- Email: [email protected]
Exosomes from differentiating human skeletal muscle cells trigger myogenesis of stem cells and provide biochemical cues for skeletal muscle regeneration
Journal of Controlled Release 222 (2016) 107–115
Contents lists available at ScienceDirect
Journal of Controlled Release journal homepage: www.elsevier.com/locate/jconrel
Exosomes from differentiating human skeletal muscle cells trigger myogenesis of stem cells and provide biochemical cues for skeletal muscle regeneration Ji Suk Choi a, Hwa In Yoon a,b, Kyoung Soo Lee a, Young Chan Choi a, Seong Hyun Yang a, In-San Kim b,c, Yong Woo Cho a,⁎ a b c
Department of Chemical Engineering, Hanyang University, Ansan, Gyeonggi-do 426-791, Republic of Korea Center for Theragnosis, Biomedical Research Institute, Korea Institute of Science and Technology (KIST), Seoul 136-791, Republic of Korea KU-KIST School, Korea University, 1 Anam-dong, Seongbuk-gu, Seoul 136-701, Republic of Korea
a r t i c l e
i n f o
Article history: Received 13 March 2015 Received in revised form 3 December 2015 Accepted 12 December 2015 Available online 14 December 2015 Keywords: Exosomes Nanovesicles Stem cells Myogenesis Muscle regeneration
a b s t r a c t Exosomes released from skeletal muscle cells play important roles in myogenesis and muscle development via the transfer of specific signal molecules. In this study, we investigated whether exosomes secreted during myotube differentiation from human skeletal myoblasts (HSkM) could induce a cellular response from human adipose-derived stem cells (HASCs) and enhance muscle regeneration in a muscle laceration mouse model. The exosomes contained various signal molecules including myogenic growth factors related to muscle development, such as insulin-like growth factors (IGFs), hepatocyte growth factor (HGF), fibroblast growth factor-2 (FGF2), and platelet-derived growth factor-AA (PDGF-AA). Interestingly, exosome-treated HASCs fused with neighboring cells at early time points and exhibited a myotube-like phenotype with increased expression of myogenic proteins (myosin heavy chain and desmin). On day 21, mRNAs of terminal myogenic genes were also up-regulated in exosome-treated HASCs. Moreover, in vivo studies demonstrated that exosomes from differentiating HSkM reduced the fibrotic area and increased the number of regenerated myofibers in the injury site, resulting in significant improvement of skeletal muscle regeneration. Our findings suggest that exosomes act as a biochemical cue directing stem cell differentiation and provide a cell-free therapeutic approach for muscle regeneration. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Exosomes are small membrane vesicles (40–120 nm in diameter) secreted in a variety of mammalian cells following the fusion of multivesicular bodies (MVBs) with the plasma membrane during endosome maturation [1]. Exosomes derived from cells contain proteins and nucleic acids that reflect their original cell sources and transfer different cellular information to neighboring cells or even to distant tissues [2–4]. The interest of scientists in the role of exosomes has expanded rapidly because of their diverse pathological and therapeutic effects. Tumorderived exosomes play important roles in tumorigenesis, metastasis, angiogenesis, and generating tumor-associated cells by transporting growth factors/receptors or oncogenes. As exosomes can be key mediators of intercellular communication in tumors, their inhibition has become a focus of research in cancer therapy [5–8]. Several groups [9–14] have shown the therapeutic potential of exosomes derived ⁎ Corresponding author at: Department of Chemical Engineering, Hanyang University ERICA Campus, 55 Hanyangdaehak-ro, Sangnok-gu, Ansan, Gyeonggi-do 426-791, Republic of Korea. E-mail address: [email protected] (Y.W. Cho).
http://dx.doi.org/10.1016/j.jconrel.2015.12.018 0168-3659/© 2015 Elsevier B.V. All rights reserved.
from various cells with secretory capacity, although the therapeutic mechanisms of exosomes are still unclear. Stem cell-derived exosomes are a complex biological cargo that contains a range of stem cellderived secretomes, making them an ideal candidate for treatment of various disorders, such as liver disease [9], kidney injury [10], lung injury [11], myocardial infarction [12,13], and cutaneous wounds [14]. In addition to stem cell-derived exosomes, nerve cell-derived exosomes enhanced axonal regeneration in the peripheral nervous system [15] and preosteoblast-exosomes also increased the gene levels of neuroectoderm differentiation in embryonic stem cells [16]. Skeletal muscle is the largest organ in the human body, consisting of aligned multinucleate myofibers surrounded by basal lamina and satellite cells located beneath muscle fibers [17]. In addition to basic functions, such as energy metabolism, homeostasis and locomotion, skeletal muscle secretes hormone-like factors, termed ‘myokines’ that regulate angiogenesis and myogenesis within the muscles (autocrine/ paracrine) or communicate systemically with other tissues (endocrine) [18,19]. Skeletal myoblasts and myotubes secrete exosome-like nanovesicles, and these nanovesicles contain various myokines, proteins, microRNAs, and mitochondrial DNA, which could act as critical signals for skeletal muscle myogenesis, homeostasis, and development
108
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
[20–22]. Other studies also have shown that C2C12-derived exosomes stimulate survival and neurite outgrowth of motor neurons by paracrine signals [23]. Despite these findings on the secretory function of muscle cells, the biological roles of skeletal muscle cell-derived exosomes are not well understood yet. In this study, we hypothesized that exosomes secreted during the differentiation of human skeletal myoblasts (HSkM) into myotubes may contain specific biochemical cues that promote and regulate the differentiation of human adipose-derived stem cells (HASCs) with myogenic potential. Moreover, these exosomes could act as a therapeutic agent associated with muscle growth/repair, leading to efficient regeneration of damaged skeletal muscle. 2. Materials and methods 2.1. Human skeletal myoblast culture and differentiation HSkM (catalog #T4068, Lot# HC0512, sourced from post-gestational tissue, usually from the quadriceps or psoas tissue) was obtained from Applied Biological Materials Inc. (Richmond, BC, Canada). Growth medium and differentiation medium were purchased from PromoCell (Heidelberg, Germany). HSkM were maintained in growth medium supplemented with 5% fetal calf serum (FCS), 50 μg/mL petuin, 10 ng/mL epidermal growth factor, 1 ng/mL basic fibroblast growth factor, 10 μg/mL insulin, 0.4 μg/mL dexamethasone, and 1000 UI/mL penicillin/streptomycin (P/S) at 37 °C in humidified air containing 5% CO2. Differentiation was induced with serum-free medium containing 10 μg/mL insulin and 1000 UI/mL P/S for 5 days.
manufacturer's protocol. The exosome pellet was resuspended in phosphate buffered saline (PBS) and purified using exosome spin columns (MW 3000, Invitrogen). Exosomes isolated from proliferating HSkM were used as a control. The exosomal total protein content was quantified using the micro bicinchoninic acid (BCA) protein assay (Pierce Biotechnology, Rockford, IL, USA). Isolated exosomes were characterized with the minimal experimental requirements for extracellular vesicles (EVs) [24]. 2.3. Dynamic light scattering Dynamic light scattering (DLS) measurements were performed at 37 °C, using Zetasizer Nano ZS90 (Malvern, Worcestershire, UK). Exosomes resuspended in PBS were placed in a UV-transparent cuvette (Sarstedt AG & Co., Germany). The size distribution plot with an x-axis showing the distribution of estimated particle radius (nm) and a y-axis showing the relative intensity of the scattered light was analyzed with software. 2.4. Transmission electron microscopy Exosomes were fixed in 0.5% glutaraldehyde solution overnight. They were subsequently centrifuged at 13000 × g for 3 min, and then the supernatant was removed. The sample was dehydrated in absolute ethanol for 10 min and collected on formvar grids stabilized with carbon (TED PELLA, Inc., Redding, CA, USA). The grids were contrasted with 1% phosphotungstic acid for 1 min and then examined using transmission electron microscopy (TEM) with a JEM-2100F field emission electron microscope (JEOL Ltd., Japan).
2.2. Exosome isolation from differentiating HSkM 2.5. Western blotting Conditioned media (CM) were collected during differentiation of HSkM and used for exosome purification (Fig. 1A). Briefly, CM were collected from cells cultured in differentiation medium every day during 5 days. Collected CM were centrifuged (10 min, 2000 ×g) to eliminate cell debris. The resulting supernatant was concentrated through an Amicon® Ultra-15 3 K centrifugal filter (Millipore, Billerica, MA, USA) at 5000 ×g for 60 min. Exosomes were isolated using the Total Exosome Isolation kit from Invitrogen (Carlsbad, CA, USA) according to the
Differentiated HSkM and exosomes were lysed in RIPA buffer (25 mM Tris–HCl, pH 7.6, 150 mM NaCl, 0.5% Trixton X-100, 1% Na-deoxycholate, 0.1% sodium dodecyl sulfate and protease inhibitor cocktail). The protein in the supernatant was quantified using the BCA protein assay, electrophoresed on a 12–15% SDS-PAGE gel, and transferred to a PVDF membrane (Millipore, Billerica, MA). The membrane was blocked with 5% BSA in T-TBS (10 mM Tris, 150 mM NaCl, and
Fig. 1. Characterization of exosomes released from differentiating human skeletal myoblasts (HSkM). (A) Summary of the protocol for exosome purification from conditioned medium (CM) of differentiating human myoblasts and the induction of stem cell myogenesis using the exosomes. (B) Dynamic light scattering (DLS) showing the particle size distribution of exosomes. (C) Transmission electron microscope (TEM) analysis of exosomes. The scale bars represent 100 nm (black) and 200 nm (white). (D) Western blot analysis of exosomes. Equal amounts of total proteins (30 μg) extracted from exosomes and cells were immunoblotted for Alix, TSG 101, CD63, CD81 and β-actin.
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
0.1% Tween 20) for 1 h at room temperature, and subsequently incubated with primary antibodies overnight at 4 °C. After vigorous washing in T-TBS, the blots were incubated with horseradish peroxidase (HRP)tagged secondary antibodies for 1 h. The labeled proteins were visualized with the ChemiDoc™ XRS imaging system (Bio-Rad Laboratories, Inc., Hercules, CA, USA) using an enhanced chemiluminescence (ECL) kit (Pierce). Primary (Alix, TSG101, CD63, CD81, and β-actin) and secondary antibodies were purchased from Santa Cruz Biotechnology (Dallas, TX, USA). All chemical reagents and protease inhibitor cocktail (P-2714) were purchased from Sigma-Aldrich (St. Louis, MO, USA). 2.6. Human adipose-derived stem cell isolation Human adipose tissue was obtained with informed consent, as approved by the Institutional Review Board of the Catholic University of Korea, College of Medicine. HASCs were obtained from the adipose tissue of healthy human females as described previously [25,26]. 2.7. Cell proliferation assays The proliferation of HASCs in different conditions was analyzed using calcein AM staining and WST-1 assay. HASCs were seeded at 3 × 104 cells per well into 96-well plates in DMEM containing with 10% FBS and 1% P/S and allowed to adhere overnight at 37 °C under 5% CO2. The cells were washed with PBS and then maintained in GM (growth medium; DMEM, 10% FBS, 1% P/S), DM (differentiation medium; DMEM with 5% horse serum, 0.1 M dexamethasone, 50 μM hydrocortisone, and 1% P/S), GFM (growth factors containing medium; serum-free DMEM with 106 pg/mL hepatocyte growth factor; HGF, 65 pg/mL vascular endothelial growth factor; VEGF, 68 pg/mL insulin like growth factor; IGF, 100 pg/mL basic fibroblast growth factor; bFGF, and 63 pg/mL interleukin-6; IL-6) or EM (serum-free DMEM containing exosomes with 0, 10, 25, 50, 100, and 200 μg/mL) for 5 days. The concentrations of recombinant growth factors (Abcam, Cambridge, UK) in the 50 μg/mL exosomes were specified in the study. The cells were stained for 30 min with the calcein AM reagents (Sigma-Aldrich) on day 3 and observed using a fluorescence microscope (IX81; Olympus, Tokyo, Japan). Green fluorescence caused by the reaction of calcein with intracellular esterase indicated live cells. These experiments were performed in triplicate. The WST-1 reagent (Roche Applied Science, Mannheim, Germany) was added to each well, and the plate was incubated at 37 °C for 1 h under 5% CO2. After gentle pipetting, the 100 μL solution was transferred to a 96-well plate. The optical density was then measured at 440 nm using a microplate spectrophotometer (PowerWave XS, Bio-Tek Instruments, Winooski, VT, USA).
