we set up rotenone intervention groups for both the AI and ASI groups (AI + Rotenone and ASI + Rotenone). The results, shown in Fig. S9, indicate that after rotenone intervention, the M1 macrophage count in the AI + Rotenone group did not significantly increase compared to the AI group (Fig. S9A). However, in the ASI + Rotenone group, M1 macrophages were significantly increased, showing a statistically significant difference compared to the ASI group (P < 0.05, Fig. S9C). More importantly, rotenone intervention significantly suppressed M2 macrophage activation in both the AI and ASI groups, leading to a marked decrease in the proportion of M2 macrophages (Figure S9B and Figure S9D). These results suggest that when rotenone inhibits IGF2's regulation of glucose metabolism, mitochondrial oxidative phosphorylation is impaired, ultimately suppressing the activation of M1 macrophages to M2 macrophages.

In conclusion, the tissue-engineered constructs used in this study appear to enhance muscle regeneration through two mechanisms: first, by activating residual satellite cells in the local microenvironment to replenish the muscle progenitor cell pool; and second, by promoting macrophage polarization toward the M2 phenotype, effectively altering the immune microenvironment during the early stages of injury. These changes create optimal conditions for subsequent muscle regeneration and fibrosis reduction.

3.7. STIM1-ASCs recellularized IGF-2 loaded adECM improves masseter volume and strength

On day 42 of the treatment, we evaluated the chewing rate, feeding frequency, and food intake of the rats to assess the effects of different scaffolds on feeding behavior. Following established methods [50], we used a high-speed camera to capture the chewing motions of the animals. As shown in Fig. 8A, one complete chewing cycle was defined as the movement from full occlusion to full opening and back to full occlusion of mouth. The number of chewing cycles per second was recorded as the chewing rate. Statistical analysis revealed that the chewing rate of the Control group was 5.16 ± 0.08 counts/second. In comparison, the ASI group showed a significant increase in chewing rate (6.15 ± 0.33 counts/second), with a statistically significant difference between the two groups (p < 0.05, Fig. 8B). We also recorded the feeding frequency of the animals over a 12-h period using the camera and weighed the remaining food afterward to determine total food intake. As shown in Fig. 6C, the Control group had a feeding frequency of 105 ± 1.15 counts in 12 h, whereas the adECM group, AI group, AS group, and ASI group showed feeding frequencies of 108 ± 4.58 counts, 114 ± 9.54 counts, 111.67 ± 4.62 counts, and 118.33 ± 5.86 counts, respectively. Among these, only the ASI group exhibited a statistically significant increase compared to the Control group. Similarly, the ASI group showed a significant increase in food intake (25.4 ± 9.54 g) over 12 h compared to the Control group (17.6 ± 2.98 g). The food intake of the ASI group was even close to that of the Native group (27.2 ± 1.84 g).

We also conducted an assessment of the bite force in rats from different treatment groups at the end of the animal experiments. The method used was adapted from Zhao et al.’s experimental protocol [50]. The results, shown in Fig. 8D, indicate that, compared to the Control group, there was no significant improvement in bite force in the adECM group, with no statistically significant difference between the two groups. This finding is inconsistent with previous studies, which suggested that dECM materials can connect the muscle ends on either side of the injury, playing a role in transmitting muscle force [36,80]. In fact, recent studies have indicated that simple dECM scaffolds are much less effective in enhancing limb muscle function than decellularized scaffolds repopulated with cells, as dECM scaffolds alone cannot efficiently regenerate muscle fibers [72]. On the other hand, recellularized dECM scaffolds exhibit higher levels of muscle regeneration [25]. The results of this study align with these findings. Specifically, in the AS and ASI groups, the animals showed a significant increase in bite force, with statistically significant differences compared to the Control group (P < 0.01). Interestingly, the bite force in the AI group also increased significantly and was even comparable to that of the ASI group. This improvement may be attributed to the dual effects of IGF2 within the dECM scaffold, which, on one hand, inhibits fibroblast activity, reducing excessive deposition of fibrous scars, and on the other hand, regulates the activation of M2 macrophages, enhancing the number of myogenic cells. These combined effects contributed to the improvement in bite force in the AI group. Additionally, no significant statistical differences were observed between the AI, AS, and ASI groups (Fig. 8D).

