Urine-derived stem cells genetically modified with IGF1 improve muscle regeneration

Abstract

Objective: In this study we aimed to determine the impact of human urine derived stem cells (USC) and genetically modified USC that were designed to overexpress myogenic growth factor IGF1 (USC^IGF), on the regenerative capacity of cardiotoxin (CTX)-injured murine skeletal muscle. Methods: We overexpressed IGF1 in USC and investigated the alterations in myogenic capacity and regenerative function in cardiotoxin-injured muscle tissues. Results: Compared with USC alone, USC^IGF1 activated the IGF1-Akt-mTOR signaling pathway, significantly improved myogenic differentiation capacity in vitro, and enhanced the secretion of myogenic growth factors and cytokines. In addition, IGF1 overexpression increased the ability of USC to fuse with skeletal myocytes to form myotubes, regulated the pro-regenerative immune response and inflammatory cytokines, and increased myogenesis in an in vivo model of skeletal muscle injury. Conclusion: Overall, USC genetically modified to overexpress IGF1 significantly enhanced skeletal muscle regeneration by regulating myogenic differentiation, paracrine effects, and cell fusion, as well as by modulating immune responses in injured skeletal muscles in vivo. This study provides a novel perspective for evaluating the myogenic function of USC as a nonmyogenic cell source in skeletal myogenesis. The combination of USC and IGF1 expression has the potential to provide a novel efficient therapy for skeletal muscle injury and associated muscular defects in patients with urinary incontinence.

Introduction

Skeletal muscles are the largest organs in the human body and account for up to 45% of the total body weight [1]. These organs are affected by frequent mild to moderate injury due to overuse, trauma, aging, and degenerative disorders. Impairment in muscle quantity and quality strongly affects physical activity and health, resulting in diminished strength and mobility and a decrease in quality of life [2]. For most mild to moderate injuries, skeletal muscle has an intrinsic regenerative capacity that requires the differentiation of local skeletal muscle progenitor cells (SMPC), and, after severe injury, reinnervation and revascularization of the muscle tissue. These processes are orchestrated by autocrine and paracrine secretion of factors involved in chemotaxis, differentiation, and growth [3]. SMPC or satellite cells, that reside under the basal lamina of mature muscle tissue play a central role in muscle regeneration, where myogenic differentiation leads to myocyte fusion and the formation of multinucleated myotubes, which further mature into contractile muscle tissue; however, the numbers of satellite cells are often insufficient, or missing, in severe muscle injuries and volumetric muscle deficits [4].

Limited regeneration and restoration of muscle function after injury often result from deficits in SMPC proliferation and differentiation, changes in cellular metabolism, cell survival, or apoptosis [5]. Therefore, the complexity of these pathological processes presents challenges for the treatment of skeletal muscle dysfunction. In recent years, the use of stem cells as a therapy for skeletal muscle injury [6-8] and congenital diseases such as muscular dystrophy [9-12] has been extensively explored. SMPC can initiate myogenesis via direct involvement in muscle fiber formation and maturation, and via indirect mechanisms such as stimulation of paracrine factors, secretion of exosomes, and regulation of the microenvironment at the site of damage [11]. In addition to SMPC, the myogenic potential of several other types of stem cells, including induced pluripotent stem cells (iPSC), bone marrow- and adipose-derived mesenchymal stem cells, and placenta- and umbilical cord-derived stem cells, has been studied [13,14]. However, the mechanisms of myogenesis involving non-muscle-derived stem cells in muscle regeneration have not been fully characterized.

Urine-derived stem cells (USC) are multipotent cells that possess myogenic differentiation capability and robust regenerative potential, making them a viable alternative in clinical therapy for skeletal muscle injury and disease [15,16]. A major advantage of using USC as a therapy for skeletal muscle injury is that USC are noninvasively derived and easily accessible, whereas invasive tissue biopsies are required to obtain SMPCs. The USC can potentially differentiate into muscle precursor cells in vitro, as we previously described [17]; however, whether the USC can contribute to skeletal muscle regeneration and maturation processes in vivo is unknown.

The insulin-like growth factor (IGF) family functions as an endocrine hormone and autocrine/paracrine growth factor family to regulate the development of tissues and organs, particularly in the anabolic and catabolic processes of skeletal muscle metabolism and growth [18]. It is well documented that IGF1 signaling is activated to promote myogenic regeneration following injury [19] and that IGF1 regulates muscle-specific protein synthesis and metabolism, especially carbohydrate and lipid metabolism, in vitro and in vivo to support the metabolic needs of muscle [19,20]. The metabolic effects of IGF1 occur via the activation of the Akt-mTOR signaling pathway, which modulates cell survival and differentiation and stimulates tissue growth and muscle size [21]. IGF1 is also associated with the AMPK-mTOR signaling pathway, which modulates the activation of reactive oxygen species (ROS) and metabolism during myogenesis. We previously showed that USC secrete a series of cytokines and chemokines, including IGF1, VEGF, IL6, IL8, and CXCL14 [22]. Thus, we hypothesized that USC-overexpressing IGF1 (USC^IGF1) would enhance myogenesis and skeletal muscle regeneration in a rodent model of injury.

In this study, we created a stable cell system using USC overexpressing IGF1 (USC^IGF1) via lentivirus transduction. We demonstrated that USC^IGF1 efficiently differentiated into a skeletal muscle lineage and promoted skeletal muscle cell maturation via heterologous organelle-associated cell fusion with mitochondria and exerted paracrine effects via the IGF1-Akt-mTOR pathway, which positively regulates myogenesis. We further demonstrated the myogenic capacity of USC^IGF1 to promote skeletal muscle regeneration by modulating immune responses in an animal model, which could provide new perspectives for the use of a combination of nonmuscle-originating stem cells and gene therapy for muscle repair in urinary system diseases, such as urinary incontinence.

Materials and methods

Cell isolation and culture

Human urine-derived stem cells (USC): The acquisition of human tissue samples was approved by the Wake Forest University Institutional Review Board (IRB00014033), and donors were all clearly aware of how we use these samples in this study. The USC were isolated and expanded as previously described [15,23]. Briefly, Urine samples (ranging from 50 ml to 200 ml) were obtained from healthy male donors (n=9, 23-36 years of age), centrifuged at 900× g for 5 min, collected and resuspended in USC medium and subsequently seeded on 24-well plates. The cells were cultured in an incubator at 37°C until colonies formed, and the clones were passaged into 10 cm tissue culture dishes for further expansion in USC expansion medium, which is composed of keratinocyte serum-free medium (KSFM) supplemented with 5% fetal bovine serum and embryonic fibroblast medium (EFM), mixed well at a ratio of 1:1 [24]. All cells were used at passages 1-4.