109
collected and prepared for immunofluorescent staining and quantitative PCR analysis. 2.10. Immunofluorescent staining Cells were washed twice with cold PBS, fixed with 4% paraformaldehyde/PBS for 1 h at 4 °C, and permeabilized with 0.3% Triton X-100 for 20 min at room temperature. Cells were incubated overnight at 4 °C with primary antibodies including desmin (Santa Cruz Biotechnology) and myosin heavy chain (MHC; Abcam, Cambridge, MA, USA), and then incubated with anti-rabbit IgG-FITC secondary antibody (Santa Cruz). Cells were counterstained with 1 μg/mL DAPI (4,6-diamidino-2phenylindole; Sigma-Aldrich) for 1 min and then observed with a fluorescence microscope (IX81, Olympus Corporation, Tokyo, Japan). The fusion index is defined as the ratio of the number of nuclei inside myotubes to the total number of nuclei at day 21. 2.11. Quantitative RT-PCR The human skeletal muscle-myogenesis & myopathy RT2 Profiler PCR Array (PAHS-099Z, SABiosciences, Valencia, CA, USA) was used to evaluate the expression of genes related to myogenesis. The total RNA was isolated from cells using the RNeasy Mini kit (Qiagen) according to the manufacturer's instructions. cDNA was synthesized from 500 ng of total RNA using the RT2 First Strand Kit (SABiosciences). 84 different genes were simultaneously amplified on a Stratagene Mx3000P (Agilent Technologies, Santa Clara, CA, USA). The thresholds and baselines were set according to the manufacturer's instructions. The fold change in gene expression (compared to controls) was calculated using SABiosciences webportal software (http://www.sabiosciences. com/pcr/arrayanalysis.php). p Values were calculated using the student's t-test and the experiments were repeated in triplicate. 2.12. Mouse model of skeletal muscle injury and exosome treatment
Differentiating HSkM exosomes were labeled with a membranelabeling dye (1′dioctadecyl-3,3,3′,3′-tetramethylindodicarbocyanine, DiD) in serum-free DMEM according to the manufacturer's protocol (Molecular Probes, Eugene, OR, USA). The labeled exosome suspension was filtered with PD MiniTrap G-10 columns (GE Healthcare Life Sciences, Pittsburgh, PA, USA) and the flow-through was used as the unbound dye control. HASCs were seeded at a density of 250 cells/mm2, incubated at 37 °C with labeled exosomes (50, 100, and 200 μg/mL) for 1 h, and then observed with a confocal microscope (Leica TCS SP8, Leica Microsystems, Buffalo Grove, IL, USA). The fluorescence intensities were quantified using an image analyzer.
All experiments with live animals were performed in compliance with the relevant laws and institutional guidelines of the Korea Institute of Science and Technology (KIST), and institutional committees approved the experiments. The muscle injury animal model was based on a previous report from Shi et al. [27]. Nude mice (male, 5 weeks old, n = 8) were anesthetized with ketamine (100 mg/kg) and xylazine (4–6 mg/kg), and an adequate depth of anesthesia was maintained such that the mice were unresponsive to tactile stimulation. For muscle injury, an anterolateral skin incision was made in both legs, and the tibialis anterior muscles were exposed. The muscles were then lacerated transversely at the mid-portion using a scalpel, creating a 5-mm wedgeshaped defect. After closure of the fascia with a suture, the exosome suspension (100 μg/100 μL in PBS) was injected into the right muscle defect and the same volume of PBS without exosome was injected into the left muscle defect as a control. For morphological evaluation of muscle regeneration after muscle laceration, muscle samples were fixed in 4% paraformaldehyde, embedded in paraffin, and sliced at 6 to 10 μm using a microtome. Sections were de-paraffinized, dehydrated, and stained with Masson's trichrome. The average area of collagen deposition and total number of myofibers with centralized nuclei within the muscle injured site were measured in three random fields (× 400) of four sections/muscle using Photoshop software (Adobe Systems Inc., San Jose, CA, USA).
2.9. Myogenic differentiation of human adipose-derived stem cells
2.13. Statistical analysis
Cells were plated at a density of 250 cells/mm2 in DMEM supplemented with 10% FBS and 1% P/S. After 72 h (~ 90% confluence), cells were rinsed with PBS and then maintained in different media. The media were changed every 3 days for up to 21 days. Cells were then
Experimental data were expressed as means ± standard deviation (SD). The student's two tailed t-test was performed with SPSS 17.0 statistical software (SPSS, Chicago, IL, USA), and statistical significance was accepted at p b 0.05 or p b 0.01.
2.8. Exosome labeling and cellular uptake
110
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
Fig. 2. The cell proliferation and cellular uptake of exosomes in human adipose-derived stem cells (HASCs). (A) Proliferation of HASC in GM (growth medium; DMEM, 10% FBS, 1% P/S), DM (differentiation medium; DMEM with 5% horse serum, 0.1 M dexamethasone, 50 μM hydrocortisone, and 1% P/S), GFM (growth factor containing medium; serum free DMEM with 106 pg/mL HGF, 65 pg/mL VEGF, 68 pg/mL IGF, 100 pg/mL bFGF, and 63 pg/mL IL-6) or EM (serum-free DMEM containing exosomes with 0, 10, 25, 50, 100, and 200 μg/mL) were visualized by calcein AM staining (A) and examined using a WST-1 assay kit (B) for 5 days. (C) Light differential interference contrast (DIC) and corresponding confocal images of HASCs after 1 h incubation with 50, 100, and 200 μg/mL of DiD-labeled exosomes, respectively. Images of DiD-labeled exosomes (red) with DAPI (blue) were visualized by merging the confocal images (Merge 1) or bright-field with confocal images (Merge 2). (D) Relative fluorescence intensity of DiD-labeled exosomes internalized in cells. Scale bars represent 200 μm (yellow) and 25 μm (white). Data are shown as mean ± standard deviation (n = 3) with significance at ⁎, ⁎⁎, ⁎⁎⁎p b 0.05. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
3. Results 3.1. Characterization of differentiating HSkM exosomes Exosomes isolated from differentiating human myoblasts were characterized in terms of size, morphology, particle yield, and surface markers (Figs. 1 and S1). Exosomes recovered with total exosome isolation reagents displayed a round or cup-like shape with a mean diameter of 113.7 nm (Fig. 1B and 1C) and had a particle yield of 1.7 × 109 particles/μg protein, comparable to that of exosomes isolated with the ultracentrifugation protocol (2.1 × 109 particles/μg protein) (Fig. S1). Western blotting for the exosomal markers revealed that the exosomes were positive for exosomal markers, including Alix, TSG101, CD63, CD81, FLOT-1, ICAM, EpCAM, and ANXA5, while non-exosomal marker, GM130 was not detected (Figs. 1D and S2). The protein
composition in exosomes was analyzed using LC–MS/MS. Proteins found in exosomes include extracellular matrix proteins, membraneassociated and cell adhesion proteins, cell structure and motility proteins, growth factors, and transmembrane signaling proteins (Table S1). Growth factors in exosomes were detected using antibody arrays. In total, 8 of 41 growth factors were significantly detected in exosomes when compared to control medium (Table S2 and Fig. S4). Interestingly, HGF, which is involved in the regulation of myogenic satellite cells, was highly expressed in exosomes. We also detected growth factors that act as potential regulators of development, function, and regeneration of skeletal muscle, including amphiregulin (AR), bFGF, IGF binding protein-3 (IGFBP-3), IGF-1, IL-6, neurotrophin-3 (NT-3), platelet-derived growth factor-AA (PDGF-AA), placenta growth factor (PLGF), transforming growth factor-beta1, 3 (TGF-β1, 3), VEGF, and VEGF receptor 3 (VEGF R3) (Table S3).
Fig. 3. Assessment of myogenesis of HASCs in differentiation medium (DM) and exosome-containing medium (EM) for 21 days. (A) HASCs were maintained in DM or EM containing 50 μg/mL of exosomes for 21 days, and were then immunostained with anti-myosin heavy chain (MHC) and anti-desmin. Cells were counterstained with DAPI (4,6-diamidino-2phenylindole), which stains nuclei blue. Scale bars represent 200 μm. (B) Immunofluorescence staining for desmin- and MHC-positive HSkM cells as a control. (C) Quantification of fusion index, representing the number of nuclei in multinucleated myotubes divided by the total number of nuclei in a field, with a myotube defined by at least three nuclei on day 21. (D) Length of multinucleated myotubes in DM or EM on day 21. (E) RT2 profiler PCR array for skeletal myogenesis of HASCs in DM or EM on day 21. The fold change of the test groups (DM and EM) compared to the control group (undifferentiated HASC) was calculated from three independent experiments using (http://www.sabiosciences.com/ pcrarraydata-analysis.php). Relative gene expression was normalized to housekeeping genes and expressed as the fold change comparing to undifferentiated HASC. ADIPOQ, FGF2, IGF1, IGF2, IL6, LEP, TGFB1, and TNF genes were evaluated as markers for skeletal autocrine signaling. MYF5, MYF6, MYOD1, MEF2C, MYOG, DMD, ACTA1, ACTN3, DAG1, DES, MYH1, MYH2, TNNT1, and TNNT3 genes were evaluated as markers for skeletal myogenesis. Data are shown as mean ± standard deviation (n = 3) with significance at ⁎, ⁎⁎p b 0.05. Abbreviations: ACTA; skeletal muscle actin alpha, ADIPOQ; adiponectin, DAG1; dystroglycan 1, DES; desmin, DMD; dystrophin, FGF2; fibroblast growth factor 2, IGF; insulin-like growth factor, LEP; leptin, IL6; interleukin 6, MEF2C; myocyte enhancer factor 2C, MYOD1; myogenic differentiation 1, MYOG; myogenin, MYF; myogenic factor, MYH; myosin heavy chain, TGFB1; transforming growth factor beta 1, TNF; tumor necrosis factor, TNNT; troponin T.
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
3.2. Cell proliferation and cellular uptake The effect of the exosome dose on cell proliferation was investigated using calcein AM staining and WST-1 assay for 5 days (Fig. 2A and 2B).
111
The proliferation of HASCs treated with exosomes was compared to the initial seeding absorbance (440 nm). On day 5, HASCs cultured in either DM, GFM or EM showed a slightly lower cell proliferation capacity than that in GM. However, exosomes significantly affect cell proliferation in a
112
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
dose dependent manner. In particular, 50 μg/mL exosome-treated cells proliferated better than cells maintained in GFM or exosome-free medium. To study the cellular uptake of differentiating HSkM exosomes at early time points, HASCs were incubated with 50 to 200 μg/mL of fluorescent dye-labeled exosomes (red) for 1 h (Fig. 2C and 2D). Exosomes were internalized into the cytoplasm of HASCs within 1 h and were partly co-localized within the nuclei. These results imply that differentiating HSkM exosomes are successfully delivered into stem cells with high cellular uptake efficiency.
the regenerated myofibers continued to grow and mature, resulting in reconstruction of muscle architecture. Damaged fibers, and inflammatory cells in the exosome (−) group were still observed with a few regenerating myofibers at day 14. The results demonstrated that the exosome-treated group had a large number of regenerative myofibers with minimal fibrosis compared with the exosome (−) groups, indicating that muscle regeneration was significantly improved by differentiating HSkM exosome treatment.