Ultrasound imaging of the masseter muscle can be used to assess muscle volume and fibrosis levels. As shown in Fig. 8C, at day 42 post-treatment, the Control group exhibited strong fibrosis signals in the injured area of the masseter muscle. While fibrosis signals were also observed in the other treatment groups, their intensity was markedly lower than that of the Control group. Notably, the injured region of the masseter muscle in the ASI group displayed not only reduced fibrosis but also imaging signals indicative of normal muscle fibers (Fig. 8E). Additionally, we calculated the cross-sectional area of the masseter muscle based on the ultrasound images. The masseter muscle cross-sectional area in the Control group was 53.90 ± 1.34 mm². Although the adECM group, AI group, and AS group showed slight increases in cross-sectional area, none of these differences were statistically significant compared to the Control group. In contrast, the ASI group exhibited a significant increase in cross-sectional area (64.35 ± 1.76 mm²), with a statistically significant difference compared to the Control group (p < 0.001, Fig. 8F).

Additionally, we measured the muscle strength of the masseter muscle in each group using a muscle force measurement system (DMT820MS, Aarhus, Denmark). The optimal muscle length (1.0 cm), voltage (20 V), and frequency (100 Hz) corresponding to the maximum muscle strength of the Native group served as standard values [25]. As shown in Fig. 8G, during tetanus contraction, the initial peak force of the masseter muscle in the Control group was 74.08 mN, with rapid fatigue or loss of muscle vitality (time required for muscle strength to decline to half the initial peak [81]). The adECM group and AI group also exhibited rapid fatigue or vitality loss, with initial peak forces of 96.44 mN and 86.53 mN, respectively. In contrast, the AS group and ASI group displayed slower fatigue or vitality loss during tetanus contraction, with significantly higher initial peak forces of 158.93 mN and 180.78 mN, respectively. Next, we normalized the maximum muscle strength values by the weight of each muscle tested. The average maximum muscle strength values for each group are shown in Fig. 8H. The Control group and adECM group had the lowest maximum muscle strengths, at 0.70 ± 0.12 mN/mg and 0.65 ± 0.05 mN/mg, respectively. The AI group (0.97 ± 0.07 mN/mg), AS group (1.20 ± 0.09 mN/mg), and ASI group (1.42 ± 0.01 mN/mg) showed significant increases in maximum muscle strength, with statistically significant differences compared to the Control group. Among these, the ASI group exhibited significantly higher maximum muscle strength than both the AI group and AS group (p < 0.05). These imaging and muscle force measurement results indicate that transplantation of the adECM scaffold alone does not prevent muscle atrophy or loss of strength, consistent with previous reports [72]. Although IGF-2-loaded adECM and STIM1-ASCs recellularized adECM scaffolds partially improved muscle atrophy and strength, their effects were inferior to those of the STIM1-ASCs recellularized IGF-2-loaded adECM scaffold.

4. Discussion

Currently, although some progress has been made in the treatment of craniofacial volumetric muscle loss (VML) using muscle tissue engineering constructs, poor muscle regeneration and severe muscle fibrosis remain the primary causes of adverse outcomes in VML [3,49]. The fundamental issues underlying these challenges are the complete absence of satellite cells in muscle tissue and the impaired transition of M1 to M2 macrophages in the immune microenvironment. This is particularly true for craniofacial muscle tissue, which contains fewer satellite cells (SCs) and exhibits slower proliferation rates [9,16]. In this study, based on these pathological and physiological characteristics, we first developed STIM1-overexpressing adipose-derived stem cells (STIM1-ASCs), a cell type with high myogenic differentiation potential, and used them to recellularize an IGF-2-loaded adECM bioscaffold. This approach provided an adequate supply of muscle progenitor cells to support muscle regeneration at both the injury site and within the scaffold. Additionally, IGF-2 was adsorbed into the adECM bioscaffold to enable its localized release at the injury site. This facilitated precise modulation of macrophage phenotypes within the microenvironment, increasing the activation of anti-inflammatory M2 macrophages, suppressing fibrotic scar formation, and promoting the regeneration of mature muscle fibers.