Human skeletal muscle progenitor cells (hSMPC): hSMPC were derived from surgical waste materials obtained from discarded donor normal gracilis muscle during plastic surgery. A total of 10⁵ cells were isolated and seeded on 10 cm dishes. The cells were maintained in growth medium supplemented with the appropriate supplements (PeproTech, Rocky Hill, NJ), and the medium was changed every 2 days for proliferation and passaging. Prior to use, hSMPC were labeled using a GFP-containing plasmid (hSMPC^GFP) (pReceiver-LV201-GFP).

Mouse skeletal muscle progenitor cells (mSMPC): GFP-expressing skeletal muscle cells (mSMPC^GFP) were isolated from 8-week-old GFP-expressing transgenic mice (C57BL/b-actin-EGFP, Jackson Laboratories [25]) and cultured as previously described [26]. To isolate skeletal muscle, connective tissues and fat were removed to dissolve the muscle tissue, which was minced into 1 mm³ pieces and digested in 0.2% collagenase type I (Worthington Biochemical Corporation, Lakewood, NJ) for 1 hour at 37°C. The digested muscle tissue was washed with phosphorus buffer solution (PBS), centrifuged, and plated on 1% Matrigel-coated plates. The isolated cells were cultured in muscle cell growth media supplemented with low glucose Dulbecco’s Modified Eagles’ Medium (DMEM; HyClone, Thermo Scientific, Logan, UT), 1% penicillin/streptomycin (Thermo Fisher Scientific, Waltham, MA), 1% chick embryo extract (Sera Laboratories International Ltd., United Kingdom), 20% fetal bovine serum (FBS, GE Healthcare, Logan, UT), and 10% horse serum at 37°C, and 5% CO₂.

IGF1 overexpression and knockdown

The human IGF-1 gene was cloned and inserted into the lentiviral shuttle plasmid Ex-A0153-LV216-mCherry and subsequently transduced into USC. The plasmid pReceiver-LV105-IGF1 was used for IGF-1 overexpression, and the empty pReceiver-LV105 vector plasmid was used as the mock control. IGF-1 was knocked down in USC (USC^shIGF1) using short hairpin RNA (psi-shIGF1-LVRH1MP), and the empty psi-LVRH1MP vector was used as control. All of the plasmids were purchased from GeneCopoeia (GeneCopoeia, Inc., Rockville, MD, USA). 293T cells were prepared as lentivirus packaging cells. The expression plasmids were mixed with the lentivirus packaging plasmids psPAX2 and pMD2G at a ratio of 3:2:1 and added to 500 μl of DMEM without antibiotics or serum. The FuGENE 6 transfection reagent (Promega, WI, USA) was used to mix the mixture with 3 times the total volume of the plasmids. The plasmids were mixed well and added to 293T cells. The viral medium was collected after 36-48 hours of culture, and USC (passage 1) were transfected at 10⁵ cells per well in 6-well plates. The USC medium was changed with puromycin at 1 μg/ml to screen for the positively transfected USC cells, which were identified by mCherry fluorescence.

Myogenic differentiation

USC cells were seeded on 100 mm dishes or on 4-well chamber slides coated with 1% hyaluronic acid gel with heparin (HA-HP, HyStem-HP, ESI-BIO, Alameda, CA) for myogenic differentiation; 10⁵ cells per dish and 3,000 cells per well were seeded. The cells were cultured in myogenic differentiation medium containing DMEM supplemented with 1% antibiotic/antimycotic solution (AA, HyClone, Logan, UT), 2% horse serum (GE Healthcare), 5% FBS, 1% insulin-transferrin-selenium solution (ITS, Lonza, Allendale, NJ), and 250 nM dexamethasone (Lonza).

To induce myogenic fusion in cocultured cells, USC^IGF1-mCherry was co-cultured with either hSMPC^GFP or mSMPC^GFP. The cells (USC^IGF1-mCherry-hSMPC^GFP) were mixed at specific ratios of 1:2, 1:1, 2:1, and 4:1 respectively, and then seeded on a 1% heparin-crosslinked hyaluronic acid gel in myogenic differentiation medium as described above. MitoTracker Red CMXRos (Thermo Fisher Scientific, Waltham, MA) was used to track mitochondria in USC^IGF1 cells following the manufacturer’s instructions. The percentage of nuclei in the syncytia with more than 2 nuclei was calculated to evaluate myotube fusion capacity. At least four visual fields per chamber slide well were evaluated for the percentage of nuclei in the syncytia as a fusion index.

Cell viability assays

USC, USC^IGF1, and USC^shIGF1 knockdown cells were seeded in 96-well plates at 3,000 cells per well in USC culture medium for 11 days. Cell viability was determined using an MTS assay kit (CellTiter 96 One Solution Reagent Promega, Madison, WI), and the absorbance was measured at a wavelength of 490 nm with a Molecular Devices SpectrumMax M5 system (Molecular Devices, LLC., San Jose, CA, USA).

Immunofluorescence and microscopy

After 2 weeks of myogenic differentiation, the cells were collected and fixed in 10% neutral buffered formalin solution for immunofluorescent labeling as previously described [27]. Primary antibodies and dilutions used included Myogenin (Abcam, ab1835), MyoD (Santa Cruz, sc32758), Desmin (Abcam, ab32362), Myosin (Abcam, ab11083), Lamin (Abcam, ab108595), Mitochondria (Abcam, ab3298), Dystrophin (Abcam, ab15277), CD206 (Proteintech, Chicago, IL, 60143-1-Ig), and CD86 (Abcam, ab119857). The secondary antibodies used included goat anti-mouse Alexa Fluor 488 (Invitrogen, 35502) and goat anti-rabbit Alexa Fluor 488 (Invitrogen, A11008) for MyoD, Myogenin, Desmin, and Myosin primary antibodies and goat anti-rabbit Alexa Fluor 647 (Thermo Fisher Scientific, A21245, A21235) and goat anti-mouse Alexa Fluor 647 (Thermo Fisher Scientific, A21235) for Lamin, Mitochondria, and Dystrophin primary antibodies. Mounting medium supplemented with DAPI (Vector Laboratories, Burlingame, CA) was used to label the nuclei. Fluorescence images were observed and recorded using a Leica DM4000B microscope system.