3.3. Myogenesis of HASCs promoted by differentiating HSkM exosomes
Skeletal muscle has the capacity to regenerate following common sports-related injuries. However, severe injuries can result in extensive fibrosis, scarring, and loss of muscle function [28]. Various approaches using the cellular and biochemical cues associated with muscle regeneration have been explored in vitro and in vivo to enhance the regeneration capacity of injured skeletal muscle [29,30]. Therapeutic approaches using myoblasts, muscle satellite cells, stem cells of various origins, and stem cells combined with growth factors have been used to improve muscle function and regeneration [31–34]. Although cell-based therapies have clear beneficial effects on skeletal muscle repair, there are still a number of concerns, such as limited survival and the reduced regenerative capacity of engrafted cells, as well as immune-mediated rejection [35]. Exosomes secreted by various cells may be an effective cell-free therapeutic agent because of their diverse biological and physiological functions as well as their key role as vehicles of intercellular communication [1,2]. Skeletal muscle cells also secrete a large number of myokines and some nano-vesicles that influence the growth, function, and development of muscle tissue [20]. Several groups have reported that exosomes secreted from skeletal muscle cells contain more than 400 different signal molecules, including microRNAs and proteins, that are involved in many cellular processes, including myogenesis by transporting and delivering a cargo of specific signals [22,36–38]. In spite of the proteomic and genomic characterization of exosomes, their biological functions remain unclear. As previously reported [20,37, 39], exosomes isolated from differentiating HSkM showed a typical round- or cup-shaped morphology ranging in diameter from 70– 120 nm and expressed specific cell surface molecules that are involved in their biogenesis, such as Alix, TSG101, CD63, CD81, FLOT-1, ICAM, EpCAM, and ANXA5 (Figs. 1, S1, and S2). Although we did not carry out in-depth proteomic and growth factor analyses on exosome contents, differentiating HSkM exosomes also contained various proteins, such as extracellular matrix (ECM) protein precursors, cell surface proteins, cell surface receptors, transmembrane signaling proteins, and myogenic growth factors, such as the HB-EGF family (AR and HB-EGF), VEGF family (VEGF-R3 and PLGF), IGFBP-3, HGF, IGF-I/II, bFGF, IL-6, NT-3, and PDGF-AA. Interestingly, HGF, which is known to be important in the activation and differentiation of muscle satellite cells, was appreciably expressed in exosomes (Tables S1, S2, S3, and Fig. S4). We hypothesized that differentiating HSkMs secrete exosomes loaded with myogenic factors and these exosomes might regulate in vitro myogenesis of HASCs. The physiological role of exosomes and their contents in regulating stem cell differentiation are not well defined; however, several groups [14,40,41] have addressed that exosomal factors derived from various cells contribute to proliferation, migration, and differentiation of cells. For example, exosomal Wnt 4 protein secreted from human mesenchymal stem cell (MSC) promotes proliferation and migration of keratinocytes and dermal fibroblasts through activation of β-catenin signaling in cells [14]. In addition, TGF-β1 contained in cancer cell exosomes induce the differentiation of ASCs and MSCs into cancer associated myofibroblast via TGF-β1/SMAD-mediated signaling pathway [40,41]. We found that exosomes significantly contribute to myogenic differentiation of HASCs (Figs. 3, S6, and S7). Stimulation by exosomes rapidly altered the morphological phenotype
To investigate the cellular function of differentiating HSkM exosomes on HASCs, HASCs were incubated in DM, GFM, and EM for 21 days and then stained for desmin and MHC, myogenesis specific markers (Fig. 3). After 7 days of culture, the HASCs treated with 50 μg/mL of exosome started to elongate and a small proportion of desmin-staining cells were present. The HASCs maintained in GFM, EM 10 or EM 25 started to change their morphology and a small proportion of MHC-staining cells were present after 14 days. However, the differentiation efficiency of HASCs in GFM, EM 10 or EM 25 was lower than that of the HASCs in EM 50 (Fig. S7). After 21 days, elongated multinucleated HASCs expressed desmin and MHC in both DM and EM 50, and the intensities of the staining significantly increased (Fig. 3A). However, high concentrations (200 μg/mL) of exosomes resulted in high cell death (Figs. 2A, 2B, and S6). The fusion index increased from 14.34% in DM to 23.35% in EM, and the myotube length also increased from 175.46 μm in DM to 204.03 μm in EM (Fig. 3C and 3D). Although the conversion efficiency of HASCs into myotubes in EM was lower than that of the HSkMs used as a positive control (fusion index, 56.67% and myotube length, 536.92 μm for 5 days), these results indicate that differentiating HSkM exosomes provide biochemical cues to stimulate myogenesis of HASCs, and their functional effects were more significant than that of conventional differentiation medium. We assessed the expression of 49 genes associated with skeletal myogenesis of HASCs treated with differentiating HSkM exosomes using qPCR arrays. Table S4 and Fig. 3E show the list and fold change of up- and down-regulated genes in HASCs grown in DM or EM for 21 days, compared to undifferentiated HASCs (p value b 0.05). The upregulated genes in EM included skeletal myogenesis-related genes (ACTA1 and MYOD1), muscle autocrine signaling-related genes (FGF2 and TNF), and muscle contractility-related genes (DAG1, DES, MYH1/2, TNNC1, and TNNT3). There were distinct differences in gene expression between HASCs differentiated in DM and EM on day 21 (Fig. 3E). Under DM conditions, the autocrine signaling genes ADIPOQ, IL6, and LEP, and the early myogenic markers MYF5 and MEF2C were significantly upregulated, compared with EM conditions. In contrast, HASCs differentiated in EM highly expressed the terminal differentiation markers MYOD1, ACTA1, DAG1, DES, MYH1, MYH2, and TNNT1. Based on these results it appears that the differentiating HSkM exosomes can quickly deliver signaling molecules that regulate the skeletal myogenic fate of HASCs, resulting in the acceleration of myogenic differentiation of HASCs. 3.4. Muscle regeneration enhanced by differentiating HSkM exosomes To investigate whether differentiating HSkM exosomes could contribute to muscle regeneration in vivo, we injected exosomes at the lacerated muscle sites. Masson trichrome staining was used to evaluate fibrosis and muscle regeneration after exosome injection (Fig. 4). At days 7 and 14, the collagen deposition in the exosome (+) groups (42.36 ± 7.23% and 15.36 ± 6.95%) was significantly smaller compared with the exosome (−) groups (60.21 ± 10.37% and 31.31 ± 6.9%). In the exosome (+) groups, there were more regenerating myofibers with centralized nuclei than in the exosome (−) groups at day 7, and
4. Discussion
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
113
Fig. 4. Histological images of skeletal muscle regeneration after muscle cell-derived exosome injection. Histological evaluation was carried out by Masson trichrome staining. (A) Normal hindlimb muscles of mice were used as a control. Seven days post-injury, newly regenerated myofibers with centralized nuclei were observed in the exosome-treated skeletal muscle, although fibrosis between regenerated myofibers was also visible. At day 14, the exosome-treated muscle showed enhanced muscle regeneration, whereas untreated skeletal muscle remained abnormal (myofibers, red-orange color; collagenous scar tissue, blue color). Scale bars represent 1 mm (red), 200 μm (black) and 100 μm (white). (B) Quantification of collagen deposition and the number of centronuclear myofibers within the muscle injured site at days 7 and 14 (⁎, ⁎⁎p b 0.05).
of HASCs at early time points and resulted in significant expression of myogenic markers (desmin and MHC) with a fusion index of 20%, when compared with HASCs in differentiation medium. Moreover, mRNA expression of myogenic genes, such as FGF2, TNF, MYOD1, DAG1, DES, MYH1/2 and TNNT1, which are related to skeletal autocrine signaling and terminal myogenic differentiation, was increased in HASCs stimulated by exosomes over 21 days (Fig. 3E). The process of myogenesis is orchestrated by complex signaling cascades initiated by extrinsic factors, including cytokines, growth factors, and signaling molecules [42]. In particular, several paracrine growth factors, including IGFI/II, FGF2, HGF, PDGF, and TGF-β play major roles in the regulation of muscle satellite cell chemotaxis, proliferation, and differentiation through the stimulation of Pax genes and myogenic regulatory factors (MRFs; MyoD, MYF5/6, myogenin) [43–45]. From a paracrine effect perspective, it seems that differentiating HSkM exosomes containing diverse myogenic factors are efficiently delivered into stem cells, and this cargo synergistically triggers myogenic signaling pathways in recipient cells, resulting in skeletal myogenic differentiation. We also investigated the therapeutic effect of differentiating HSkM exosomes using a muscle injury model. Skeletal muscle regeneration is a complex process that is orchestrated by a collaboration of many regulatory factors. After muscle injury, some cytokines and growth factors are released from inflammatory and stromal cells. These autocrine/
paracrine factors could not only create a regenerative niche for the migration, activation, and differentiation of myoblasts and quiescent satellite cells, which is critical for muscle regeneration, but may also balance regeneration and fibrosis [46,47]. We found that differentiating HSkM exosomes accelerate skeletal muscle regeneration in a laceration mouse model by reducing the collagen deposition in injured muscle and increasing the number of regenerated myofibers after exosome injection (Fig. 4). As mentioned above, differentiating HSkM exosomes are rich in various myogenic factors such as HGF. HGF is a potent chemoattractant for muscle stem cell activation and migration to injury sites [17]. Although HGF has a dose-dependent effect on skeletal muscle repair, it has synergistic effects with other myogenic growth factors, such as FGF2, IGF1, and TGF-β, which in combination significantly increase muscle repair and regeneration compared with the individual growth factors alone [43,48]. Therefore, these results support our hypothesis that the myogenic factors contained in exosomes could effectively provide a niche favorable to the skeletal muscle regeneration by stimulating the differentiation of myoblasts and stem cells (Fig. S5). However, it remains unclear as to which factors in exosomes play a key role for controlling cell fate and promoting skeletal muscle regeneration. Moreover, we should consider the adverse effects that can arise from high doses of exosomes (e.g., cell apoptosis) because complex molecules contained in exosomes can stimulate different signaling
114
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
pathways. Nonetheless, we expect that exosomes can be an alternative approach to stem cell-based therapy and show promise as a therapeutic agent for skeletal muscle regeneration. 5. Conclusion Exosomes derived from differentiating HSkM can induce in vitro myogenesis of HASCs and improve in vivo muscle regeneration after skeletal muscle injury. Although the mechanism for the biological effects of exosomes remains unclear, exosomes may regulate skeletal myogenesis through the transfer of diverse myogenic factors, thereby controlling stem cell fate toward myogenesis and improving muscle regeneration. Therefore, we propose that exosomes can provide a cell-free therapeutic approach for regenerative medicine and for skeletal muscle regeneration in particular. Acknowledgments This work was supported by the Basic Research Program (Grant No. 2012-008294) and the Bio & Medical Technology Development Program (Grant No. 2011-0019774) through the National Research Foundation of Korea (NRF) funded by the Korean government (MEST). This study was also supported by a grant from the National R&D Program for Cancer Control, Ministry of Health and Welfare, Republic of Korea (1420390). Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.jconrel.2015.12.018. References [1] C. Théry, L. Zitvogel, S. Amigorena, Exosomes: composition, biogenesis and function, Nat. Rev. Immunol. 2 (2002) 569–579. [2] G. Raposo, W. Stoorvogel, Extracellular vesicles: exosomes, microvesicles, and friends, J. Cell Biol. 200 (2013) 373–383. [3] R. van der Meel, M.H. Fens, P. Vader, W.W. van Solinge, O. Eniola-Adefeso, R.M. Schiffelers, Extracellular vesicles as drug delivery systems: lessons from the liposome field, J. Control. Release 195 (2014) 72–85. [4] S.M. van Dommelen, P. Vader, S. Lakhal, S.A. Kooijmans, W.W. van Solinge, M.J. Wood, R.M. Schiffelers, Microvesicles and exosomes: opportunities for cell-derived membrane vesicles in drug delivery, J. Control. Release 161 (2012) 635–644. [5] J. Webber, R. Steadman, M.D. Mason, Z. Tabi, A. Clayton, Cancer exosomes trigger fibroblast to myofibroblast differentiation, Cancer Res. 70 (2010) 9621–9630. [6] L. Pascucci, V. Coccè, A. Bonomi, D. Ami, P. Ceccarelli, E. Ciusani, L. Viganò, A. Locatelli, F. Sisto, S.M. Doglia, E. Parati, M.E. Bernardo, M. Muraca, G. Alessandri, G. Bondiolotti, A. Pessina, Paclitaxel is incorporated by mesenchymal stromal cells and released in exosomes that inhibit in vitro tumor growth: a new approach for drug delivery, J. Control. Release 192 (2014) 262–270. [7] T. Smyth, M. Kullberg, N. Malik, P. Smith-Jones, M.W. Graner, T.J. Anchordoquy, Biodistribution and delivery efficiency of unmodified tumor-derived exosomes, J. Control. Release 199 (2015) 145–155. [8] P. Vader, X.O. Breakefield, M.J. Wood, Extracellular vesicles: emerging targets for cancer therapy, Trends Mol. Med. 20 (2014) 385–393. [9] S. Lemoinne, D. Thabut, C. Housset, R. Moreau, D. Valla, C.M. Boulanger, P.E. Rautou, The emerging roles of microvesicles in liver diseases, Nat. Rev. Gastroenterol. Hepatol. 11 (2014) 350–361. [10] S. Gatti, S. Bruno, M.C. Deregibus, A. Sordi, V. Cantaluppi, C. Tetta, G. Camussi, Microvesicles derived from human adult mesenchymal stem cells protect against ischaemia-reperfusion-induced acute and chronic kidney injury, Nephrol. Dial. Transplant. 26 (2011) 1474–1483. [11] Y.G. Zhu, X.M. Feng, J. Abbott, X.H. Fang, Q. Hao, A. Monsel, J.M. Qu, M.A. Matthay, J.W. Lee, Human mesenchymal stem cell microvesicles for treatment of Escherichia coli endotoxin-induced acute lung injury in mice, Stem Cells 32 (2014) 116–125. [12] R.C. Lai, F. Arslan, M.M. Lee, N.S. Sze, A. Choo, T.S. Chen, M. Salto-Tellez, L. Timmers, C.N. Lee, R.M. El Oakley, G. Pasterkamp, D.P. de Kleijn, S.K. Lim, Exosome secreted by MSC reduces myocardial ischemia/reperfusion injury, Stem Cell Res. 4 (2010) 214–222. [13] S.H. Ranganath, O. Levy, M.S. Inamdar, J.M. Karp, Harnessing the mesenchymal stem cell secretome for the treatment of cardiovascular disease, Cell Stem Cell 10 (2012) 244–258. [14] B. Zhang, M. Wang, A. Gong, X. Zhang, X. Wu, Y. Zhu, H. Shi, L. Wu, W. Zhu, H. Qian, W. Xu, HucMSC-exosome mediated-Wnt4 signaling is required for cutaneous wound healing, Stem Cells 33 (2014) 2158–2168.