From the perspective of the biological scaffold, numerous studies have demonstrated that skeletal muscle decellularized extracellular matrix (dECM) may contain a biochemical composition that provides a favorable environment for the regeneration of new skeletal muscle and promotes positive remodeling characteristics and myogenesis in vivo [38]. As a result, skeletal muscle dECMs are considered an ideal candidate scaffold for muscle recovery. However, given the limitations of skeletal muscle dECM, the application of adipose-derived extracellular matrix (adECM) is more compelling in terms of both manipulation techniques and scaffold quality. On one hand, skeletal muscle scalability presents a number of technological challenges and difficulties in achieving fully perfused decellularization for transplantability, which is much more complex than techniques previously reported for other partial-organ perfusion decellularizations [39]. In contrast, the structure of adipose tissue is relatively loose, and low-concentration decellularization reagents can fully removeadipocytes, thus minimizing damage to the natural structure and components of the dECM [44,82]. Moreover, adipose tissue's blood supply vessels are relativelysimple, enhancing decellularization efficiency and reducing processing time. Furthermore, adipose dECM has greater potential for clinical application due to its abundant sources, high yields, and easier transformation into clinical practice.

More importantly, the debate over the superiority of skeletal muscle-specific ECM scaffolds for the constructive remodeling of damaged muscle tissue continues. Although no study has directly compared the repair effects of sub-organ scale skeletal muscle dECM and large-volume adipose dECM for VML, several studies provide valuable insights. Fernando et al. compared the treatment of VML with adipose tissue grafts and skeletal muscle dECM. Histological evaluation suggested that the VML defect receiving adipose tissue transplantation was filled with a large amount of newly formed muscle tissue, despite the abnormal structure of muscle fibers [83]. However, skeletal muscle dECM resulted in muscle fibrosis and inflammation, indicating failed muscle tissue regeneration [84]. Similar results have also been reported by other studies [85,86]. Gamba et al. transplanted diaphragmatic muscle dECM into a rabbit abdominal wall defect model. After 90 days of treatment, there was no sign of skeletal muscle tissue regeneration within the biological scaffold, and the diaphragmatic muscle decellularized matrix had been reshaped into fibrous tissue [86].

In this study, we employed a method of perfusing a decellularization solution through the adipose tissue vascular pedicle to prepare large-volume adECM biological scaffolds. This method offers several advantages: 1) The single blood vessel supply allows the decellularization solution to be evenly perfused throughout the scaffold, requiring less solution, reducing processing time, improving decellularization efficiency, and ensuring more consistent batch stability [39]; 2) The retention of the vascular pedicle provides a natural conduit for the subsequent recellularization of the scaffold with STIM1-ASCs, ensuring uniform cell seeding and preventing the formation of non-functional muscle islands [25]; 3) After transplantation, microsurgical techniques can be used to anastomose the scaffold's vascular pedicle with adjacent blood vessels, facilitating the rapid supply of nutrients to the scaffold's interior and supporting the metabolic activities necessary for cell differentiation and proliferation within the scaffold [40].

Nonetheless, studies suggest that the muscle fiber arrangement within large-volume adECM scaffolds tends to be disordered, which differs significantly from the regular, layered arrangement of muscle fibers in normal muscle tissue [87,88]. The ability of muscle to generate strong contraction force is partially dependent on the highly organized alignment and connection of muscle fibers. Therefore, muscle fibers that are not aligned within the adECM scaffolds are not conducive to efficient muscle contraction. To address this issue, researchers have explored various strategies, including the application of external forces such as tensile and magnetic forces to promote the formation of organized muscle tissue with aligned muscle fibers during regeneration [89,90]. However, several challenges remain, such as clarifying key parameters like the frequency and duration of tensile and magnetic forces. More importantly, since muscle contraction force and modes of contraction vary across different muscle subunits, it is crucial to apply personalized approaches when evaluating the effectiveness of external interventions, based on the anatomical and physiological characteristics of different muscle regions [91].