Western blot analyses

After 2 weeks of myogenic differentiation, the cultured cells and mouse skeletal muscle tissues were collected for protein isolation and western blot analysis. Total protein was collected using RIPA buffer containing a 1% protease/phosphatase inhibitor cocktail (Pierce, Rockford, IL). Total protein (10 μg) was separated using 15% precast gels (Bio-Rad Laboratories, Hercules, CA) for IGF1 detection, and 10% precast gels for the detection of the other proteins. The gels were run at 100 V for 1 hour, after which the proteins were transferred to a nitrocellulose membrane at 12 V for 1 hour. Membranes were blocked with 5% nonfat milk in PBST (0.1% Tween 20 in PBS) prior to incubation with the following primary antibodies at 4°C overnight: IGF1 (Abcam, ab9572), p70 S6 Kinase (Cell Signaling Technology, 2708), Phospho-p70 S6 Kinase (Cell Signaling Technology, 9208), S6 (Cell Signaling Technology, 2317), Phospho-S6 (Cell Signaling Technology, 4858), 4EBP1 (Cell Signaling Technology, 9644), Phospho-4EBP1 (Cell Signaling Technology, 9455), mTOR (Cell Signaling Technology, 2972), Phospho-mTOR (Cell Signaling Technology, 2971), GSK-3β (P-GSK3β, Cell Signaling Technology, 12456), Phospho-GSK 3β (P-GSK3β, Cell Signaling Technology, 5558), Akt (Cell Signaling Technology, 4691), Phospho-Akt (P-Akt, Thr308, Cell Signaling Technology, 13038), Phospho-Akt (P-Akt, Ser473, Cell Signaling Technology, 4060), FoxO1α (Abcam, ab39670), Phospho-FoxO1/FoxO3α (Cell Signaling Technology, 9464), Acetyl-CoA Carboxylase (ACC, Cell Signaling Technology, 3676), Phospho-Acetyl-CoA Carboxylase (P-ACC, Cell Signaling Technology, 11818), Raptor (Cell Signaling Technology, 2280), Phospho-Raptor (Cell Signaling Technology, 2083), AMPK (Cell Signaling Technology, 5831), Phospho-AMPK (Cell Signaling Technology, 2535), Phospho-ULK (Cell Signaling Technology, 37762), LC3I/II (Cell Signaling Technology, 4108), and β-actin (Abcam, ab8227). HRP-conjugated anti-rabbit secondary antibody (Abcam, ab288151) and HRP-conjugated anti-mouse secondary antibody (Abcam, ab205719) were used for incubation for 1 hour at room temperature. The immunoblots were developed using Supersignal West Femto Maximum Sensitive Substrate (Thermo Fisher Scientific) within 1 min of incubation at room temperature.

For the quantification of inflammatory cytokines, tissues from injured tibialis anterior muscles, with or without cell therapy, were minced and homogenized using a tissue disruptor with RIPA buffer. Total protein was collected, separated by western blot analysis as previously described, and probed with primary antibodies against IL-1β (Abcam, ab200478), IL-6 (Abcam, ab259341), and TNFα (Abcam, ab92324), with β-actin as control. The gray values of the protein blots were measured by ImageJ software (National Institutes of Health) to represent the relative protein expression level compared to that of β-actin.

Proteome profile arrays

USC^IGF1 and USC (control) cells were seeded in 6-well plates as 2×10⁵ cells per well. After cell attachment, the culture medium was replaced with 1 ml of high-glucose DMEM without FBS or other growth factors. After 24 hours of incubation, the supernatant was collected and filtered through a 0.45 μm filter to collect the medium, after which the cell fragments were removed. Proteome profiling was performed using a Proteome Profiler Human Angiogenesis Array Kit (R&D, Minneapolis, MN) following the provided protocol. The captured proteins were visualized using chemiluminescent detection reagents and recorded by an FUJI imaging system. The data were measured and quantified using ImageJ software (NIH, Bethesda, MA, USA).

qRT-PCR analyses

RNA was extracted from USC^IGF1 and USC (control) cells as described above, and then reverse transcribed into cDNA using the ReverTra Ace-α Reverse Transcriptase Kit (Toyobo, Japan) following the manufacturer’s instructions. cDNA (10 μl) was used for each reaction using a Fluorescent Real-Time Quantitative RT-PCR Kit (Roche). ANGPTL4, CKLF, UBE2C, PRC1, DCBLD2, IGF1, ANLN, CCNB2, PTTG3P, CEP55, COL1A1, CDC20, TOP2A, TYMS, WNT5A, KCNIP4, SYNE2, GABARAPL1, MAOA, SERPINA1, TMEM37, SERPINA3, BBOX1, CXCL14, SLPI, MMP7, APOE, DUSP23, IFITM1, HOPX, and the housekeeping gene GAPDH were used as controls (the primer sequences are provided in Supplementary Table 1). The amplification program consisted of denaturation at 94°C for 30 s, annealing at 60°C for 30 s, and extension at 72°C for 1 min for 45 cycles. The Ct value of each sample was obtained and calculated to determine the relative gene expression level as 2^(-ΔCt).

Microarray analysis

RNA was isolated from cultured USC (control) and USC^IGF1 cells using TRIzol reagent (Invitrogen, Carlsbad, CA). RNA samples were subsequently sent to Wake Forest Baptist Hospital (NC, USA) for microarray hybridization. The RNA samples from the different groups were hybridized to an Illumina BeadChip Array for Gene Expression (HumanHT-12_v4_BeadChip Gene Expression Array, Part Number 15011997). There were three replicates of both experimental groups (Supplementary Table 2). Genes with a fold change (FC) ≥1.5 were filtered by a t-test (P<0.05), and enrichment analysis including Gene Ontology (GO), biological process (BP), cell component (CC), and molecular function (MF) enrichment, was performed using Metascape [28].

Mitochondrial stress and respiration tests

To determine the effect of IGF1 overexpression on the mitochondrial function of USC, the oxygen consumption rate (OCR) of complexes I-V was measured in USC^IGF1 and USC (control) cells using a Seahorse XF96 Extracellular Flux Analyzer (Seahorse Bioscience, North Billerica, MA, USA). The assay medium was supplemented with 25 mM glucose, 2 mM L-glutamine, and 1 mM sodium pyruvate according to the manufacturer’s instructions. Inhibitors of mitochondrial function (oligomycin, carbonyl cyanide-p-trifluoromethoxyphenyl-hydrazone (FCCP)), and a combination of rotenone/antimycin A were added following the manufacturer’s instructions. Basal respiration was measured after subtracting nonmitochondrial respiration. Cell respiration-associated ATP production and proton leakage were calculated by the OCR complex I-V in the presence of oligomycin. The maximal respiration was measured by the OCR following the addition of FCCP. The spare respiration capacity was calculated as the difference between the maximal and basal respiration.