[15] M.A. Lopez-Verrilli, F. Picou, F.A. Court, Schwann cell-derived exosomes enhance axonal regeneration in the peripheral nervous system, Glia 61 (2013) 1795–1806. [16] R. Nair, L. Santos, S. Awasthi, T. von Erlach, L.W. Chow, S. Bertazzo, M.M. Stevens, Extracellular vesicles derived from preosteoblasts influence embryonic stem cell differentiation, Stem Cells Dev. 23 (2014) 1625–1635. [17] N.J. Turner, S.F. Badylak, Regeneration of skeletal muscle, Cell Tissue Res. 347 (2012) 759–774. [18] B.K. Pedersen, M.A. Febbraio, Muscles, exercise and obesity: skeletal muscle as a secretory organ, Nat. Rev. Endocrinol. 8 (2012) 457–465. [19] J. Henningsen, K.T. Rigbolt, B. Blagoev, B.K. Pedersen, I. Kratchmarova, Dynamics of the skeletal muscle secretome during myoblast differentiation, Mol. Cell. Proteomics 9 (2010) 2482–2496. [20] D.P. Romancino, G. Paterniti, Y. Campos, A. De Luca, V. Di Felice, A. d'Azzo, A. Bongiovanni, Identification and characterization of the nano-sized vesicles released by muscle cells, FEBS Lett. 587 (2013) 1379–1384. [21] M. Guescini, D. Guidolin, L. Vallorani, L. Casadei, A.M. Gioacchini, P. Tibollo, M. Battistelli, E. Falcieri, L. Battistin, L.F. Agnati, V. Stocchi, C2C12 myoblasts release micro-vesicles containing mtDNA and proteins involved in signal transduction, Exp. Cell Res. 316 (2010) 1977–1984. [22] A. Forterre, A. Jalabert, K. Chikh, S. Pesenti, V. Euthine, A. Granjon, E. Errazuriz, E. Lefai, H. Vidal, S. Rome, Myotube-derived exosomal miRNAs downregulate Sirtuin1 in myoblasts during muscle cell differentiation, Cell Cycle 13 (2014) 78–89. [23] R.D. Madison, C. McGee, R. Rawson, G.A. Robinson, Extracellular vesicles from a muscle cell line (C2C12) enhance cell survival and neurite outgrowth of a motor neuron cell line (NSC-34), J. Extracell. Vesicles 3 (2014) 22865. [24] J. Lötvall, A.F. Hill, F. Hochberg, E.I. Buzás, D. Di Vizio, C. Gardiner, Y.S. Gho, I.V. Kurochkin, S. Mathivanan, P. Quesenberry, S. Sahoo, H. Tahara, M.H. Wauben, K.W. Witwer, C. Théry, Minimal experimental requirements for definition of extracellular vesicles and their functions: a position statement from the International Society for Extracellular Vesicles, J. Extracell. Vesicles 3 (2014) 26913. [25] J.S. Choi, B.S. Kim, J.D. Kim, Y.C. Choi, E.K. Lee, K. Park, H.Y. Lee, Y.W. Cho, In vitro expansion of human adipose-derived stem cells in a spinner culture system using human extracellular matrix powders, Cell Tissue Res. 345 (2011) 415–423. [26] J.S. Choi, H.J. Yang, B.S. Kim, J.D. Kim, J.Y. Kim, B. Yoo, K. Park, H.Y. Lee, Y.W. Cho, Human extracellular matrix (ECM) powders for injectable cell delivery and adipose tissue engineering, J. Control. Release 139 (2009) 2–7. [27] M. Shi, M. Ishikawa, N. Kamei, T. Nakasa, N. Adachi, M. Deie, T. Asahara, M. Ochi, Acceleration of skeletal muscle regeneration in a rat skeletal muscle injury model by local injection of human peripheral blood-derived CD133-positive cells, Stem Cells 27 (2009) 949–960. [28] M. Tabebordbar, E.T. Wang, A.J. Wagers, Skeletal muscle degenerative diseases and strategies for therapeutic muscle repair, Annu. Rev. Pathol. 8 (2013) 441–475. [29] D. Kuraitis, C. Giordano, M. Ruel, A. Musarò, E.J. Suuronen, Exploiting extracellular matrix-stem cell interactions: a review of natural materials for therapeutic muscle regeneration, Biomaterials 33 (2012) 428–443. [30] U.G. Longo, M. Loppini, A. Berton, F. Spiezia, N. Maffulli, V. Denaro, Tissue engineered strategies for skeletal muscle injury, Stem Cells Int. 2012 (2012) 175038. [31] D. Briggs, J.E. Morgan, Recent progress in satellite cell/myoblast engraftment — relevance for therapy, FEBS J. 280 (2013) 4281–4293. [32] R. Darabi, K. Gehlbach, R.M. Bachoo, S. Kamath, M. Osawa, K.E. Kamm, M. Kyba, R.C. Perlingeiro, Functional skeletal muscle regeneration from differentiating embryonic stem cells, Nat. Med. 14 (2008) 134–143. [33] J.H. Hwang, I.G. Kim, S. Piao, A.R. Jung, J.Y. Lee, K.D. Park, J.Y. Lee, Combination therapy of human adipose-derived stem cells and basic fibroblast growth factor hydrogel in muscle regeneration, Biomaterials 34 (2013) 6037–6045. [34] C. Borselli, H. Storrie, F. Benesch-Lee, D. Shvartsman, C. Cezar, J.W. Lichtman, H.H. Vandenburgh, D.J. Mooney, Functional muscle regeneration with combined delivery of angiogenesis and myogenesis factors, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 3287–3292. [35] G. Cossu, F. Mavilio, Myogenic stem cells for the therapy of primary myopathies: wishful thinking or therapeutic perspective? J. Clin. Invest. 105 (2000) 1669–1674. [36] S. Hartwig, S. Raschke, B. Knebel, M. Scheler, M. Irmler, W. Passlack, S. Muller, F.G. Hanisch, T. Franz, X. Li, H.D. Dicken, K. Eckardt, J. Beckers, M.H. de Angelis, C. Weigert, H.U. Häring, H. Al-Hasani, D.M. Ouwens, J. Eckel, J. Kotzka, S. Lehr, Secretome profiling of primary human skeletal muscle cells, Biochim. Biophys. Acta 1844 (2014) 1011–1017. [37] A. Forterre, A. Jalabert, E. Berger, M. Baudet, K. Chikh, E. Errazuriz, J. De Larichaudy, S. Chanon, M. Weiss-Gayet, A.M. Hesse, M. Record, A. Geloen, E. Lefai, H. Vidal, Y. Couté, S. Rome, Proteomic analysis of C2C12 myoblast and myotube exosome-like vesicles: a new paradigm for myoblast–myotube cross talk? PLoS One 9 (2014) e84153. [38] X.C. Chan, J.C. McDermott, K.W. Siu, Identification of secreted proteins during skeletal muscle development, J. Proteome Res. 6 (2007) 698–710. [39] M.C. Le Bihan, A. Bigot, S.S. Jensen, J.L. Dennis, A. Rogowska-Wrzesinska, J. Lainé, V. Gache, D. Furling, O.N. Jensen, T. Voit, V. Mouly, G.R. Coulton, G. Butler-Browne, In-depth analysis of the secretome identifies three major independent secretory pathways in differentiating human myoblasts, J. Proteome 77 (2012) 344–356. [40] J.A. Cho, H. Park, E.H. Lim, K.W. Lee, Exosomes from breast cancer cells can convert adipose tissue-derived mesenchymal stem cells into myofibroblast-like cells, Int. J. Oncol. 40 (2012) 130–138. [41] R. Chowdhury, J.P. Webber, M. Gurney, M.D. Mason, Z. Tabi, A. Clayton, Cancer exosomes trigger mesenchymal stem cell differentiation into pro-angiogenic and pro-invasive myofibroblasts, Oncotarget 6 (2015) 715–731. [42] J. von Maltzahn, N.C. Chang, C.F. Bentzinger, M.A. Rudnicki, Wnt signaling in myogenesis, Trends Cell Biol. 22 (2012) 602–609.
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115 [43] M.K. DO, T. Suzuki, B. Gerelt, Y. Sato, W. Mizunoya, M. Nakamura, Y. Ikeuchi, J.E. Anderson, R. Tatsumi, Time-coordinated prevalence of extracellular HGF, FGF2 and TGF-β3 in crush-injured skeletal muscle, Anim. Sci. J. 83 (2012) 712–717. [44] Y.H. Song, J.L. Song, P. Delafontaine, M.P. Godard, The therapeutic potential of IGF-I in skeletal muscle repair, Trends Endocrinol. Metab. 26 (2013) 310–319. [45] T. Braun, M. Gautel, Transcriptional mechanisms regulating skeletal muscle differentiation, growth and homeostasis, Nat. Rev. Mol. Cell Biol. 12 (2011) 349–361.
115
[46] M. Karalaki, S. Fili, A. Philippou, M. Koutsilieris, Muscle regeneration: cellular and molecular events, In Vivo 23 (2009) 779–796. [47] F. Relaix, P.S. Zammit, Stellite cells are essential for skeletal muscle regeneration: the cell on the edge returns centre stage, Development 139 (2012) 2845–2856. [48] Y.M. Ju, A. Atala, J.J. Yoo, S.J. Lee, In situ regeneration of skeletal muscle tissue through host cell recruitment, Acta Biomater. 10 (2014) 4332–4339.