In addition to some of the strategies we mentioned above, such as magnetic field stimulation, recent studies have also indicated that mechanical stimulation has potential to address these challenges [91]. In most studies, muscle precursor cells are seeded onto biological scaffold materials with suitable elastic modulus, and these scaffolds undergo regular dynamic and static stretching and compression. The goal is to guide the regeneration of muscle cells to grow in alignment with the direction of mechanical stimuli, leading to the orderly arrangement of regenerated muscle fibers [[92], [93], [94]]. YOO et al. applied pre-mechanical stimulation to decellularized scaffolds seeded with cells, which effectively facilitated the orderly alignment of muscle fibers after in vivo transplantation [95]. Moreover, active or passive muscle movement after transplantation is beneficial for the orderly remodeling of muscle fibers. Both preclinical and clinical studies have shown that post-surgery exercise accelerates muscle fiber regeneration and promotes the rearrangement of fibers along the direction of force application [36,[96], [97], [98]].

From the perspective of the seed cells, ASCs have emerged as a promising candidate for muscle tissue engineering in recent years. However, their low myogenic differentiation efficiency limits their clinical application [99]. Previous studies have attempted to promote myogenic differentiation by regulating the expression of transcription factors in ASCs, such as myogenic factor 5 (Myf5), myogenic differentiation 1 (MyoD), myogenin (MyoG), and myogenic regulatory factor 4 (MRF4, also known as Myf6) [23,100]. Alternatively, co-culturing ASCs with myogenic cells has been explored to improve myofiber fusion and increase the proportion of mature myofibers [101]. However, these methods have not resulted in high levels of myogenic differentiation in ASCs. In this study, we identified that STIM1 may play a role in the myogenic differentiation of ASCs based on single-cell sequencing data [25]. To further validate this, we used lentiviral transfection to stably overexpress the STIM1 gene in ASCs. After in vitro myogenic induction (under both 2D and 3D culture conditions), STIM1-ASCs exhibited significantly enhanced myotube formation compared to ASCs. Furthermore, we used TEM to assess the maturity of muscle fibers formed by STIM1-ASCs. The results shown in Fig. S7 indicate that not all cells are able to differentiate and fuse into mature muscle fibers or myotubes, because we did not observed the clearly defined and well-aligned sarcomeres or Z-bands, which are strong evidence of muscle fiber maturation [102]. Thus, the majority of muscle fibers formed by STIM1-ASCs are at a mid-to-late stage of maturation.

From the safety perspective of lentiviral transfection technology, the STIM1 gene sequence in this study was integrated into the ASCs' genome, which could cause uncertainty in the direction of cell differentiation and may even result in adverse outcomes such as tumorigenesis. Therefore, to demonstrate the in vivo biosafety of STIM1-ASCs, we performed histological staining on the major organs (heart, liver, spleen, lungs, kidneys) of the treatment groups receiving STIM1-ASCs (AI and ASI). H&E staining results, as shown in Fig. S10, revealed no significant signs of inflammation, necrosis, or tumor formation in the major organs of the AI and ASI groups compared to the Native group. Despite this, adenovirus vectors, which are already used in clinical applications [103], appear more suitable for transfecting STIM1 into ASCs, as they do not integrate foreign gene sequences into the cell's genome. This avoids uncertain differentiation and reduces the risk of tumorigenicity, offering an alternative to STIM1-ASCs.

In this study, IGF-2 was loaded into the adECM bioscaffold through a co-incubation method. IGF-2 release experiments demonstrated that adECM scaffolds loaded with different concentrations of IGF-2 could sustain its release over 7 days, effectively covering the macrophage phenotype transition window. Additionally, the release profile showed a burst effect within the first 3 h, with up to 45 % of IGF-2 released in the 5 ng/mL group. This release pattern is acceptable from the perspective of IGF-2's mechanism of action in muscle repair, as high concentrations of IGF-2 in the early injury phase promote the reactivation and proliferation of quiescent SCs, driving them into a differentiation state [31]. Fig. 7A and B confirm the proliferative effect of IGF-2 on muscle stem cells, as both the AI group and ASI group showed increased numbers of SCs (Pax3⁺/Ki67⁺ cells). Interestingly, compared to the AI group, the ASI group exhibited a significantly higher number of Pax3⁺/Ki67⁺ cells. This effect is likely not solely attributable to IGF-2. According to previous reports, ASCs can differentiate into Pax3⁺ muscle progenitor cells under specific myogenic induction conditions, which may also contribute to this outcome [77,78].