Skeletal muscle injury model

All animals were treated in accordance with ethics approval for the animal study from the Institutional Animal Care and Use Committee (IACUC) of Guangzhou Huateng Biomedical Technology Co., Ltd. In total, 39 female BALB/c-nu/nu mice (aged 6 weeks, approximately 25 g in weight) from the Guangdong Pharmaceutical Undergraduate Laboratory Animal Center were used for USC^IGF1-mCherry transplantation experiments. All mice were housed in standard light, temperature, humidity, and food conditions. Mice were divided into three groups to be treated with USC^IGF1-mCherry, USC^mCherry, or PBS (3 mice/group). To create the skeletal muscle injury model, the left and right tibialis anterior (TA) muscles were injected with 20 μl (0.2 mg/ml) cardiotoxin (CTX). After 24 hours, the injured left and right TA muscles were injected with USC^IGF1 (2×10⁵ cells), USC^mCherry (control) or PBS for each group [29,30]. All experiments were performed under isoflurane anaesthesia for the mice, at the end of the experiment, CO₂ euthanasia was performed for the mice in the cage (3 L/min), continued exposure to CO₂ for at least 20 minutes after cessation of respiration and confirmation of death followed IACUC guidelines. TA muscles were collected from the left and right legs of each mouse at the end of the experiment, fixed in 4% PFA, embedded in cryosections and subjected to immunofluorescence staining, or fixed in 10% neutral buffered formalin solution for paraffin sectioning.

Electrophysiological assessment

Muscle contractility of the TA muscles from the different groups was assessed via quantitative electromyography at each time point (1, 3, 5, and 7 days after cell injection). The nerve trunks were sequentially exposed at the proximal and distal ends to detect and recorde muscle contractility, and compound muscle action potentials (CMAPs) on the ipsilateral side were recorded under electrical stimuli (0.5 V in strength, 0.01 min in pulse width) using a biological function experimental system (BL-420F, TaiMeng, Chengdu, China).

Histochemical evaluation

Paraffin sections of TA muscles were prepared for hematoxylin-eosin (H&E) staining. Sections were dewaxed at 56°C, rinsed in xylene, and rehydrated through 70%, 80%, and 90% alcohol prior to staining with an H&E solution kit (Solarbio, Beijing, China). TA muscle cryosections were used for the detection of M1 and M2 macrophage associated markers using the primary antibodies against CD86 (Abcam, ab119857) and CD206 (ProteinTech, Chicago, IL, 60143-1-Ig), respectively. The area of inflammation, myotube area, and number of immune cells and total cells were measured by ImageJ software. Inflammation and myotube area were determined as the immune cell infiltration area and muscle area with syncytia and the myotube phenotype within 1 mm² of each cryosection. Immune cells and total cells were counted as hematoxylin-stained nuclei with an immune cell phenotype and as total stained nuclei within 1 mm². Four randomly selected cryosections from muscle samples from each group were used for the calculations.

Statistical analysis

All analyses were based on at least 3 independent experiments. The data for all groups and comparisons were analyzed using a one-way analysis of variance (ANOVA) and Tukey’s test as post-hoc analyses. The results are presented as the mean ± standard error of the mean, and p values of *P<0.05 or **P<0.01 were calculated for significant differences. Calculations and statistical analyses were performed using GraphPad Prism Software 6.0 (GraphPad Software Inc., La Jolla, CA, USA).

Results

IGF1 overexpression and knockdown

Human USC were transduced with lentivirus encoding human IGF1 (USC^IGF1). Rapid cell proliferation was observed for USC^IGF1, which resulted in multilayer cells and dome-like structures by Day 7 of culture (Figure 1A). MTS assays were used to demonstrate increased cell survival in USC^IGF1 cells compared to USC and USC^Mock cells over time in culture (Figure 1B). In contrast, USC with IGF1 knockdown (USC^shIGF) exhibited little proliferation, and most of the cells died after 7 days of culture (Figure 1B). The overexpression of IGF1 in cultured cells was assessed via western blot analysis in USC^Mock, USC, and USC^IGF1 cells, increased expression of IGF1 in USC^IGF1 was confirmed by western blot analysis (Figure 1C).

Figure 1.

Changes in the morphology of USC^IGF1 in vitro. A. Proliferation of normal USC, USC with mock vehicle (USC^mock), USC with IGF1 overexpression (USC^IGF1), and USC with IGF1 knockdown (USC^IGF1) for 1 day, 3 days, 5 days and 7 days. B. Growth curves of USC^IGF1, normal USC, and USC^mock cells. C. Protein expression of IGF1 in normal USC, USC^mock, and USC^IGF1. Bar =100 μm. Data are shown for 3 independent experiments with 3 replicates. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01.

Myogenic differentiation of USC^IGF1

USC^IGF1 showed a distinct myogenic differentiation capacity, as demonstrated by the expression of myogenic markers, as indicated by immunofluorescence staining (Figure 2A). After culture for 2 weeks in differentiation medium, USC^IGF1 produced multinucleated myotubes, whereas no distinct myotubes were observed from USC cultured under the same conditions (Figure 2A). Compared with those of hSMPC, the multinucleated myotubes of USC^IGF1 were clearly delineated by brightfield imaging (Figure 2B). After 2 weeks of differentiation, USC^IGF1, but not USC, exhibited increased expression of later-stage myogenic differentiation genes, such as myogenin, desmin, and myosin (Figure 2C). These data suggested that IGF1 expression can significantly enhance the later stages of myogenic differentiation and maturation capacity of USC.

Figure 2.

Myogenic marker expression of differentiated USC^IGF1. A. Immunostaining analysis of the myogenic proteins (Myogenin, Desmin, Myosin) in differentiated USC^IGF1 cells compared to those in normal USC, USC^mock and human skeletal muscle progenitor cells (SMPC). B. Phase contrast images of USC at 14 days of differentiation (left) and the magnified area showing the multinucleated myotube structure (right). C. RT-PCR analysis of the expression of myogenesis-associated proteins in USC^IGF1 after 14 days of induced differentiation. Bar =50 μm. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05.