Contents lists available at ScienceDirect
Journal of Controlled Release journal homepage: www.elsevier.com/locate/jconrel
Exosomes from differentiating human skeletal muscle cells trigger myogenesis of stem cells and provide biochemical cues for skeletal muscle regeneration Ji Suk Choi a, Hwa In Yoon a,b, Kyoung Soo Lee a, Young Chan Choi a, Seong Hyun Yang a, In-San Kim b,c, Yong Woo Cho a,⁎ a b c
Department of Chemical Engineering, Hanyang University, Ansan, Gyeonggi-do 426-791, Republic of Korea Center for Theragnosis, Biomedical Research Institute, Korea Institute of Science and Technology (KIST), Seoul 136-791, Republic of Korea KU-KIST School, Korea University, 1 Anam-dong, Seongbuk-gu, Seoul 136-701, Republic of Korea
a r t i c l e
i n f o
Article history: Received 13 March 2015 Received in revised form 3 December 2015 Accepted 12 December 2015 Available online 14 December 2015 Keywords: Exosomes Nanovesicles Stem cells Myogenesis Muscle regeneration
a b s t r a c t Exosomes released from skeletal muscle cells play important roles in myogenesis and muscle development via the transfer of specific signal molecules. In this study, we investigated whether exosomes secreted during myotube differentiation from human skeletal myoblasts (HSkM) could induce a cellular response from human adipose-derived stem cells (HASCs) and enhance muscle regeneration in a muscle laceration mouse model. The exosomes contained various signal molecules including myogenic growth factors related to muscle development, such as insulin-like growth factors (IGFs), hepatocyte growth factor (HGF), fibroblast growth factor-2 (FGF2), and platelet-derived growth factor-AA (PDGF-AA). Interestingly, exosome-treated HASCs fused with neighboring cells at early time points and exhibited a myotube-like phenotype with increased expression of myogenic proteins (myosin heavy chain and desmin). On day 21, mRNAs of terminal myogenic genes were also up-regulated in exosome-treated HASCs. Moreover, in vivo studies demonstrated that exosomes from differentiating HSkM reduced the fibrotic area and increased the number of regenerated myofibers in the injury site, resulting in significant improvement of skeletal muscle regeneration. Our findings suggest that exosomes act as a biochemical cue directing stem cell differentiation and provide a cell-free therapeutic approach for muscle regeneration. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Exosomes are small membrane vesicles (40–120 nm in diameter) secreted in a variety of mammalian cells following the fusion of multivesicular bodies (MVBs) with the plasma membrane during endosome maturation [1]. Exosomes derived from cells contain proteins and nucleic acids that reflect their original cell sources and transfer different cellular information to neighboring cells or even to distant tissues [2–4]. The interest of scientists in the role of exosomes has expanded rapidly because of their diverse pathological and therapeutic effects. Tumorderived exosomes play important roles in tumorigenesis, metastasis, angiogenesis, and generating tumor-associated cells by transporting growth factors/receptors or oncogenes. As exosomes can be key mediators of intercellular communication in tumors, their inhibition has become a focus of research in cancer therapy [5–8]. Several groups [9–14] have shown the therapeutic potential of exosomes derived ⁎ Corresponding author at: Department of Chemical Engineering, Hanyang University ERICA Campus, 55 Hanyangdaehak-ro, Sangnok-gu, Ansan, Gyeonggi-do 426-791, Republic of Korea. E-mail address: [email protected] (Y.W. Cho).
http://dx.doi.org/10.1016/j.jconrel.2015.12.018 0168-3659/© 2015 Elsevier B.V. All rights reserved.
from various cells with secretory capacity, although the therapeutic mechanisms of exosomes are still unclear. Stem cell-derived exosomes are a complex biological cargo that contains a range of stem cellderived secretomes, making them an ideal candidate for treatment of various disorders, such as liver disease [9], kidney injury [10], lung injury [11], myocardial infarction [12,13], and cutaneous wounds [14]. In addition to stem cell-derived exosomes, nerve cell-derived exosomes enhanced axonal regeneration in the peripheral nervous system [15] and preosteoblast-exosomes also increased the gene levels of neuroectoderm differentiation in embryonic stem cells [16]. Skeletal muscle is the largest organ in the human body, consisting of aligned multinucleate myofibers surrounded by basal lamina and satellite cells located beneath muscle fibers [17]. In addition to basic functions, such as energy metabolism, homeostasis and locomotion, skeletal muscle secretes hormone-like factors, termed ‘myokines’ that regulate angiogenesis and myogenesis within the muscles (autocrine/ paracrine) or communicate systemically with other tissues (endocrine) [18,19]. Skeletal myoblasts and myotubes secrete exosome-like nanovesicles, and these nanovesicles contain various myokines, proteins, microRNAs, and mitochondrial DNA, which could act as critical signals for skeletal muscle myogenesis, homeostasis, and development
108
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
[20–22]. Other studies also have shown that C2C12-derived exosomes stimulate survival and neurite outgrowth of motor neurons by paracrine signals [23]. Despite these findings on the secretory function of muscle cells, the biological roles of skeletal muscle cell-derived exosomes are not well understood yet. In this study, we hypothesized that exosomes secreted during the differentiation of human skeletal myoblasts (HSkM) into myotubes may contain specific biochemical cues that promote and regulate the differentiation of human adipose-derived stem cells (HASCs) with myogenic potential. Moreover, these exosomes could act as a therapeutic agent associated with muscle growth/repair, leading to efficient regeneration of damaged skeletal muscle. 2. Materials and methods 2.1. Human skeletal myoblast culture and differentiation HSkM (catalog #T4068, Lot# HC0512, sourced from post-gestational tissue, usually from the quadriceps or psoas tissue) was obtained from Applied Biological Materials Inc. (Richmond, BC, Canada). Growth medium and differentiation medium were purchased from PromoCell (Heidelberg, Germany). HSkM were maintained in growth medium supplemented with 5% fetal calf serum (FCS), 50 μg/mL petuin, 10 ng/mL epidermal growth factor, 1 ng/mL basic fibroblast growth factor, 10 μg/mL insulin, 0.4 μg/mL dexamethasone, and 1000 UI/mL penicillin/streptomycin (P/S) at 37 °C in humidified air containing 5% CO2. Differentiation was induced with serum-free medium containing 10 μg/mL insulin and 1000 UI/mL P/S for 5 days.
manufacturer's protocol. The exosome pellet was resuspended in phosphate buffered saline (PBS) and purified using exosome spin columns (MW 3000, Invitrogen). Exosomes isolated from proliferating HSkM were used as a control. The exosomal total protein content was quantified using the micro bicinchoninic acid (BCA) protein assay (Pierce Biotechnology, Rockford, IL, USA). Isolated exosomes were characterized with the minimal experimental requirements for extracellular vesicles (EVs) [24]. 2.3. Dynamic light scattering Dynamic light scattering (DLS) measurements were performed at 37 °C, using Zetasizer Nano ZS90 (Malvern, Worcestershire, UK). Exosomes resuspended in PBS were placed in a UV-transparent cuvette (Sarstedt AG & Co., Germany). The size distribution plot with an x-axis showing the distribution of estimated particle radius (nm) and a y-axis showing the relative intensity of the scattered light was analyzed with software. 2.4. Transmission electron microscopy Exosomes were fixed in 0.5% glutaraldehyde solution overnight. They were subsequently centrifuged at 13000 × g for 3 min, and then the supernatant was removed. The sample was dehydrated in absolute ethanol for 10 min and collected on formvar grids stabilized with carbon (TED PELLA, Inc., Redding, CA, USA). The grids were contrasted with 1% phosphotungstic acid for 1 min and then examined using transmission electron microscopy (TEM) with a JEM-2100F field emission electron microscope (JEOL Ltd., Japan).
2.2. Exosome isolation from differentiating HSkM 2.5. Western blotting Conditioned media (CM) were collected during differentiation of HSkM and used for exosome purification (Fig. 1A). Briefly, CM were collected from cells cultured in differentiation medium every day during 5 days. Collected CM were centrifuged (10 min, 2000 ×g) to eliminate cell debris. The resulting supernatant was concentrated through an Amicon® Ultra-15 3 K centrifugal filter (Millipore, Billerica, MA, USA) at 5000 ×g for 60 min. Exosomes were isolated using the Total Exosome Isolation kit from Invitrogen (Carlsbad, CA, USA) according to the
Differentiated HSkM and exosomes were lysed in RIPA buffer (25 mM Tris–HCl, pH 7.6, 150 mM NaCl, 0.5% Trixton X-100, 1% Na-deoxycholate, 0.1% sodium dodecyl sulfate and protease inhibitor cocktail). The protein in the supernatant was quantified using the BCA protein assay, electrophoresed on a 12–15% SDS-PAGE gel, and transferred to a PVDF membrane (Millipore, Billerica, MA). The membrane was blocked with 5% BSA in T-TBS (10 mM Tris, 150 mM NaCl, and
Fig. 1. Characterization of exosomes released from differentiating human skeletal myoblasts (HSkM). (A) Summary of the protocol for exosome purification from conditioned medium (CM) of differentiating human myoblasts and the induction of stem cell myogenesis using the exosomes. (B) Dynamic light scattering (DLS) showing the particle size distribution of exosomes. (C) Transmission electron microscope (TEM) analysis of exosomes. The scale bars represent 100 nm (black) and 200 nm (white). (D) Western blot analysis of exosomes. Equal amounts of total proteins (30 μg) extracted from exosomes and cells were immunoblotted for Alix, TSG 101, CD63, CD81 and β-actin.
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
0.1% Tween 20) for 1 h at room temperature, and subsequently incubated with primary antibodies overnight at 4 °C. After vigorous washing in T-TBS, the blots were incubated with horseradish peroxidase (HRP)tagged secondary antibodies for 1 h. The labeled proteins were visualized with the ChemiDoc™ XRS imaging system (Bio-Rad Laboratories, Inc., Hercules, CA, USA) using an enhanced chemiluminescence (ECL) kit (Pierce). Primary (Alix, TSG101, CD63, CD81, and β-actin) and secondary antibodies were purchased from Santa Cruz Biotechnology (Dallas, TX, USA). All chemical reagents and protease inhibitor cocktail (P-2714) were purchased from Sigma-Aldrich (St. Louis, MO, USA). 2.6. Human adipose-derived stem cell isolation Human adipose tissue was obtained with informed consent, as approved by the Institutional Review Board of the Catholic University of Korea, College of Medicine. HASCs were obtained from the adipose tissue of healthy human females as described previously [25,26]. 2.7. Cell proliferation assays The proliferation of HASCs in different conditions was analyzed using calcein AM staining and WST-1 assay. HASCs were seeded at 3 × 104 cells per well into 96-well plates in DMEM containing with 10% FBS and 1% P/S and allowed to adhere overnight at 37 °C under 5% CO2. The cells were washed with PBS and then maintained in GM (growth medium; DMEM, 10% FBS, 1% P/S), DM (differentiation medium; DMEM with 5% horse serum, 0.1 M dexamethasone, 50 μM hydrocortisone, and 1% P/S), GFM (growth factors containing medium; serum-free DMEM with 106 pg/mL hepatocyte growth factor; HGF, 65 pg/mL vascular endothelial growth factor; VEGF, 68 pg/mL insulin like growth factor; IGF, 100 pg/mL basic fibroblast growth factor; bFGF, and 63 pg/mL interleukin-6; IL-6) or EM (serum-free DMEM containing exosomes with 0, 10, 25, 50, 100, and 200 μg/mL) for 5 days. The concentrations of recombinant growth factors (Abcam, Cambridge, UK) in the 50 μg/mL exosomes were specified in the study. The cells were stained for 30 min with the calcein AM reagents (Sigma-Aldrich) on day 3 and observed using a fluorescence microscope (IX81; Olympus, Tokyo, Japan). Green fluorescence caused by the reaction of calcein with intracellular esterase indicated live cells. These experiments were performed in triplicate. The WST-1 reagent (Roche Applied Science, Mannheim, Germany) was added to each well, and the plate was incubated at 37 °C for 1 h under 5% CO2. After gentle pipetting, the 100 μL solution was transferred to a 96-well plate. The optical density was then measured at 440 nm using a microplate spectrophotometer (PowerWave XS, Bio-Tek Instruments, Winooski, VT, USA).