Moreover, interleukin-4 (IL-4) and IL-13 have been shown to promote tissue repair through macrophage polarization, including in skeletal muscle, blood vessels, and other tissues [104,105]. Additionally, studies have demonstrated that insulin-like growth factor (IGF-1) promotes the proliferation, differentiation, and maturation of muscle satellite cells and myogenic precursor cells [106]. As a result, researchers often supplement tissue-engineered materials with IL-4 and IGF-1. IL-4 acts as an early factor to promote macrophage polarization, while IGF-1 serves as an active factor in the early to mid-stage to activate satellite cells or precursor cells and promote myogenic differentiation [33]. According to the literature, when muscle is injured, muscle cells secrete large amounts of IGF-2 into the microenvironment, activating muscle satellite cells and promoting their proliferation to compensate for the deficiency of myogenic precursor cells [107]. Subsequently, IGF-2 no longer regulates the proliferation of satellite cells but instead regulates the polarization of macrophages from M1 to M2 type in the microenvironment [108]. However, the above-mentioned role of IGF-2 has been discovered in independent experiments, and no study has simultaneously observed the spatiotemporal role of IGF-2 in VML repair. This study not only confirmed that the role of IGF-2 in a VML model is temporally and spatially coupled with the myogenesis process but also designed a muscle tissue engineering construct that simultaneously regulates macrophage polarization and STIM1-ASCs myogenic differentiation. Compared to previous studies, our approach uses a single active factor to regulate two key biological processes. From the perspective of tissue engineering material construction, this study reduces the complexity of scaffolds designed to deliver two separate cytokines, thus advancing its clinical translation potential.

IGF-2's role in skeletal muscle growth and development was demonstrated early on [109], but the hypothesis that IGF-2 influences skeletal muscle regeneration by altering macrophage phenotypes and metabolic states has only been proposed in recent years. Fang et al. found that muscle satellite cells promote the activation of M2 macrophages in the immune microenvironment by secreting IGF-2, thereby reducing inflammation in inflammatory bowel disease tissues [30]. Zhang et al. discovered that silencing insulin-like growth factor mRNA-binding protein 3 (IGF2BP3) expression, which suppresses trophoblast cell secretion of IGF-2, leads to a decrease in IL-10 levels in the microenvironment, inhibition of the NF-κB signaling pathway, and ultimately a reduction in the activation ratio of M2 macrophages [110], indirectly reflecting the regulatory effect of IGF-2 on M2 macrophage activation.

Regarding the mechanism by which IGF-2 regulates macrophage metabolic reprogramming and alters their polarization, study in experimental autoimmune encephalomyelitis models have shown that IGF-2-treated macrophages display an increased polarization towards the M2 phenotype [108]. This is accompanied by a significant increase in oxidative phosphorylation (OCR/ECAR ratio), higher mitochondrial membrane potential, and an elevated ATP/ADP ratio, a state that cannot even be reversed by M1 macrophage activator LPS. Further research indicated that the sustained activation of mitochondrial complex V in M2 macrophages, which is significantly correlated with the activation of the Akt signaling pathway, may account for the maintenance of high oxidative phosphorylation levels [108].

Importantly, in this study, we have validated the mechanism by which IGF-2 regulates macrophage metabolic reprogramming. Based on transcriptomic sequencing results, we identified that the adECM scaffold carrying IGF-2 (adECM + IGF2) may alter the mitochondrial energy metabolism in macrophages through the PI3K-Akt signaling pathway. We then intervened with cells using a PI3K-Akt signaling pathway inhibitor (LY294002). The results showed that under LY294002 intervention, the levels of p-PI3K and p-AKT proteins in macrophages from the adECM + IGF2+LY294002 group were significantly reduced, accompanied by a significant decrease in the mitochondrial OXPHOS levels, as indicated by a lower OCR/ECAR ratio. Simultaneously, the mRNA expression levels of key enzymes involved in mitochondrial oxidative phosphorylation, ATP5A1, Ndufb5, and COX4I1 were decreased. These results suggest that IGF-2 regulates macrophage metabolic reprogramming through the PI3K-Akt signaling pathway.