Secretory profile of USC^IGF1

The angiogenic secretory profiles of both USC^IGF1 and USC were analyzed and compared to further characterize the proteomic effects of IGF1 overexpression in USC. In total, 11 of the 22 growth factors and cytokines (i.e. angiogenin, angiopoietin2, CXCL16, endostatin, endothelin1, GM-CSF, IGFBP2, IL8, MCP1, PDGF-AA, and CXCL14) were found to be more highly secreted by USC^IGF1 than USC (Figure 3A, 3B). The greatest differences in secreted factors included interleukin-8 (IL8) and monocyte chemotactic factor-1 (MCP1), which were found to be strongly increased in USC^IGF1 (Figure 3B). All of the upregulated and downregulated factors are listed in Table 1. Notably, the IGF1 secretion level in the medium did not significantly differ between USC^IGF1 and USC (Figure 3C), which suggested that the alteration in USC^IGF1 function may be due to increased receptor uptake and transcriptional activity rather than an increased concentration of IGF1 in the extracellular environment.

Figure 3.

Angiogenic factors secreted by USC^IGF1. (A) Angiogenic protein array analysis of cytokines and growth factors secreted by USC^IGF1 compared with those secreted by USC. (B) Quantification of the gray values of the angiogenic factor dots detected in (A). (C) Concentration of extracellular IGF1 secreted into the medium within 7 days of culture. One-way analysis of variance (ANOVA) and Student’s t-test were used for statistical analysis, *P<0.05, **P<0.01, ***P<0.001, N.S. indicates no significant difference.

Table 1.

Upregulated and downregulated angiogenesis factors secreted by USC^IGF1 compared to USC

Up regulated	Down regulated
MCP-1	TSP-1
IL8	uPA
CXCL14	IGFBP3
IGFBP2	IGFBP1
PDGF-AA	DPPIV
CXCL16	FGF2
Endostatin	Amphiregulin
Endothelin-1
Angiogenin
Angiopotoietin-2
GM-CSF
MMP-9
VEGF

Myofusion

Myofusion assays were performed between the mCherry expressing USC^IGF1 and hSMPC^GFP. The number of fused cells significantly increased when USC^IGF1 was cocultured with hSMPC^GFP (i.e. 23 fused myotubes out of 2×10⁵ USC^IGF1) at a 1:2 ratio after 7 days of myogenic differentiation (Figure 4A, 4B). Heterogenous cell fusion between USC^IGF1 and mSMPC^GFP was originally detected 3 days after the switch to differentiation media and continued to occur on Day 14. The distinct fusion of human USC^IGF1 with mSMPC^GFP was assessed by immunostaining human mitochondria, lamin, and dystrophin 3 (DYS3) in the fused myotubes (Figure 4C). In contrast, the USC and USC^mCherry controls developed only 1 detectable myotube out of 2×10⁵ cells. Cytoplasm and mitochondria were observed in the fused myotubes with mCherry-labeled USC^IGF and hSMPC^GFP (Figure 4D).

Figure 4.

USC^IGF1 was fused with human skeletal muscle progenitor cells (hSMPC) and mouse skeletal muscle progenitor cells (mSMPC) in vitro. When cocultured in myogenic induction medium. A. Human skeletal muscle progenitor cells at ratios of 1:2, 1:1, 2:1, and 4:1 for coculture and induced differentiation. The nuclei were counterstained with DAPI, and the white arrow indicates multinucleated myotubes formed by the fusion of USC^IGF1-mCherry and hSMPC^GFP; Bar =50 μm. B. Fusion rate of hSMPC^GFP and USC^IGF1-mCherry in mixed culture. C. USC^IGF1 mixed with mouse skeletal muscle progenitor mSMPC-GFP cells at a ratio of 2:1 in coculture: USC^IGF1 was stained with the human-specific antibodies Lamin, Mitochondria, and Dystrophin (DYS). The white arrow indicates the location of myotubes that were heterogeneously formed by the fusion of USC^IGF1 and mSMPC^GFP. D. USC^IGF1-mCherry and hSMPC^GFP fused to form multinucleated myotubes, and the enlarged area indicates the field of fused myotubes in the right column (upper layer). Mitochondria pre-stained USC^IGF1 (USC^IGF1-mitotracker) was fused with hSMPC^GFP to form multinucleated myotubes, and the enlarged area indicates the area of fused myotubes in the right column (lower layer). Bar =50 μm. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05.

Gene expression alterations in USC^IGF1

mRNA microarray analysis was used to determine changes in the expression of the USC^IGF1 gene compared to that in mock USC. Modulated gene expression was ranked by fold change with P<0.01. The upregulated genes that fit these criteria included CKLF, ANGPTL4, PTTG1, C6orf173, UBE2C, ASPM, PRC1, DCBLD2, IGF1, PLAT, ANLN, CCNB2, PTTG3P, CEP55, HMMR, COL1A1, CDC20, TOP2A, TYMS, WNT5A, LTBP2, and TPM1, and the downregulated genes included KANIP4, SYNE2, GABARAPL1, MAOA, SERPINA3, BBOX1, CXCL14, and SLPI (Figure 5A). Further analysis of the cellular component (CC) (Figure 5B), biological process (BP) (Supplementary Figure 1A), and molecular function (MF) (Supplementary Figure 1B) terms, which revealed the cellular location of the active genes, showed that the most prominent components were involved in mitochondria and associated respiratory and metabolic processes, enzyme activation, and cadherin binding. We selected 15 genes whose expression was upregulated and 15 genes whose expression was downregulated in USC^IGF1 versus USC and confirmed these changes by RT-PCR (Figure 5C, 5D).

Figure 5.

The genes with the greatest change in expression after IGF1 overexpression. A. Heatmap of the gene clusters with significant expression changes (log (FC)>2, P<0.01) by GO analysis for RNA microarray. B. Cell component analysis for gene clusters with significant expression changes (log (FC)>2, P<0.01). C. qRT-PCR analysis of the top 15 genes upregulated in USC^IGF1 compared with those of normal USC. D. qRT-PCR analysis of the top 15 genes associated with the genes downregulated in USC^IGF1 compared with those in normal USC. Data are shown for 3 independent experiments with 3 replicates. One-way analysis of variance (ANOVA) and Student’s t-test were used for statistical analysis, *P<0.05, **P<0.01.