109
collected and prepared for immunofluorescent staining and quantitative PCR analysis. 2.10. Immunofluorescent staining Cells were washed twice with cold PBS, fixed with 4% paraformaldehyde/PBS for 1 h at 4 °C, and permeabilized with 0.3% Triton X-100 for 20 min at room temperature. Cells were incubated overnight at 4 °C with primary antibodies including desmin (Santa Cruz Biotechnology) and myosin heavy chain (MHC; Abcam, Cambridge, MA, USA), and then incubated with anti-rabbit IgG-FITC secondary antibody (Santa Cruz). Cells were counterstained with 1 μg/mL DAPI (4,6-diamidino-2phenylindole; Sigma-Aldrich) for 1 min and then observed with a fluorescence microscope (IX81, Olympus Corporation, Tokyo, Japan). The fusion index is defined as the ratio of the number of nuclei inside myotubes to the total number of nuclei at day 21. 2.11. Quantitative RT-PCR The human skeletal muscle-myogenesis & myopathy RT2 Profiler PCR Array (PAHS-099Z, SABiosciences, Valencia, CA, USA) was used to evaluate the expression of genes related to myogenesis. The total RNA was isolated from cells using the RNeasy Mini kit (Qiagen) according to the manufacturer's instructions. cDNA was synthesized from 500 ng of total RNA using the RT2 First Strand Kit (SABiosciences). 84 different genes were simultaneously amplified on a Stratagene Mx3000P (Agilent Technologies, Santa Clara, CA, USA). The thresholds and baselines were set according to the manufacturer's instructions. The fold change in gene expression (compared to controls) was calculated using SABiosciences webportal software (http://www.sabiosciences. com/pcr/arrayanalysis.php). p Values were calculated using the student's t-test and the experiments were repeated in triplicate. 2.12. Mouse model of skeletal muscle injury and exosome treatment
Differentiating HSkM exosomes were labeled with a membranelabeling dye (1′dioctadecyl-3,3,3′,3′-tetramethylindodicarbocyanine, DiD) in serum-free DMEM according to the manufacturer's protocol (Molecular Probes, Eugene, OR, USA). The labeled exosome suspension was filtered with PD MiniTrap G-10 columns (GE Healthcare Life Sciences, Pittsburgh, PA, USA) and the flow-through was used as the unbound dye control. HASCs were seeded at a density of 250 cells/mm2, incubated at 37 °C with labeled exosomes (50, 100, and 200 μg/mL) for 1 h, and then observed with a confocal microscope (Leica TCS SP8, Leica Microsystems, Buffalo Grove, IL, USA). The fluorescence intensities were quantified using an image analyzer.
All experiments with live animals were performed in compliance with the relevant laws and institutional guidelines of the Korea Institute of Science and Technology (KIST), and institutional committees approved the experiments. The muscle injury animal model was based on a previous report from Shi et al. [27]. Nude mice (male, 5 weeks old, n = 8) were anesthetized with ketamine (100 mg/kg) and xylazine (4–6 mg/kg), and an adequate depth of anesthesia was maintained such that the mice were unresponsive to tactile stimulation. For muscle injury, an anterolateral skin incision was made in both legs, and the tibialis anterior muscles were exposed. The muscles were then lacerated transversely at the mid-portion using a scalpel, creating a 5-mm wedgeshaped defect. After closure of the fascia with a suture, the exosome suspension (100 μg/100 μL in PBS) was injected into the right muscle defect and the same volume of PBS without exosome was injected into the left muscle defect as a control. For morphological evaluation of muscle regeneration after muscle laceration, muscle samples were fixed in 4% paraformaldehyde, embedded in paraffin, and sliced at 6 to 10 μm using a microtome. Sections were de-paraffinized, dehydrated, and stained with Masson's trichrome. The average area of collagen deposition and total number of myofibers with centralized nuclei within the muscle injured site were measured in three random fields (× 400) of four sections/muscle using Photoshop software (Adobe Systems Inc., San Jose, CA, USA).
2.9. Myogenic differentiation of human adipose-derived stem cells
2.13. Statistical analysis
Cells were plated at a density of 250 cells/mm2 in DMEM supplemented with 10% FBS and 1% P/S. After 72 h (~ 90% confluence), cells were rinsed with PBS and then maintained in different media. The media were changed every 3 days for up to 21 days. Cells were then
Experimental data were expressed as means ± standard deviation (SD). The student's two tailed t-test was performed with SPSS 17.0 statistical software (SPSS, Chicago, IL, USA), and statistical significance was accepted at p b 0.05 or p b 0.01.
2.8. Exosome labeling and cellular uptake
110
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
Fig. 2. The cell proliferation and cellular uptake of exosomes in human adipose-derived stem cells (HASCs). (A) Proliferation of HASC in GM (growth medium; DMEM, 10% FBS, 1% P/S), DM (differentiation medium; DMEM with 5% horse serum, 0.1 M dexamethasone, 50 μM hydrocortisone, and 1% P/S), GFM (growth factor containing medium; serum free DMEM with 106 pg/mL HGF, 65 pg/mL VEGF, 68 pg/mL IGF, 100 pg/mL bFGF, and 63 pg/mL IL-6) or EM (serum-free DMEM containing exosomes with 0, 10, 25, 50, 100, and 200 μg/mL) were visualized by calcein AM staining (A) and examined using a WST-1 assay kit (B) for 5 days. (C) Light differential interference contrast (DIC) and corresponding confocal images of HASCs after 1 h incubation with 50, 100, and 200 μg/mL of DiD-labeled exosomes, respectively. Images of DiD-labeled exosomes (red) with DAPI (blue) were visualized by merging the confocal images (Merge 1) or bright-field with confocal images (Merge 2). (D) Relative fluorescence intensity of DiD-labeled exosomes internalized in cells. Scale bars represent 200 μm (yellow) and 25 μm (white). Data are shown as mean ± standard deviation (n = 3) with significance at ⁎, ⁎⁎, ⁎⁎⁎p b 0.05. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
3. Results 3.1. Characterization of differentiating HSkM exosomes Exosomes isolated from differentiating human myoblasts were characterized in terms of size, morphology, particle yield, and surface markers (Figs. 1 and S1). Exosomes recovered with total exosome isolation reagents displayed a round or cup-like shape with a mean diameter of 113.7 nm (Fig. 1B and 1C) and had a particle yield of 1.7 × 109 particles/μg protein, comparable to that of exosomes isolated with the ultracentrifugation protocol (2.1 × 109 particles/μg protein) (Fig. S1). Western blotting for the exosomal markers revealed that the exosomes were positive for exosomal markers, including Alix, TSG101, CD63, CD81, FLOT-1, ICAM, EpCAM, and ANXA5, while non-exosomal marker, GM130 was not detected (Figs. 1D and S2). The protein
composition in exosomes was analyzed using LC–MS/MS. Proteins found in exosomes include extracellular matrix proteins, membraneassociated and cell adhesion proteins, cell structure and motility proteins, growth factors, and transmembrane signaling proteins (Table S1). Growth factors in exosomes were detected using antibody arrays. In total, 8 of 41 growth factors were significantly detected in exosomes when compared to control medium (Table S2 and Fig. S4). Interestingly, HGF, which is involved in the regulation of myogenic satellite cells, was highly expressed in exosomes. We also detected growth factors that act as potential regulators of development, function, and regeneration of skeletal muscle, including amphiregulin (AR), bFGF, IGF binding protein-3 (IGFBP-3), IGF-1, IL-6, neurotrophin-3 (NT-3), platelet-derived growth factor-AA (PDGF-AA), placenta growth factor (PLGF), transforming growth factor-beta1, 3 (TGF-β1, 3), VEGF, and VEGF receptor 3 (VEGF R3) (Table S3).
Fig. 3. Assessment of myogenesis of HASCs in differentiation medium (DM) and exosome-containing medium (EM) for 21 days. (A) HASCs were maintained in DM or EM containing 50 μg/mL of exosomes for 21 days, and were then immunostained with anti-myosin heavy chain (MHC) and anti-desmin. Cells were counterstained with DAPI (4,6-diamidino-2phenylindole), which stains nuclei blue. Scale bars represent 200 μm. (B) Immunofluorescence staining for desmin- and MHC-positive HSkM cells as a control. (C) Quantification of fusion index, representing the number of nuclei in multinucleated myotubes divided by the total number of nuclei in a field, with a myotube defined by at least three nuclei on day 21. (D) Length of multinucleated myotubes in DM or EM on day 21. (E) RT2 profiler PCR array for skeletal myogenesis of HASCs in DM or EM on day 21. The fold change of the test groups (DM and EM) compared to the control group (undifferentiated HASC) was calculated from three independent experiments using (http://www.sabiosciences.com/ pcrarraydata-analysis.php). Relative gene expression was normalized to housekeeping genes and expressed as the fold change comparing to undifferentiated HASC. ADIPOQ, FGF2, IGF1, IGF2, IL6, LEP, TGFB1, and TNF genes were evaluated as markers for skeletal autocrine signaling. MYF5, MYF6, MYOD1, MEF2C, MYOG, DMD, ACTA1, ACTN3, DAG1, DES, MYH1, MYH2, TNNT1, and TNNT3 genes were evaluated as markers for skeletal myogenesis. Data are shown as mean ± standard deviation (n = 3) with significance at ⁎, ⁎⁎p b 0.05. Abbreviations: ACTA; skeletal muscle actin alpha, ADIPOQ; adiponectin, DAG1; dystroglycan 1, DES; desmin, DMD; dystrophin, FGF2; fibroblast growth factor 2, IGF; insulin-like growth factor, LEP; leptin, IL6; interleukin 6, MEF2C; myocyte enhancer factor 2C, MYOD1; myogenic differentiation 1, MYOG; myogenin, MYF; myogenic factor, MYH; myosin heavy chain, TGFB1; transforming growth factor beta 1, TNF; tumor necrosis factor, TNNT; troponin T.
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
3.2. Cell proliferation and cellular uptake The effect of the exosome dose on cell proliferation was investigated using calcein AM staining and WST-1 assay for 5 days (Fig. 2A and 2B).
111
The proliferation of HASCs treated with exosomes was compared to the initial seeding absorbance (440 nm). On day 5, HASCs cultured in either DM, GFM or EM showed a slightly lower cell proliferation capacity than that in GM. However, exosomes significantly affect cell proliferation in a
112
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
dose dependent manner. In particular, 50 μg/mL exosome-treated cells proliferated better than cells maintained in GFM or exosome-free medium. To study the cellular uptake of differentiating HSkM exosomes at early time points, HASCs were incubated with 50 to 200 μg/mL of fluorescent dye-labeled exosomes (red) for 1 h (Fig. 2C and 2D). Exosomes were internalized into the cytoplasm of HASCs within 1 h and were partly co-localized within the nuclei. These results imply that differentiating HSkM exosomes are successfully delivered into stem cells with high cellular uptake efficiency.
the regenerated myofibers continued to grow and mature, resulting in reconstruction of muscle architecture. Damaged fibers, and inflammatory cells in the exosome (−) group were still observed with a few regenerating myofibers at day 14. The results demonstrated that the exosome-treated group had a large number of regenerative myofibers with minimal fibrosis compared with the exosome (−) groups, indicating that muscle regeneration was significantly improved by differentiating HSkM exosome treatment.