Furthermore, combining the macrophage immufluorescence staining results from day 7 with previous findings, the IGF-2-loaded bioscaffold groups (AI group and ASI group) indeed induced an increase in M2 macrophages and a decrease in M1 macrophages, consistent with the results of flow cytometry and Seahorse analysis. However, the adECM group did not demonstrate the ability to alter macrophage phenotypes. According to previous studies [74,111], dECM materials can promote macrophage polarization toward the M2 phenotype, which contradicts our findings. In this study, the low activation ratio of M2 macrophages in the adECM group may have contributed to the more severe muscle fibrosis and lower muscle strength observed later. We hypothesize that the differences in results may be related to the following two factors, 1) The form in which adECM is applied: adECM is often used in the form of hydrogels, bioinks, or electrospun nanofibers [74,111,112]. In contrast, in this study, adECM was applied as a large-volume biological scaffold. 2) The decellularization method of adECM: Literature reports indicate that the use of different decellularization agents (such as SDS or SDC) and varying concentrations of these agents during the decellularization process can affect the material's microstructure and the retention of bioactive components, which in turn alters the adECM's ability to influence macrophage polarization [113,114].

Additionally, the selection of the optimal IGF-2 concentration (10 ng/mL) seems to be controversial. Based on the experimental results showing that IGF-2 promotes the polarization of macrophages to the M2 phenotype, we found that 20 ng/mL of IGF-2 also promotes M2 macrophage activation and alters mitochondrial metabolic reprogramming, although the enhancement of mitochondrial oxidative phosphorylation is weaker than in the 10 ng/mL group. Furthermore, at a concentration of 20 ng/mL, IGF-2 also effectively inhibits fibroblast proliferation, migration, and collagen deposition, with no significant statistical difference between the two concentrations. In the animal experiments, we used adECM scaffolds loaded with IGF-2 (10 ng/mL) because 10 ng/mL of IGF-2 significantly promoted myogenic differentiation of STIM1-ASCs, which is crucial for effective muscle tissue regeneration within the scaffold.

After reviewing the literature [115], we initially hypothesized that the higher concentration of IGF-2 might affect the cell viability of STIM1-ASCs, thereby impairing their myogenic differentiation potential. To investigate this, we treated STIM1-ASCs with adECM + IGF-2 (0 ng/mL), adECM + IGF-2 (5 ng/mL), adECM + IGF-2 (10 ng/mL), and adECM + IGF-2 (20 ng/mL) for 48 h, followed by a live/dead cell staining assay. The results, shown in Fig. S11, revealed that compared to the control group (adECM + IGF-2 (0 ng/mL)), the adECM + IGF-2 (20 ng/mL) group had relatively fewer viable cells, with a noticeable increase in dead cells, whereas the adECM + IGF-2 (5 ng/mL) and adECM + IGF-2 (10 ng/mL) groups showed no significant difference in dead cell numbers compared to the control group. These results suggest that the high concentration of IGF-2 (20 ng/mL) may negatively impact the viability of STIM1-ASCs, thus leading to a lower efficiency in promoting myogenic differentiation in this group compared to the adECM + IGF-2 (10 ng/mL) group. As for why adECM + IGF-2 (20 ng/mL) did not reduce the viability of macrophages and fibroblasts, we speculate that after lentiviral infection, the sensitivity of STIM1-ASCs to IGF-2 may be reduced [116].

Although the biological scaffold constructed in this project shows promising antifibrotic and muscle regeneration effects, there are still several issues to address in the clinical translation process. The first concern is the long-term stability of the scaffold in vivo. While there are currently no clinical reports on the use of adipose decellularized biological scaffolds for treating volumetric muscle loss (VML), data from other tissue-derived decellularized biological scaffolds, such as small intestine submucosa, dermis, and bladder decellularized scaffolds, seem to provide valuable references regarding in vivo stability and degradation rates. A clinical cohort study involving 13 VML patients found that, 8 months after transplantation with different types of decellularized scaffolds, imaging techniques (CT, MRI, and ultrasound) and histological staining confirmed that some remnants of the decellularized scaffolds remained, even though some scaffolds had remodeled into tissue consistent with muscle imaging signals [36]. Although a direct comparison of degradation rates or in vivo stability between various decellularized tissue scaffolds is not available, the evidence suggests that the longer remodeling cycle of adipose decellularized scaffolds should be sufficient to meet the time required for muscle tissue repair.