Mitochondrial stress and respiration

The Seahorse XF cell Mito Stress Test (HST) showed distinct differences in the oxygen consumption rate (OCR) between USC^IGF1 and USC, with USC^IGF1 showing greater maximal respiration (P<0.01) and ATP production (P<0.01) and greater spare respiration capacity and nonmitochondrial oxygen consumption than USC. However, no significant differences in the basal respiration capacity and proton leak were found between the USC and USC^IGF1 (Figure 6).

Figure 6.

The oxygen consumption rate (OCR) increased in USC^IGF1 patient. (A) OCR profile plot, (B) basal respiration, (C) ATP-linked respiration, (D) maximal respiration, (E) spare capacity, (F) proton leakage, and (G) nonmitochondrial respiration. n=4, One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01, N.S. indicates no significant difference.

The mTOR and AMPK signaling pathways in the myogenic differentiation of USC^IGF1

The Akt-mTOR signaling pathway is a known effector of IGF signaling. We investigated the role of IGF1-Akt-mTOR signaling in mediating the potential of USC^IGF1 to fuse with SMPCs (Figure 7). Protein harvested from hSMPC was run alongside USC and USC^IGF1 as a control. Of the proteins that are downstream of mTOR, S6K, 4EBP1 and their active phosphorylated forms p-S6K and p-4EBP1 were upregulated in the USC^IGF1 group. The levels of proteins in the Akt pathway, including phosphorylated p-Akt (S473), p-GSK3β, and p-FoxO1/3α were increased in the USC^IGF1 group. In contrast, Akt (p-T308) was downregulated in the USC^IGF1 group compared with the control USC group. Interestingly, proteins associated with autophagy that are regulated by mTOR signaling are known to be involved in skeletal muscle metabolism and cell fusion. Our data showed that the autophagy-associated proteins phosphorylated ULK (p-ULK), LC3I, and LC3II were significantly upregulated in USC^IGF1 compared to USC.

Figure 7.

The signaling pathways involved in the myogenesis of USC^IGF1. USC^IGF1 cells at passage 4 were examined to detect the protein expression and phosphorylation of the mTOR, Akt, AMPK and autophagy pathways and were compared with those of the corresponding normal USC as a negative control and human skeletal muscle progenitor cells as a positive control. The lower row shows the grey values of the western blot results. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01, N.S. indicates no significant difference.

Effect of USC^IGF1 on skeletal muscle tissue repair

To evaluate the potential contribution of USC^IGF1 in tissue repair, we injected USC^IGF and USC cells with mCherry fluorescence (USC^IGF-mCherry) into the CTX-injured tibialis anterior (TA) muscles of nude mice. H&E staining confirmed muscle injury and showed tissue regeneration at 7 days after USC^mCherry and USC^IGF1-mCherry injection (Figure 8A). There was significantly less inflammatory cell infiltration at the site of injury in muscle tissue injected with USC^IGF1 than in that injected with PBS or USC 7 days after injection (Figure 8B), normal skeletal muscle was used as a control (Supplementary Figure 2).

Figure 8.

Histochemical analyses of CTX injured skeletal muscle after USC^IGF1 therapy. A. Hematoxylin and eosin (HE) staining of CTX-injured TA muscle within 7 days after USC^IGF1 injection. Bar =100 μm. Mark 1: immune cell infiltration zone. B. Ratio of the inflammatory area to the myotube area and quantification of immune cells among the total cells within 1 mm². Data are shown for 3 independent experiments with 3 replicates. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01, ***P<0.001, N.S. indicates no significant difference.

Immunofluorescence staining for CD86 in M1 macrophages (proinflammatory) and for CD206 in M2 macrophages (anti-inflammatory) revealed distinct differences in macrophage subtypes between the the USC^mCherry (high M1) and USC^IGF1-mCherry (high M2) groups (Figure 9A, 9B). Desmin, an intermediate filament associated with the contractile sarcomeric complex in skeletal muscle was used to indicate regenerating and mature myofibers after injury. Muscles from the USC and USC^IGF1 treatment groups presented more Desmin+ myofibers at the site of injury than those from the PBS- injected group (Figure 9C). Quantification of the results revealed decreased expression of CD86, an M1 macrophage marker, and increased expression of CD206, an M2 macrophage marker, in CTX-injured skeletal muscle tissues within 7 days of injury, as indicated by the significantly increased CD206/CD86 ratio in the USC^IGF1 group as compared to the USC^mCherry and PBS groups. Quantification of Desmin staining revealed significantly greater desmin expression in the USC^IGF1-mCherry group than in the USC^mCherry and PBS groups at 1, 3, and 5 days postinjection (Figure 9D). TA tissues with USC^IGF1-mCherry injected therapy after injury showed Desmin positive staining within 7 days (Figure 9E). Electromyography demonstrated an increased amplitude, corresponding to muscle contraction, in muscles that had been injected with USC^IGF1-mCherry compared to those in the other treatment groups (Figure 10A, 10B).

Figure 9.

In vivo detection of USC^IGF1 in a nude mouse model of CTX-induced tibialis muscle injury. A. Immunofluorescence staining of the proinflammatory marker CD86 (green) in tibialis-injured nude mouse models within 7 days after injection of USC^IGF1, USC or PBS; nuclei were stained with DAPI. B. Immunofluorescence staining of the anti-inflammatory marker CD206 (green) in tibialis-injured nude mouse models within 7 days after injection of USC^IGF1, USC or PBS; nuclei were stained with DAPI. C. Immunofluorescence staining of the myogenesis marker Desmin (green) in tibialis injury model nude mice within 7 days after injection of USC^IGF1, USC or PBS; nuclei were stained with DAPI. D. Calculations of the CD206/CD86 ratio and the desmin-positive staining ratio in tibialis injury nude mice models within 7 days after injection of USC^IGF1, USC or PBS; nuclei were stained with DAPI. Representative images are shown from 3 independent experiments with at least 3 replicates. Histograms are the means ± SEM of four experiments under each independent culture condition. E. Desmin (green) staining for USC^IGF1 (mCherry) within 7 days of USC^IGF1 therapy. The results are presented as the mean ± SEM. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01, and N.S. indicate no significant difference. Bar =100 μm.

Figure 10.