3.3. Myogenesis of HASCs promoted by differentiating HSkM exosomes
Skeletal muscle has the capacity to regenerate following common sports-related injuries. However, severe injuries can result in extensive fibrosis, scarring, and loss of muscle function [28]. Various approaches using the cellular and biochemical cues associated with muscle regeneration have been explored in vitro and in vivo to enhance the regeneration capacity of injured skeletal muscle [29,30]. Therapeutic approaches using myoblasts, muscle satellite cells, stem cells of various origins, and stem cells combined with growth factors have been used to improve muscle function and regeneration [31–34]. Although cell-based therapies have clear beneficial effects on skeletal muscle repair, there are still a number of concerns, such as limited survival and the reduced regenerative capacity of engrafted cells, as well as immune-mediated rejection [35]. Exosomes secreted by various cells may be an effective cell-free therapeutic agent because of their diverse biological and physiological functions as well as their key role as vehicles of intercellular communication [1,2]. Skeletal muscle cells also secrete a large number of myokines and some nano-vesicles that influence the growth, function, and development of muscle tissue [20]. Several groups have reported that exosomes secreted from skeletal muscle cells contain more than 400 different signal molecules, including microRNAs and proteins, that are involved in many cellular processes, including myogenesis by transporting and delivering a cargo of specific signals [22,36–38]. In spite of the proteomic and genomic characterization of exosomes, their biological functions remain unclear. As previously reported [20,37, 39], exosomes isolated from differentiating HSkM showed a typical round- or cup-shaped morphology ranging in diameter from 70– 120 nm and expressed specific cell surface molecules that are involved in their biogenesis, such as Alix, TSG101, CD63, CD81, FLOT-1, ICAM, EpCAM, and ANXA5 (Figs. 1, S1, and S2). Although we did not carry out in-depth proteomic and growth factor analyses on exosome contents, differentiating HSkM exosomes also contained various proteins, such as extracellular matrix (ECM) protein precursors, cell surface proteins, cell surface receptors, transmembrane signaling proteins, and myogenic growth factors, such as the HB-EGF family (AR and HB-EGF), VEGF family (VEGF-R3 and PLGF), IGFBP-3, HGF, IGF-I/II, bFGF, IL-6, NT-3, and PDGF-AA. Interestingly, HGF, which is known to be important in the activation and differentiation of muscle satellite cells, was appreciably expressed in exosomes (Tables S1, S2, S3, and Fig. S4). We hypothesized that differentiating HSkMs secrete exosomes loaded with myogenic factors and these exosomes might regulate in vitro myogenesis of HASCs. The physiological role of exosomes and their contents in regulating stem cell differentiation are not well defined; however, several groups [14,40,41] have addressed that exosomal factors derived from various cells contribute to proliferation, migration, and differentiation of cells. For example, exosomal Wnt 4 protein secreted from human mesenchymal stem cell (MSC) promotes proliferation and migration of keratinocytes and dermal fibroblasts through activation of β-catenin signaling in cells [14]. In addition, TGF-β1 contained in cancer cell exosomes induce the differentiation of ASCs and MSCs into cancer associated myofibroblast via TGF-β1/SMAD-mediated signaling pathway [40,41]. We found that exosomes significantly contribute to myogenic differentiation of HASCs (Figs. 3, S6, and S7). Stimulation by exosomes rapidly altered the morphological phenotype
To investigate the cellular function of differentiating HSkM exosomes on HASCs, HASCs were incubated in DM, GFM, and EM for 21 days and then stained for desmin and MHC, myogenesis specific markers (Fig. 3). After 7 days of culture, the HASCs treated with 50 μg/mL of exosome started to elongate and a small proportion of desmin-staining cells were present. The HASCs maintained in GFM, EM 10 or EM 25 started to change their morphology and a small proportion of MHC-staining cells were present after 14 days. However, the differentiation efficiency of HASCs in GFM, EM 10 or EM 25 was lower than that of the HASCs in EM 50 (Fig. S7). After 21 days, elongated multinucleated HASCs expressed desmin and MHC in both DM and EM 50, and the intensities of the staining significantly increased (Fig. 3A). However, high concentrations (200 μg/mL) of exosomes resulted in high cell death (Figs. 2A, 2B, and S6). The fusion index increased from 14.34% in DM to 23.35% in EM, and the myotube length also increased from 175.46 μm in DM to 204.03 μm in EM (Fig. 3C and 3D). Although the conversion efficiency of HASCs into myotubes in EM was lower than that of the HSkMs used as a positive control (fusion index, 56.67% and myotube length, 536.92 μm for 5 days), these results indicate that differentiating HSkM exosomes provide biochemical cues to stimulate myogenesis of HASCs, and their functional effects were more significant than that of conventional differentiation medium. We assessed the expression of 49 genes associated with skeletal myogenesis of HASCs treated with differentiating HSkM exosomes using qPCR arrays. Table S4 and Fig. 3E show the list and fold change of up- and down-regulated genes in HASCs grown in DM or EM for 21 days, compared to undifferentiated HASCs (p value b 0.05). The upregulated genes in EM included skeletal myogenesis-related genes (ACTA1 and MYOD1), muscle autocrine signaling-related genes (FGF2 and TNF), and muscle contractility-related genes (DAG1, DES, MYH1/2, TNNC1, and TNNT3). There were distinct differences in gene expression between HASCs differentiated in DM and EM on day 21 (Fig. 3E). Under DM conditions, the autocrine signaling genes ADIPOQ, IL6, and LEP, and the early myogenic markers MYF5 and MEF2C were significantly upregulated, compared with EM conditions. In contrast, HASCs differentiated in EM highly expressed the terminal differentiation markers MYOD1, ACTA1, DAG1, DES, MYH1, MYH2, and TNNT1. Based on these results it appears that the differentiating HSkM exosomes can quickly deliver signaling molecules that regulate the skeletal myogenic fate of HASCs, resulting in the acceleration of myogenic differentiation of HASCs. 3.4. Muscle regeneration enhanced by differentiating HSkM exosomes To investigate whether differentiating HSkM exosomes could contribute to muscle regeneration in vivo, we injected exosomes at the lacerated muscle sites. Masson trichrome staining was used to evaluate fibrosis and muscle regeneration after exosome injection (Fig. 4). At days 7 and 14, the collagen deposition in the exosome (+) groups (42.36 ± 7.23% and 15.36 ± 6.95%) was significantly smaller compared with the exosome (−) groups (60.21 ± 10.37% and 31.31 ± 6.9%). In the exosome (+) groups, there were more regenerating myofibers with centralized nuclei than in the exosome (−) groups at day 7, and
4. Discussion
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
113
Fig. 4. Histological images of skeletal muscle regeneration after muscle cell-derived exosome injection. Histological evaluation was carried out by Masson trichrome staining. (A) Normal hindlimb muscles of mice were used as a control. Seven days post-injury, newly regenerated myofibers with centralized nuclei were observed in the exosome-treated skeletal muscle, although fibrosis between regenerated myofibers was also visible. At day 14, the exosome-treated muscle showed enhanced muscle regeneration, whereas untreated skeletal muscle remained abnormal (myofibers, red-orange color; collagenous scar tissue, blue color). Scale bars represent 1 mm (red), 200 μm (black) and 100 μm (white). (B) Quantification of collagen deposition and the number of centronuclear myofibers within the muscle injured site at days 7 and 14 (⁎, ⁎⁎p b 0.05).
of HASCs at early time points and resulted in significant expression of myogenic markers (desmin and MHC) with a fusion index of 20%, when compared with HASCs in differentiation medium. Moreover, mRNA expression of myogenic genes, such as FGF2, TNF, MYOD1, DAG1, DES, MYH1/2 and TNNT1, which are related to skeletal autocrine signaling and terminal myogenic differentiation, was increased in HASCs stimulated by exosomes over 21 days (Fig. 3E). The process of myogenesis is orchestrated by complex signaling cascades initiated by extrinsic factors, including cytokines, growth factors, and signaling molecules [42]. In particular, several paracrine growth factors, including IGFI/II, FGF2, HGF, PDGF, and TGF-β play major roles in the regulation of muscle satellite cell chemotaxis, proliferation, and differentiation through the stimulation of Pax genes and myogenic regulatory factors (MRFs; MyoD, MYF5/6, myogenin) [43–45]. From a paracrine effect perspective, it seems that differentiating HSkM exosomes containing diverse myogenic factors are efficiently delivered into stem cells, and this cargo synergistically triggers myogenic signaling pathways in recipient cells, resulting in skeletal myogenic differentiation. We also investigated the therapeutic effect of differentiating HSkM exosomes using a muscle injury model. Skeletal muscle regeneration is a complex process that is orchestrated by a collaboration of many regulatory factors. After muscle injury, some cytokines and growth factors are released from inflammatory and stromal cells. These autocrine/
paracrine factors could not only create a regenerative niche for the migration, activation, and differentiation of myoblasts and quiescent satellite cells, which is critical for muscle regeneration, but may also balance regeneration and fibrosis [46,47]. We found that differentiating HSkM exosomes accelerate skeletal muscle regeneration in a laceration mouse model by reducing the collagen deposition in injured muscle and increasing the number of regenerated myofibers after exosome injection (Fig. 4). As mentioned above, differentiating HSkM exosomes are rich in various myogenic factors such as HGF. HGF is a potent chemoattractant for muscle stem cell activation and migration to injury sites [17]. Although HGF has a dose-dependent effect on skeletal muscle repair, it has synergistic effects with other myogenic growth factors, such as FGF2, IGF1, and TGF-β, which in combination significantly increase muscle repair and regeneration compared with the individual growth factors alone [43,48]. Therefore, these results support our hypothesis that the myogenic factors contained in exosomes could effectively provide a niche favorable to the skeletal muscle regeneration by stimulating the differentiation of myoblasts and stem cells (Fig. S5). However, it remains unclear as to which factors in exosomes play a key role for controlling cell fate and promoting skeletal muscle regeneration. Moreover, we should consider the adverse effects that can arise from high doses of exosomes (e.g., cell apoptosis) because complex molecules contained in exosomes can stimulate different signaling
114
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115
pathways. Nonetheless, we expect that exosomes can be an alternative approach to stem cell-based therapy and show promise as a therapeutic agent for skeletal muscle regeneration. 5. Conclusion Exosomes derived from differentiating HSkM can induce in vitro myogenesis of HASCs and improve in vivo muscle regeneration after skeletal muscle injury. Although the mechanism for the biological effects of exosomes remains unclear, exosomes may regulate skeletal myogenesis through the transfer of diverse myogenic factors, thereby controlling stem cell fate toward myogenesis and improving muscle regeneration. Therefore, we propose that exosomes can provide a cell-free therapeutic approach for regenerative medicine and for skeletal muscle regeneration in particular. Acknowledgments This work was supported by the Basic Research Program (Grant No. 2012-008294) and the Bio & Medical Technology Development Program (Grant No. 2011-0019774) through the National Research Foundation of Korea (NRF) funded by the Korean government (MEST). This study was also supported by a grant from the National R&D Program for Cancer Control, Ministry of Health and Welfare, Republic of Korea (1420390). Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.jconrel.2015.12.018. References [1] C. Théry, L. Zitvogel, S. Amigorena, Exosomes: composition, biogenesis and function, Nat. Rev. Immunol. 2 (2002) 569–579. [2] G. Raposo, W. Stoorvogel, Extracellular vesicles: exosomes, microvesicles, and friends, J. Cell Biol. 200 (2013) 373–383. [3] R. van der Meel, M.H. Fens, P. Vader, W.W. van Solinge, O. Eniola-Adefeso, R.M. Schiffelers, Extracellular vesicles as drug delivery systems: lessons from the liposome field, J. Control. Release 195 (2014) 72–85. [4] S.M. van Dommelen, P. Vader, S. Lakhal, S.A. Kooijmans, W.W. van Solinge, M.J. Wood, R.M. Schiffelers, Microvesicles and exosomes: opportunities for cell-derived membrane vesicles in drug delivery, J. Control. Release 161 (2012) 635–644. [5] J. Webber, R. Steadman, M.D. Mason, Z. Tabi, A. Clayton, Cancer exosomes trigger fibroblast to myofibroblast differentiation, Cancer Res. 70 (2010) 9621–9630. [6] L. Pascucci, V. Coccè, A. Bonomi, D. Ami, P. Ceccarelli, E. Ciusani, L. Viganò, A. Locatelli, F. Sisto, S.M. Doglia, E. Parati, M.E. Bernardo, M. Muraca, G. Alessandri, G. Bondiolotti, A. Pessina, Paclitaxel is incorporated by mesenchymal stromal cells and released in exosomes that inhibit in vitro tumor growth: a new approach for drug delivery, J. Control. Release 192 (2014) 262–270. [7] T. Smyth, M. Kullberg, N. Malik, P. Smith-Jones, M.W. Graner, T.J. Anchordoquy, Biodistribution and delivery efficiency of unmodified tumor-derived exosomes, J. Control. Release 199 (2015) 145–155. [8] P. Vader, X.O. Breakefield, M.J. Wood, Extracellular vesicles: emerging targets for cancer therapy, Trends Mol. Med. 20 (2014) 385–393. [9] S. Lemoinne, D. Thabut, C. Housset, R. Moreau, D. Valla, C.M. Boulanger, P.E. Rautou, The emerging roles of microvesicles in liver diseases, Nat. Rev. Gastroenterol. Hepatol. 11 (2014) 350–361. [10] S. Gatti, S. Bruno, M.C. Deregibus, A. Sordi, V. Cantaluppi, C. Tetta, G. Camussi, Microvesicles derived from human adult mesenchymal stem cells protect against ischaemia-reperfusion-induced acute and chronic kidney injury, Nephrol. Dial. Transplant. 26 (2011) 1474–1483. [11] Y.G. Zhu, X.M. Feng, J. Abbott, X.H. Fang, Q. Hao, A. Monsel, J.M. Qu, M.A. Matthay, J.W. Lee, Human mesenchymal stem cell microvesicles for treatment of Escherichia coli endotoxin-induced acute lung injury in mice, Stem Cells 32 (2014) 116–125. [12] R.C. Lai, F. Arslan, M.M. Lee, N.S. Sze, A. Choo, T.S. Chen, M. Salto-Tellez, L. Timmers, C.N. Lee, R.M. El Oakley, G. Pasterkamp, D.P. de Kleijn, S.K. Lim, Exosome secreted by MSC reduces myocardial ischemia/reperfusion injury, Stem Cell Res. 4 (2010) 214–222. [13] S.H. Ranganath, O. Levy, M.S. Inamdar, J.M. Karp, Harnessing the mesenchymal stem cell secretome for the treatment of cardiovascular disease, Cell Stem Cell 10 (2012) 244–258. [14] B. Zhang, M. Wang, A. Gong, X. Zhang, X. Wu, Y. Zhu, H. Shi, L. Wu, W. Zhu, H. Qian, W. Xu, HucMSC-exosome mediated-Wnt4 signaling is required for cutaneous wound healing, Stem Cells 33 (2014) 2158–2168.