Moreover, although the adipose decellularized scaffold retains the vascular pedicle, even after vascular pedicle anastomosis surgery, the endothelial cells of the internal vessels are completely removed, which leads to blood leakage and fails to meet the metabolic needs of the cells within the scaffold [8]. Therefore, the implanted scaffold primarily relies on the infiltration growth of host blood vessels, but this process is relatively slow. Scholars have proposed pre-vascularization strategies by perfusing endothelial cells and supporting cells (such as fibroblasts) through the vascular pedicle into decellularized scaffolds, and some progress has been made in this area [117]. This direction will guide our next phase of research, which involves pre-vascularizing and pre-myogenesis the scaffold in vitro, with the goal of further enhancing its therapeutic efficacy for VML in vivo, thereby increasing its clinical translation potential.

In addition, the preparation process of large-volume adipose decellularized scaffolds is more complex and requires more time and cost. For example, compared to the large adipose flaps obtained from abdominal wall plastic surgery, which retain their pedicle, the fragmented adipose tissue obtained through liposuction has advantages in terms of decellularization effectiveness, time, and cost. At the same time, mass production of pedicled adipose decellularized scaffolds is also challenging. Considering that there is currently no standardized decellularization protocol, batch-to-batch variations in decellularized scaffolds are inevitable. Therefore, further research and exploration are needed to address these issues in the future.

5. Conclusion

In this study, we designed an adipose-derived extracellular matrix (adECM) bioscaffold recellularized with STIM1-overexpressing adipose-derived stem cells (STIM1-ASCs) and loaded with the cytokine IGF-2, tailored to the pathophysiological characteristics of craniofacial muscle (masseter) VML repair. On one hand, we leveraged the high myogenic differentiation potential of STIM1-ASCs to address the shortage of progenitor cells in VML-injured muscle tissue. On the other hand, we utilized the dual role of IGF-2 in muscle repair: promoting the activation and proliferation of residual progenitor cells at the injury site and precisely regulating the polarization of M2 macrophages within the local microenvironment. The later fundamentally alleviated the severe fibrosis typically associated with VML injuries. More importantly, IGF-2 was shown to alter macrophage glucose metabolism, inducing metabolic reprogramming by enhancing mitochondrial respiration, ultimately driving the phenotype transition from M1 to M2 macrophages.

CRediT authorship contribution statement

Wei Liang: Writing – review & editing, Writing – original draft, Methodology, Investigation, Funding acquisition, Formal analysis. Rigele Ao: Writing – review & editing, Writing – original draft, Investigation, Data curation. Mengli Xu: Methodology, Investigation. Mengying Jin: Visualization, Methodology, Investigation. Meng Han: Writing – original draft, Visualization, Methodology. Zimo Wang: Visualization, Investigation. Wanwen Dang: Methodology, Investigation. Hongxu Wu: Methodology. Weibo Lin: Methodology. Yonghuan Zhen: Supervision. Tao Xu: Conceptualization. Yang An: Writing – review & editing, Funding acquisition, Conceptualization.

Ethics approval and consent to participate

The animal protocols were approved by the Peking University Biomedical Ethics Committee (No. A2024081). All in vivo experiments were conducted in accordance with the National Institutes of Health guidelines for the care and use of laboratory animals.

Declaration of competing interest

The authors have no financial interest to declare in relation to the content of this article.

Acknowledgement

National Natural Science Foundation of China (NO. 82503035); Beijing Natural Science Foundation (NO. 7254442); China Postdoctoral Science Foundation (NO. 2024M750129); Postdoctoral Fellowship Program of CPSF (NO. GZC20230151); Peking University Medicine Sailing Program for Young Scholars' Scientific & Technological Innovation (NO. BMU2024YFJHPY030). Peking University Third Hospital Fund for Interdisciplinary Research (NO. BMU2025XY032).