Electrophysiological features of USC^IGF1 injected into the CTX-injured TA site. (A) Electrophysiological analyses and calculation of the relative amplitude ratio in CTX injured TA muscle within 7 days after USC^IGF1 injection therapy compared to injections of mSPC, USC or PBS. (B) Relative amplitube ratio of electromyography from (A) compared with normal skeletal muscle. One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01, ***P<0.001, N.S. indicates no significant difference. Bar =100 μm.

Finally, the levels of the inflammation-associated cytokines IL1β, IL6, and TNFα in the injured TA were examined at 1, 3, 5, and 7 days after PBS, USC^mCherry, and USC^IGF1-mCherry injection (Figure 11A). Quantification of these inflammatory cytokines showed that muscles treated with USC^IGF1-mCherry had significantly lower levels of these proinflammatory cytokines postinjection than did those treated with PBS or USC^mCherry (Figure 11B).

Figure 11.

Inflammation cytokine expression in CTX-injured mouse TAs after USC and USC^IGF1 treatment. (A) Expression of IL1β, IL6, and TNFα at 1 d, 3 d, 5 d, 7 d after USC and USC^IGF1 were injected into the TAs of CTX-injured mice. (B) Quantification of the expression of IL1β, IL6, and TNFα at 1 d, 3 d, 5 d, 7 d in (A). One-way analysis of variance (ANOVA) and Tukey’s test were used for statistical analysis, *P<0.05, **P<0.01, ***P<0.001, N.S. indicates no significant difference.

Taken together, these results showed that USC^IGF1 enhanced recovery from the CTX-induced inflammatory response in the skeletal muscle of nude mice and promoted the regeneration of skeletal muscle after injury.

Discussion

USC is a novel and easily accessible source of adult stem cells that possess a potent differentiation capacity, suggesting their utility as a regenerative therapy. USC are capable of myogenic differentiation under specific induction conditions and secrete a series of angiogenic proteins and growth factors that are able to enhance regeneration of skeletal muscle after injury [17]. However, the mechanism through which USC is involved in myogenesis has not been fully defined.

Primary factors affecting skeletal muscle regeneration commonly include the secretion of chemokines that recruit host stem cells to the site of injury, myogenic differentiation of local stem cells, and the ability of myogenic cells to fuse to form new myofibers [31]. In this study, we verified that the overexpression of IGF1 in USC greatly improved their ability to undergo myo-differentiation and cell fusion to form myotubes and modulated the secretome profile of the USC. These changes are likely due to increased signaling through the IGF1-Akt-mTOR signaling pathway, which promotes an anti-inflammatory tissue environment, resulting in enhanced muscle regeneration after acute injury.

IGF1 plays a critical role in promoting cell viability, myogenesis, and muscle regeneration after injury [32]. In this study, we examined the characteristics of USC after IGF1 overexpression and found that the altered genes were involved in the regulation of the cell cycle, protein synthesis, angiogenesis, and inflammatory and immune regulation processes. The genes that were upregulated the most in our screen included ANGPTL4, CCND1, IL8, SERPINE1, and IL6. ANGPTL4 functions as a serum hormone that regulates insulin sensitivity, glucose homeostasis, and lipid metabolism. CCND1 functions as a CDK kinase that is required for the cell cycle G1/S transition. IL8 serves as a chemotactic factor that guides inflammatory and immune reactions and triggers the Akt signaling pathway. SERPINE1 is related to angiogenic gene expression, cell adhesion and ECM remodeling. IL6 acts in a paracrine manner and is secreted in the microenvironment to induce SC proliferation and fusion [33]. Increasing exogenous IL6 stimulates myogenesis in muscle stem cells, myonuclear accretion and myotube maturation [34].

USC^IGF1 secreted angiogenic factors, of which MCP1, IL8, CXCL14, and IGFBP2 were the most highly upregulated. MCP1 regulates the immune response by attracting monocytes and basophils and activating the Akt signaling pathway. IL8 and CXCL14 secretion was high, but although IGF1 overexpression in USC promoted the increased secretion of these two immune response regulators, USC^IGF1 downregulated the gene expression of CXCL14. Intriguingly, we found that IGF1 was not secreted at a distinctly high level by USC^IGF1 and that IGFBP2 was significantly upregulated. These findings indicated that the changes in myogenic capacity we observed in USC in which IGF1 was overexpressed may not be due to increased secretion of IGF but rather to systemic regulation of combined gene and protein networks in the downstream signaling pathways triggered by IGF1.

Skeletal muscle injuries generally follow three phases as they heal: 1) the acute inflammatory and degenerative phase, 2) the tissue repair phase, 3) the tissue remodeling phase [35]. It is critical to treat acute muscle impairment during the inflammatory phase; 3-5 days after injury is often the optimal window for timely treatment [36]. In the present study, after injection into the injured muscle, USC^IGF1 migrated to the site of injury, resulting in decreased expression of inflammatory cytokines and promotion of macrophage polarization from the M1 to M2 phenotype 1 week after injury, as determined by an increase in the CD206/CD86 (M2/M1) ratio. Compared with USC alone, USC^IGF1 recruited more anti-inflammatory macrophages (expressing CD206) during an earlier stage of regeneration. These findings indicated that USC^IGF1 has an anti-inflammatory effect on the recovery of skeletal muscle tissue after injury. IGF1 is involved not only in myogenic regeneration but also in neurogenesis, revascularization and immune regulation in vivo [22,37]. In further studies, we will evaluate the long-term therapeutic effects of this combination of stem cell therapy and gene therapy on the signaling pathways mediating IGF-1-induced muscle and nerve tissue regeneration and angiogenesis.

IGF1, promyogenic factor, significantly enhanced myogenic differentiation and cell fusion between USC and skeletal muscle cells, as demonstrated by the significant increase in fusion compared to that in USC alone (1 out of 2×10⁵ USC versus 23 out of 2×10⁵ USC^IGF1). In addition, mitochondria from USC^IGF1 were observed in the cytoplasm of fusion cells and multinucleated myotubes, and healthy mitochondria from exogenous stem cells regulate myogenesis [38], which provides an effective strategy for stem cell therapy for muscle dysfunction. Recent evidence suggests that mitochondrial fission and fusion are associated with mitochondrial dynamics, which modulate the shape, size, and number of mitochondria and contribute to mitochondrial respiration [39,40]. Mitochondrial respiration, as indicated by the OCR, can be used to measure mitochondrial bioenergetics [41]. In addition, intercellular mitochondrial transfer occurs during cell fusion [42]; thus, the significant difference in spare respiratory capacity may be associated with the energy consumption of mitochondrial transfer via cell fusion. An increase in the OCR in USC^IGF1 is associated with a shift in mitochondrial dynamics toward fusion, which indicates the connection between cellular metabolism and mitochondrial functions.