[15] M.A. Lopez-Verrilli, F. Picou, F.A. Court, Schwann cell-derived exosomes enhance axonal regeneration in the peripheral nervous system, Glia 61 (2013) 1795–1806. [16] R. Nair, L. Santos, S. Awasthi, T. von Erlach, L.W. Chow, S. Bertazzo, M.M. Stevens, Extracellular vesicles derived from preosteoblasts influence embryonic stem cell differentiation, Stem Cells Dev. 23 (2014) 1625–1635. [17] N.J. Turner, S.F. Badylak, Regeneration of skeletal muscle, Cell Tissue Res. 347 (2012) 759–774. [18] B.K. Pedersen, M.A. Febbraio, Muscles, exercise and obesity: skeletal muscle as a secretory organ, Nat. Rev. Endocrinol. 8 (2012) 457–465. [19] J. Henningsen, K.T. Rigbolt, B. Blagoev, B.K. Pedersen, I. Kratchmarova, Dynamics of the skeletal muscle secretome during myoblast differentiation, Mol. Cell. Proteomics 9 (2010) 2482–2496. [20] D.P. Romancino, G. Paterniti, Y. Campos, A. De Luca, V. Di Felice, A. d'Azzo, A. Bongiovanni, Identification and characterization of the nano-sized vesicles released by muscle cells, FEBS Lett. 587 (2013) 1379–1384. [21] M. Guescini, D. Guidolin, L. Vallorani, L. Casadei, A.M. Gioacchini, P. Tibollo, M. Battistelli, E. Falcieri, L. Battistin, L.F. Agnati, V. Stocchi, C2C12 myoblasts release micro-vesicles containing mtDNA and proteins involved in signal transduction, Exp. Cell Res. 316 (2010) 1977–1984. [22] A. Forterre, A. Jalabert, K. Chikh, S. Pesenti, V. Euthine, A. Granjon, E. Errazuriz, E. Lefai, H. Vidal, S. Rome, Myotube-derived exosomal miRNAs downregulate Sirtuin1 in myoblasts during muscle cell differentiation, Cell Cycle 13 (2014) 78–89. [23] R.D. Madison, C. McGee, R. Rawson, G.A. Robinson, Extracellular vesicles from a muscle cell line (C2C12) enhance cell survival and neurite outgrowth of a motor neuron cell line (NSC-34), J. Extracell. Vesicles 3 (2014) 22865. [24] J. Lötvall, A.F. Hill, F. Hochberg, E.I. Buzás, D. Di Vizio, C. Gardiner, Y.S. Gho, I.V. Kurochkin, S. Mathivanan, P. Quesenberry, S. Sahoo, H. Tahara, M.H. Wauben, K.W. Witwer, C. Théry, Minimal experimental requirements for definition of extracellular vesicles and their functions: a position statement from the International Society for Extracellular Vesicles, J. Extracell. Vesicles 3 (2014) 26913. [25] J.S. Choi, B.S. Kim, J.D. Kim, Y.C. Choi, E.K. Lee, K. Park, H.Y. Lee, Y.W. Cho, In vitro expansion of human adipose-derived stem cells in a spinner culture system using human extracellular matrix powders, Cell Tissue Res. 345 (2011) 415–423. [26] J.S. Choi, H.J. Yang, B.S. Kim, J.D. Kim, J.Y. Kim, B. Yoo, K. Park, H.Y. Lee, Y.W. Cho, Human extracellular matrix (ECM) powders for injectable cell delivery and adipose tissue engineering, J. Control. Release 139 (2009) 2–7. [27] M. Shi, M. Ishikawa, N. Kamei, T. Nakasa, N. Adachi, M. Deie, T. Asahara, M. Ochi, Acceleration of skeletal muscle regeneration in a rat skeletal muscle injury model by local injection of human peripheral blood-derived CD133-positive cells, Stem Cells 27 (2009) 949–960. [28] M. Tabebordbar, E.T. Wang, A.J. Wagers, Skeletal muscle degenerative diseases and strategies for therapeutic muscle repair, Annu. Rev. Pathol. 8 (2013) 441–475. [29] D. Kuraitis, C. Giordano, M. Ruel, A. Musarò, E.J. Suuronen, Exploiting extracellular matrix-stem cell interactions: a review of natural materials for therapeutic muscle regeneration, Biomaterials 33 (2012) 428–443. [30] U.G. Longo, M. Loppini, A. Berton, F. Spiezia, N. Maffulli, V. Denaro, Tissue engineered strategies for skeletal muscle injury, Stem Cells Int. 2012 (2012) 175038. [31] D. Briggs, J.E. Morgan, Recent progress in satellite cell/myoblast engraftment — relevance for therapy, FEBS J. 280 (2013) 4281–4293. [32] R. Darabi, K. Gehlbach, R.M. Bachoo, S. Kamath, M. Osawa, K.E. Kamm, M. Kyba, R.C. Perlingeiro, Functional skeletal muscle regeneration from differentiating embryonic stem cells, Nat. Med. 14 (2008) 134–143. [33] J.H. Hwang, I.G. Kim, S. Piao, A.R. Jung, J.Y. Lee, K.D. Park, J.Y. Lee, Combination therapy of human adipose-derived stem cells and basic fibroblast growth factor hydrogel in muscle regeneration, Biomaterials 34 (2013) 6037–6045. [34] C. Borselli, H. Storrie, F. Benesch-Lee, D. Shvartsman, C. Cezar, J.W. Lichtman, H.H. Vandenburgh, D.J. Mooney, Functional muscle regeneration with combined delivery of angiogenesis and myogenesis factors, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 3287–3292. [35] G. Cossu, F. Mavilio, Myogenic stem cells for the therapy of primary myopathies: wishful thinking or therapeutic perspective? J. Clin. Invest. 105 (2000) 1669–1674. [36] S. Hartwig, S. Raschke, B. Knebel, M. Scheler, M. Irmler, W. Passlack, S. Muller, F.G. Hanisch, T. Franz, X. Li, H.D. Dicken, K. Eckardt, J. Beckers, M.H. de Angelis, C. Weigert, H.U. Häring, H. Al-Hasani, D.M. Ouwens, J. Eckel, J. Kotzka, S. Lehr, Secretome profiling of primary human skeletal muscle cells, Biochim. Biophys. Acta 1844 (2014) 1011–1017. [37] A. Forterre, A. Jalabert, E. Berger, M. Baudet, K. Chikh, E. Errazuriz, J. De Larichaudy, S. Chanon, M. Weiss-Gayet, A.M. Hesse, M. Record, A. Geloen, E. Lefai, H. Vidal, Y. Couté, S. Rome, Proteomic analysis of C2C12 myoblast and myotube exosome-like vesicles: a new paradigm for myoblast–myotube cross talk? PLoS One 9 (2014) e84153. [38] X.C. Chan, J.C. McDermott, K.W. Siu, Identification of secreted proteins during skeletal muscle development, J. Proteome Res. 6 (2007) 698–710. [39] M.C. Le Bihan, A. Bigot, S.S. Jensen, J.L. Dennis, A. Rogowska-Wrzesinska, J. Lainé, V. Gache, D. Furling, O.N. Jensen, T. Voit, V. Mouly, G.R. Coulton, G. Butler-Browne, In-depth analysis of the secretome identifies three major independent secretory pathways in differentiating human myoblasts, J. Proteome 77 (2012) 344–356. [40] J.A. Cho, H. Park, E.H. Lim, K.W. Lee, Exosomes from breast cancer cells can convert adipose tissue-derived mesenchymal stem cells into myofibroblast-like cells, Int. J. Oncol. 40 (2012) 130–138. [41] R. Chowdhury, J.P. Webber, M. Gurney, M.D. Mason, Z. Tabi, A. Clayton, Cancer exosomes trigger mesenchymal stem cell differentiation into pro-angiogenic and pro-invasive myofibroblasts, Oncotarget 6 (2015) 715–731. [42] J. von Maltzahn, N.C. Chang, C.F. Bentzinger, M.A. Rudnicki, Wnt signaling in myogenesis, Trends Cell Biol. 22 (2012) 602–609.
J.S. Choi et al. / Journal of Controlled Release 222 (2016) 107–115 [43] M.K. DO, T. Suzuki, B. Gerelt, Y. Sato, W. Mizunoya, M. Nakamura, Y. Ikeuchi, J.E. Anderson, R. Tatsumi, Time-coordinated prevalence of extracellular HGF, FGF2 and TGF-β3 in crush-injured skeletal muscle, Anim. Sci. J. 83 (2012) 712–717. [44] Y.H. Song, J.L. Song, P. Delafontaine, M.P. Godard, The therapeutic potential of IGF-I in skeletal muscle repair, Trends Endocrinol. Metab. 26 (2013) 310–319. [45] T. Braun, M. Gautel, Transcriptional mechanisms regulating skeletal muscle differentiation, growth and homeostasis, Nat. Rev. Mol. Cell Biol. 12 (2011) 349–361.
115
[46] M. Karalaki, S. Fili, A. Philippou, M. Koutsilieris, Muscle regeneration: cellular and molecular events, In Vivo 23 (2009) 779–796. [47] F. Relaix, P.S. Zammit, Stellite cells are essential for skeletal muscle regeneration: the cell on the edge returns centre stage, Development 139 (2012) 2845–2856. [48] Y.M. Ju, A. Atala, J.J. Yoo, S.J. Lee, In situ regeneration of skeletal muscle tissue through host cell recruitment, Acta Biomater. 10 (2014) 4332–4339.