Footnotes

Peer review under the responsibility of editorial board of Bioactive Materials.

^{Appendix A}

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bioactmat.2025.08.019.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Multimedia component 1

mmc1.docx^{(2MB, docx)}

Data availability

Data will be made available on request.

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PERMALINK

Bifunctional adECM bioscaffold with STIM1-ASCs and IGF-2 promotes functional masseter VML repair via myogenesis and fibrosis suppression

Wei Liang

Rigele Ao

Mengli Xu

Mengying Jin

Meng Han

Zimo Wang

Wanwen Dang

Hongxu Wu

Weibo Lin

Yonghuan Zhen

Tao Xu

Yang An

Abstract

Graphical abstract

Highlights

1. Introduction

2. Materials and methods

2.1. Preparation of the adECM with vascular pedicle

2.2. Characterization of adECM

2.2.1. cDNA content

2.2.2. Glycosaminoglycan (GAG) content

2.2.3. Swelling rate and degradation rate

2.2.4. Histological staining

2.2.5. Scanning electron microscopy (SEM)

2.2.6. Mechanical properties assessment

2.2.7. SDS-polyacrylamide gel electrophoresis (SDS-PAGE)

2.3. IGF-2 loaded adECM regulates macrophage polarization

2.3.1. The efficiency of adECM loading IGF-2

2.3.2. IGF-2 release analysis of adECM

2.3.3. Bone marrow-derived macrophages (BMDMs) cell isolation and culture

2.3.4. IGF-2-loaded adECM scaffolds regulates macrophage polarization in vitro

2.3.5. Quantitative real-time polymerase chain reaction (RT-qPCR)

2.3.6. Seahorse analysis

2.3.7. Western blotting

2.3.8. RNA sequencing (RNA-seq)

2.4. IGF-2-loaded adECM scaffolds alter the function of human normal skin fibroblasts by promoting M2 macrophage polarization

2.5. Myogenic differentiation of STIM1-ASCs inside IGF-2 loaded adECM

2.5.1. Preparation of STIM1-ASCs and myogenic differentiation

2.5.2. SCs isolation and culture

2.5.3. STIM1-ASCs and SCs myogenic differentiation

2.5.4. Cytotoxicity test of adECM

2.5.5. Distribution and proliferation of ASCs inside adECM

2.5.6. Myogenic differentiation of STIM1-ASCs inside IGF-2 loaded adECM

2.6. Masseter VML injury reconstruction with STIM1-ASCs recellularized IGF-2-loaded adECM

2.6.1. Masseter VML injury model

2.6.2. Histologic evaluation of masseter

2.6.3. Imaging evaluation of masseter

2.6.4. Mastication rate measurement and eating behaviour analyses

2.6.5. Contraction strength evaluation of masseter muscle

2.6.6. Bite force measurement

2.7. Statistical analysis

3. Results

3.1. Preparation and characterization of adECM with vascular pedicle

Fig. 1.

3.2. The IGF-2 loaded adECM induce the polarization of macrophage into M2 macrophage phenotype

Fig. 2.

Table 1.

Fig. 3.

3.3. The IGF-2 loaded adECM regulates fibroblast function by promoting M2 macrophage activation

Fig. 4.

3.4. Myogenic differentiation of STIM1-ASCs when in IGF-2 loaded adECM

Fig. 5.

3.5. STIM1-ASCs recellularized IGF-2 loaded adECM treatment for masseter VML

Fig. 6.

3.6. STIM1-ASCs recellularized IGF-2 loaded adECM promotes Pax3+ proliferation and M2 polarization in vivo

Fig. 7.

3.7. STIM1-ASCs recellularized IGF-2 loaded adECM improves masseter volume and strength

Fig. 8.

4. Discussion

5. Conclusion

CRediT authorship contribution statement

Ethics approval and consent to participate

Declaration of competing interest

Acknowledgement

Footnotes

Appendix A. Supplementary data

Data availability

References

3.6. STIM1-ASCs recellularized IGF-2 loaded adECM promotes Pax3⁺ proliferation and M2 polarization in vivo