There are 2 primary signaling pathways involved in regulating protein synthesis involved in skeletal muscle regeneration: 1) the IGF1-Akt-mTOR pathway, which acts as an inducer of protein synthesis; and 2) the Myostatin-Smad2/3 pathway, which acts as an inhibitor of protein synthesis [20]. Previous studies have shown that mitochondria play key roles in myogenic processes through the AMPK-FoxO3 and Akt signaling pathways, which regulate the repair of atrophied muscle. Intercellular transfer of mitochondria following cell fusion enhances muscle healing after injury by replacing the damaged mitochondria, which restores normal homeostasis within the cell and reduces the production of inflammatory factors while accelerating phagocytic ability [43-45]. Therefore, it is important to determine the roles of the IGF1-Akt-mTOR pathway and the transfer of healthy mitochondria via cell fusion in alleviating the myogenesis of human USC for muscle repair by preventing cellular apoptosis and death, promoting cell growth, and promoting myodifferentiation around the injured region.

IGF1 participates in skeletal muscle development by activating the Akt signaling pathway and the downstream mTOR signaling pathway. The activated signaling pathways and their associated downstream pathways indicate the pivotal role of IGF1 in the protein interaction network and enhanced cell survival and differentiation. In this study, we found accelerated cell fusion and mitochondrial transfer from USC to fused human or mouse myotubes with IGF1 overexpression, and the clustered gene pool focused on mitochondria-mediated cell metabolism. An increasing number of studies have shown that AMPK/AKT-mTOR signaling is critical for modulating mitochondrial activity. The mTOR complex controls mitochondria-associated endoplasmic reticulum membrane integrity and mitochondrial function through the protein phosphorylation process mediated by Akt activity [46].

The IGF1-Akt-mTOR pathway, which stimulates myogenesis, is the major signaling pathway that regulates skeletal muscle growth [19,47,48]. Severe myopathies and a reduction in fast muscle fibers were observed in postnatal mice harboring a muscle-specific mTOR knockout [49]. These studies indicate the prominent role of the IGF1-Akt-mTOR pathway in modulating skeletal muscle development and regeneration and emphasize IGF1 as a switch that promotes myogenesis. However, the mechanisms of IGF1 activation and the potential function of IGF1-stimulated modulation in the muscle microenvironment mediated by exogenous adult stem cells have not been fully illustrated. In our study, we found that IGF1 overexpression in USC modified the downstream gene and protein expression profiles of these cells though activation of Akt signaling via phosphorylation of the S473 site. IGF1 binding promotes skeletal muscle development by activating the downstream mTOR pathway through phosphorylation of S6K and upregulation of 4EBP1, sequentially affecting autophagy-associated signaling and regulating protein synthesis [50]. In addition, phosphorylated S6K increases ribosomal protein-related transcription, thus supporting the required cell metabolism in skeletal muscle cells [51].

Through its downstream effector FoxO3, Akt connects growth factor signaling to myofiber formation and degradation and consequently activates AMPK signaling to control cell metabolism [52]. The activation of AMPK leads to functional recovery of mitochondrial transfer to myoblasts and damaged myotubes. In this way, USC^IGF1 proliferation and differentiation were modulated, thus facilitating the myogenesis of USC^IGF1 in vitro and further fusion with myocytes to form mature myotubes. Interestingly, we found that the cell fusion rate of USC and SMPC did not increase when recombinant IGF1 protein was added to the induction medium at concentration ranging from 10 to 100 μg/ml (data not shown), suggesting that exogenous IGF1 is not limiting factor in the myogenic process but rather a more complex network associated with USC^IGF1 and accompanying intracellular alterations in cell signaling and/or metabolism may play a crucial role in muscle regeneration, especially during the maturation period. In addition to its function in myogenic regeneration, IGF1 has also been implicated in neurogenesis, revascularization, and immune regulation in vivo [22,37].

While CTX-induced injury offers a controlled environment, it may not fully represent the complexity of diverse human muscle injuries. Further studies in larger animal models with clinically relevant injuries are needed for broader translational potential. In addition, the study primarily focused on short-term regeneration. Long-term studies are crucial to assess sustained functional recovery and potential adverse effects. Furthermore, although the study demonstrates myogenic potential, further characterization of long-term differentiation stability and potential safety of USCIGF1 such as tumorigenicity is needed. While IGF1 shows promise, exploring the combined effects of additional factors or optimized gene delivery systems could enhance efficacy.

Based on the promising results, further preclinical studies in larger animals with clinically relevant injuries are warranted to pave the way for clinical trials. Investigating the synergistic effects of USC^IGF1 with other therapeutic approaches, such as physical therapy or biomaterials, could improve overall muscle regeneration outcomes. Exploring patient-specific USC lines or tailoring growth factor expression based on individual needs could personalize treatment for optimal outcomes. The findings on immunomodulatory effects of USC^IGF1 could be explored for treating muscle diseases with inflammatory components.

This study provides a significant step forward in understanding the potential of USC^IGF1 for muscle regeneration. Addressing the limitations through further research and exploring the identified prospective avenues will be crucial for translating this promising therapy into clinical reality for various muscle injuries and defects. In further studies, we will evaluate the long-term therapeutic effects of this combination of stem cell therapy and gene therapy on the signaling pathways mediating IGF1-induced muscle and nerve tissue regeneration and angiogenesis for potential use in treating urinary incontinence.

Conclusion

Our data demonstrated that USC^IGF1 can affect myogenesis via three major mechanisms: 1) the secretion of cytokines, growth factors and chemokines to repair tissue injury and promote in situ myogenesis; 2) the direct interaction of USC^IGF1 with muscle cells, promoting cell fusion to form myotubes; and 3) myogenic differentiation and myotube formation. The present study revealed the essential role of IGF1 in regulating USC function in myogenesis and skeletal muscle regeneration via the IGF1-Akt-mTOR signaling pathway. However, stem cell or gene therapy alone has limitation in promoting long-term recovery from muscle injury. A combination of USC and IGF1 therapy might provide an efficient approach for skeletal muscle regeneration in the treatment of neuromuscular injury or other types of muscle-associated defects, such as stress urinary incontinence caused by urethral sphincter muscle dysfunction due to childbirth in women or radical prostatectomy in men.

Disclosure of conflict of interest

None.